Peptidases and Neuropeptide Processing (Methods in Neurosciences)

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Methods in Neurosciences Volume 23

Peptidases and Neuropeptide Processing

Methods in Neurosciences Editor-in-Chief

P. Michael Conn

Methods in Neurosciences Volume 23

Peptidases and Neuropeptide Processing

Edited by A. Ian Smith Peptide Biology Laboratory Baker Medical Research Institute Prahran, Victoria Australia

ACADEMIC PRESS San Diego New York

Boston

London

Sydney Tokyo Toronto

This book is printed on acid-free paper.

Copyright 9 1995 by ACADEMIC PRESS, INC. 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. Academic Press, Inc. A Division of Harcourt Brace & Company 525 B Street, Suite 1900, San Diego, California 92101-4495

United Kingdom Edition published by Academic Press Limited 24-28 Oval Road, London NW1 7DX

International Standard Serial Number: 1043-9471 International Standard Book Number: 0-12-185293-8

PRINTED IN THE UNrIED STATES OF AMERICA 95 96 97 98 99 00 EB 9 8 7 6

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4

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Table of Contents

Contributors to Volume 23 Preface Volumes in Series

ix ~ 1 7 6 1 7 6

Xlll XV

Section I Molecular Approaches for the Study of Intracellular Processing Enzymes 1. Molecular Strategies for Identifying Processing Enzymes Nabil G. Seidah 2. In Situ Hybridization Techniques to Map Processing

Enzymes 16

Martin K.-H. Schiifer and Robert Day

3. Analysis of Ontogeny of Processing Enzyme Gene Expression and Regulation 45

Min Zheng and John E. Pintar

4. Use of Vaccinia Virus Vectors to Study Neuropeptide Processing 65

Judy K. VanSlyke, Laurel Thomas, and Gary Thomas

5. Overexpression of Neuropeptide Precursors and Processing Enzymes 94

Iris Lindberg and Yi Zhou

6. Use of Antisense RNA to Block Peptide-Processing Enzyme Expression Richard E. Mains

109

Section II Immunological and Biochemical Approaches to the Study of Peptide-Processing Pathways 7. Combination of High-Performance Liquid Chromatography and Radioimmunoassay for Characterization of Peptide-Processing Pathways A. Ian Smith and Rebecca A. Lew

125

8. Development and Use of Two-Site Immunometric Assays for Examining Peptide-Processing Pathways Steven R. Crosby

140

vi

TABLE OF CONTENTS 9. Methods for Identification of Neuropeptide-Processing Pathways Paul Cohen, Mohamed Rholam, and Hamadi Boussetta

155

10. Immunological and Related Techniques for Studying Neurohypophyseal Peptide-Processing Pathways Harold Gainer, Mark O. Lively, and Mariana Morris

195

11. Approaches to Assessing Ontogeny of Processing Enzymes Richard G. Allen and Julianne Stack

208

12. Measurement, Distribution, and Subcellular Localization of Peptide-Amidating Activity Rebecca A. Lew and A. Ian Smith

219

13. Methods for Studying Carboxypeptidase E Lloyd D. Fricker

237

14. Characterization of Endothelin-Converting Enzymes Terry J. Opgenorth, Sadao Kimura, and Jinshyun R. Wu-Wong

251

15. In Vivo Approaches for Studying Peptide Processing Arthur Shulkes

266

Section III Identification and Characterization of Extracellular Processing Enzymes in the Central Nervous System 16. Identification and Characterization of Central Nervous System Peptidase Activities John R. McDermont and Alison M. Gibson

281

17. Strategies for Characterizing, Cloning, and Expressing Soluble Endopeptidases Marc J. Glucksman and James L. Roberts

296

18. Proteolytic Processing and Amyloid Protein Precursor of Alzheimer's Disease D. H. Small, G. Reed, S. J. Fuller, A. Weidemann, K. Beyreuther, and C. L. Masters

317

19. Strategies for Measurement of Angiotensin and Bradykinin Peptides and Their Metabolites in Central Nervous System and Other Tissues Duncan J. Campbell, Anne C. Lawrence, Athena Kladis, and Ann-Maree Duncan

328

20. Distribution and Roles of Endopeptidase 24.11 Anthony J. Turner and Kay Barnes

344

TABLE OF CONTENTS

vii

21. Identification and Distribution of Endopeptidase 24.16 in the Central Nervous System F. Checler, P. Dauch, H. Barelli, V. Dive, Y. Masuo, B. Vincent, and J. P. Vincent

363

22. Autoradiographic Techniques to Map Angiotensin-Converting Enzyme in Brain and Other Tissues Siew Yeen Chai and Frederick A. O. Mendelsohn

Index

383 399

This Page Intentionally Left Blank

Contributors to Volume 23

Article numbers are in parentheses following the names of contributors. Affiliations listed are current.

RICHARD G. ALLEN (11), Center of Research on Occupational and Environmental Toxicology, Oregon Health Sciences University, Portland, Oregon 97201 H. BARELLI (21), Institut de Pharmacologie Mol6culaire et Cellulaire, Centre National de la Recherche Scientifique, 06560 Valbonne, France KaY BARNES (20), Department of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT, United Kingdom K. BEYREtJTHEI~ (18), Center for Molecular Biology, University of Heidelberg, D-6900 Heidelberg, Germany HaMaoi BOUSSETTA (9), Biochimie des Signaux R6gulateurs Cellulaires et Mol6culaires, Universit6 Pierre et Marie Curie, F-75006 Paris, France DUNCAN J. CAMVBEU~ (19), St. Vincent's Institute of Medical Research, Fitzroy, Victoria 3065, Australia SIEW YEEN CI-IAI (22), Department of Medicine, University of Melbourne, Austin Hospital, Melbourne, Victoria 3048, Australia F. CI-IECLErt(21), Institut de Pharmacologie Mol6culaire et Cellulaire, Centre National de la Recherche Scientifique, 06560 Valbonne, France PAUL COHEN (9), Biochimie des Signaux R6gulateurs Cellulaires et Mol6culaires, Universit6 Pierre et Marie Curie, F-75006 Paris, France Sa'wVEr~ R. CROSBY (8), School of Biomolecular Sciences, Liverpool John Moores University, Liverpool L3 3AF, United Kingdom P. Dauci-i (21), Institut de Pharmacologie Mol6culaire et Cellulaire, Centre National de la Recherche Scientifique, 06560 Valbonne, France ROBERT DAY (2), Laboratory of Biochemical Neuroendocrinology, Clinical Research Institute of Montreal, Montreal, Quebec, Canada H2W 1R7 V. D I w (21), CEN de Saclay, 91191 Gif s/s Yvette, France ANN-MArtEE DUNCAr~ (19), St. Vincent's Institute of Medical Research, Fitzroy, Victoria 3065, Australia

ix

CONTRIBUTORS TO VOLUME 23

LLOYD D. FRICKER (13), Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, New York 10461 S. J. FULLER (18), Department of Pathology, University of Melbourne, Parkville, Victoria 3052, Australia HAROLD GAINER (10), Laboratory of Neurochemistry, National Institute of Neurological Disorders and Strokes, National Institutes of Health, Bethesda, Maryland 20892 ALISON M. GIBSON (16), Medical Research Council, Neurochemical Pathology Unit, Newcastle General Hospital, Newcastle Upon Tyne NE4 6BE, United Kingdom MARC J. GLUCKSMAN (17), Fishberg Research Center in Neurobiology, Mount Sinai School of Medicine, New York, New York 10029 SADAO KIMURA (14), Center for Biomedical Science, School of Medicine, Chiba University, Chiba 260, Japan ATHENA KLADIS (19), St. Vincent's Institute of Medical Research, Fitzroy, Victoria 3065, Australia ANNE C. LAWRENCE (19), Department of Biology, Medawar Building, University College London, London WC1E 6BT, United Kingdom REBECCA A. LEW (7, 12), Peptide Biology Laboratory, Baker Medical Research Institute, Prahran, Victoria 3181, Australia IRIS LINDBERG (5), Department of Biochemistry and Molecular Biology, Louisiana State University Medical Center, New Orleans, Louisiana 70112 MARK O. LIVELY (10), Department of Biochemistry, Bowman Gray School of Medicine, Wake Forest University, Winston-Salem, North Carolina 27157 RICHARD E. MAINS (6), Department of Neuroscience, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21205 C. L. MASTERS (18), Department of Pathology, University of Melbourne, Parkville, Victoria 3052, Australia Y. MASUO (21), Takeda Chemical Industries, Ltd., Pharmaceutical Group, Tsukuda 300-42, Japan JOHN R. MCDERMONT (16), Medical Research Council, Neurochemical Pathology Unit, Newcastle General Hospital, Newcastle Upon Tyne NE4 6BE, United Kingdom

CONTRIBUTORS TO VOLUME 23

xi

FREDERICK A. O. MENDELSOHN (22), Department of Medicine, University of Melbourne, Austin Hospital, Melbourne, Victoria 3048, Australia MARIANA MORRIS (10), Department of Physiology and Pharmacology, Bowman Gray School of Medicine, Wake Forest University, Winston-Salem, North Carolina 27157 TERRY J. OPGENORTH (14), Aging and Degenerate Disease Research, Abbott

Laboratories, Abbott Park, Illinois 60064 JOHN E. PINTAR (3), Department of Neuroscience and Cell Biology, University of Medicine and Dentistry of New Jersey-Robert Wood Johnson Medical School, Piscataway, New Jersey 08854 G. REED (18), Department of Pathology, University of Melbourne, Parkville, Victoria 3052, Australia MOHAMED RHOLAM (9), Biochimie des Signaux R6gulateurs Cellulaires et Mol6culaires, Universit6 Pierre et Marie Curie, F-75006 Paris, France JAMES L. ROBERTS (17), Fishberg Research Center in Neurobiology, Mount Sinai School of Medicine, New York, New York 10029 MARTIN K.-H. SCH,g,FER (2), Department of Anatomy and Cell Biology, Phillips University of Marburg, D-35037 Marburg, Germany NABIL G. SEIDAH (1), Laboratory of Biochemical Neuroendocrinology, Clinical Research Institute of Montreal, Montreal, Quebec, Canada H2W 1R7 ARTHUR SHULKES (15), Department of Surgery, University of Melbourne, Melbourne, Victoria 3084, Australia D. H. SMALL (18), Department of Pathology, University of Melbourne, Parkville, Victoria 3052, Australia A. IAN SMITH (7, 12), Peptide Biology Laboratory, Baker Medical Research Institute, Prahran, Victoria 318 l, Australia JULIANNE STACK (11), The Vollum Institute, Oregon Health Sciences University, Portland, Oregon 97201 GARY THOMAS (4), Vollum Institute, Oregon Health Sciences University, Portland, Oregon 97201 LAUREL THOMAS (4), Vollum Institute, Oregon Health Sciences University, Portland, Oregon 97201

~176

Xll

CONTRIBUTORS TO VOLUME 23

ANTHONY J. TURNER (20), Department of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT, United Kingdom JUDY K. VANSLYKE (4), Vollum Institute, Oregon Health Sciences University, Portland, Oregon 97201 B. VINCENT (21), Institut de Pharmacologie Mol6culaire et Cellulaire, Centre National de la Recherche Scientifique, 06560 Valbonne, France J. P. VINCENT (21), Institut de Pharmacologie Mol6culaire et Cellulaire, Centre National de la Recherche Scientifique, 06560 Valbonne, France A. WEIDEMANN (18), Center for Molecular Biology, University of Heidelberg, D-6900 Heidelberg, Germany JINSHYUN R. Wu-WON6 (14), Aging and Degenerative Diseases Research, Abbott Laboratories, Abbott Park, Illinois 60064 MIN ZHENG (3), Department of Anatomy and Cell Biology, Columbia University College of Physicians and Surgeons, New York, New York 10032 YI ZHOU (5), Department of Biochemistry and Molecular Biology, Louisiana State University Medical Center, New Orleans, Louisiana 70112

Preface

The generation of bioactive peptides from inactive precursor molecules involves a series of highly ordered, enzyme-mediated processing events. The posttranslational modifications can occur within the cell at the point of secretion or postsecretion. The precise pattern of processing for any given precursor also can vary, depending on the site of expression and/or stage of development, reflecting the differential expression of processing enzymes. The last decade or so has seen the characterization of both peptide products and the majority of the processing enzymes involved in their production, thus facilitating the development of various biochemical, immunological, and molecular probes necessary to characterize these complex pathways in greater detail. The aim of this volume in the Methods in Neurosciences series is to describe in a very practical way the "state-of-the-art" technology being developed and applied in the field of peptidases and neuropeptide processing. It is divided into three sections. The first, "Molecular Approaches for the Study of Intracellular Processing Enzymes," covers strategies for the molecular characterization of processing enzymes, including cloning, expression, localization by in situ hybridization, and the use of antisense mRNA to block enzyme expression. The second, "Immunological and Biochemical Approaches to the Study of Peptide-Processing Pathways," describes the combination of more classical approaches such as immunoassays, HPLC, and the use of specifically modified substrates to characterize both the precise pattern of peptide products in a given tissue and the regulation and distribution of the enzymes involved in their generation. Finally, the last section, "Identification and Characterization of Extracellular Processing Enzymes in the Central Nervous System," is designed to provide an insight into, as well as strategies for, the investigation of this exciting and developing area in which extracellular enzymes can generate, modulate, or terminate peptide signals in the central nervous system. In this book, like others in the series, the authors have been encouraged to provide chapters that reflect the latest techniques being developed in their laboratories, with their own specific scientific interests providing the practical application. Each chapter provides sufficient detail to allow the experimental procedures to be easily duplicated, although, for practical reasons, lengthy operating procedures for common laboratory equipment have been omitted. Absolute conditions for any given experiment are inevitably determined empirically; however, it is hoped that this volume will provide both

xiii

xiv

PREFACE

the student and experienced researcher a valuable starting point in developing strategies for the study of peptidases and neuropeptide processing. I would like to express my appreciation to the Baker Medical Research Institute for supporting the production of this work. Appreciation is also expressed to my fellow authors for the high standard of their contributions and for meeting their deadlines. A. IAN SMITH

Methods in Neurosciences

Volume 1 Gene Probes Edited by P. Michael Conn Volume 2 Cell Culture Edited by P. Michael Conn Volume 3 Quantitative and Qualitative Microscopy Edited by P. Michael Conn Volume 4 Electrophysiology and Microinjection Edited by P. Michael Conn Volume 5 Neuropeptide Technology: Gene Expression and Neuropeptide Receptors Edited by P. Michael Conn Volume 6 Neuropeptide Technology: Synthesis, Assay, Purification, and Processing Edited by P. Michael Conn Volume 7 Lesions and Transplantation Edited by P. Michael Conn Volume 8 Neurotoxins Edited by P. Michael Conn Volume 9 Gene Expression in Neural Tissues Edited by P. Michael Conn Volume 10 Computers and Computations in the Neurosciences Edited by P. Michael Conn Volume 11 Receptors: Model Systems and Specific Receptors Edited by P. Michael Conn Volume 12 Receptors: Molecular Biology, Receptor Subclasses, Localization, and Ligand Design Edited by P. Michael Conn Volume 13 Neuropeptide Analogs, Conjugates, and Fragments Edited by P. Michael Conn Volume 14 Paradigms for the Study of Behavior Edited by P. Michael Conn Volume 15 Photoreceptor Cells Edited by Paul A. Hargrave Volume 16 Neurobiology of Cytokines (Part A) Edited by Errol B. De Souza Volume 17 Neurobiology of Cytokines (Part B) Edited by Errol B. De Souza Volume 18 Lipid Metabolism in Signaling Systems Edited by John N. Fain Volume 19 Ion Channels of Excitable Membranes Edited by Toshio Narahashi

XV

xvi

VOLUMES IN SERIES

Volume 20 Pulsatility in Neuroendocrine Systems Edited by Jon E. Levine Volume 21 Providing Pharmacological Access to the Brain: Alternate Approaches Edited by Thomas R. Flanagan, Dwaine F. Emerich, and Shelley R. Winn Volume 22 Neurobiology of Steroids Edited by E. Ronald deKloet and Win Sutanto Volume 23 Peptidases and Neuropeptide Processing Edited by A. Ian Smith Volume 24 Neuroimmunology (in preparation) Edited by M. Ian Phillips and Dwight E. Evans Volume 25 Receptor Molecular Biology (in preparation) Edited by Stuart C. Sealfon Volume 26 PCR in Neuroscience (in preparation) Edited by Gobinda Sarkar

Section I

Molecular Approaches for the Study of Intracellular Processing Enzymes

This Page Intentionally Left Blank

[1]

Molecular Strategies for Identifying Processing Enzymes Nabil G. Seidah

Introduction In the early 1960s it was proposed that polypeptide hormones are first synthesized as inactive precursors that require specific cleavage after pairs of basic residues (such as LysArg-, ArgArg-, LysLys-, and ArgLys-) in order to release the active hormone. Since then this model has been extended to other precursors, as it is also applicable to progrowth factors, proneurotrophic factors, hormonal receptors, adhesion molecules, retroviral surface glycoproteins, proenzymes, and even certain protoxins. The elaboration of the structures of many precursors as well as their biosynthetic products also revealed that processing C terminal to single basic residues such as Arg(and less frequently Lys-) as well as after multiple basic residues (three or more) occurs in about 20% of the processed sites utilized in vivo. Therefore, it was of great interest to identify the proteinase(s) responsible for such proprotein processing and to define whether cleavage after monobasic residues and C terminal to pairs of basic residues was performed by the same enzyme(s). The search for the physiologically important processing enzymes, termed "proprotein convertases" or "PCs," was laborious and a number of laboratories, including our own, participated actively in this hunt (1). The major breakthrough came in 1984, with the molecular identification of the convertase responsible for the activation of the yeast a-mating factor and killer toxin. The proteinase identified by genetic complementation of a K E X 2 mutant strain was found to be a subtilisin-like serine proteinase (2, 3) and is now called "kexin." The search for the mammalian counterpart of kexin took about 5 years, before it was realized by computer database searches for sequence identity to kexin that a partial human genomic sequence encoding a protein called furin had already been reported by Roebroek et al. in 1986 (4). In the reported DNA sequence only the active site serine and the catalytically important asparagine residue found in all subtilisin-like proteases were identified. The complete sequence of the 5' end of the gene was completed in 1989 and it comprised the other two active site residues, aspartate and histidine (5).

Discovery of PC1 and PC2 Alignment of the amino acid sequences of furin and kexin within their catalytic domains revealed a number of segments exhibiting a high degree of Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

4

I

MOLECULAR APPROACHES hFurin

SGVTQRDLNVKAAWAQGYTGHGIVVSILDDGIEKNHPDLAGNYDPGASFD

174

yKexin

PSFPGSDINVLDLWYNNITGAGVV~~D_CzLDYENEDLKDNFCAEGSWD

196 224

9Z. . . .

J:ll"

I "-'Jl

I:l''J'llJ::

~:'11

:J s : : : l Z l

hFurin

VNDQDPDPEPRYTQMNDNRHGTRCAGEVAAVANNGVCGVGVAYNARIGGV

yKexin

F N D N T N L P K P R . . .L S D D Y H G T R C A G E I A A K K G N N F C G V G V G Y N A K I SG I 243

hFurin

R M L D G E V T D A V D A R S L G L N P N H I H I Y S A S W G P D D D G K T V H G P A R L A E E A F 274

yKexin

RI L S G D I T T E D E A A S L I Y G L D V N D IYS C S W G P A D D G R H L Q G P S D L V K K A L

hFurin

II o~ F R G V S Q G R G G L G S I F V W A S G N G G R E H D S C N C D G Y T N S IYTLS I S S A T Q F G

yKexin

V K G V T E G R D S KGAI Y V F A S G N G G T R G D N C N Y D G Y T N S I YS I T I GAI D H K D

9II..

I.II

I'I.I-'I..

:.I:

@

IIIIIIII:II

:I II ..

9:II.,II'-I.I'I'IIIIII

AS

@

.I..IIIII:III:I:I:

9 .III.IIII.III-"II.

I...I:

I.II-IIIIIIII.:.I'..:

(.-m I

(..

293

9

324 343

hFurin

NVPWYSEACSSTLATTYSSGNQNEKQ IVTTDLRQKCTESHTGTSASAPLA

374

yKexin

LHPPYSEGCSAVMAVTYSSG.. SGEYIHSSDINGRCSNSHGGTSAAAPLA

391

I III'II..'I.IIIII

.'. I ..I.'..'I.-II.IIII.IIII

FIG. 1 Alignment of the amino acid sequences of human furin and yeast kexin within the catalytic domain. The active site residues Asp", His", and Sera are emphasized, as well as the catalytically important Asne. The sense (S) and antisense (AS) oligonucleotides used to identify human PC2 are shown, as well as the primers (I and II) that were first used to identify mouse PC1 and PC2.1, Identical sequence; :, highly similar in sequence.

sequence identity (Fig. 1). In 1989, the partial sequence of furin (from the catalytically important Asn* up to the C terminus) (4) and the full sequence ofkexin (3) were known. Accordingly, on the basis of the concept of sequence conservation around the active sites of serine proteinases, polymerase chain reaction (PCR) amplification of mRNA (reverse transcriptase-PCR or RTPCR) allowed two laboratories simultaneously to isolate for the first time other mammalian homologs of kexin, known as PC1 (6, 7) and PC2 (6, 8), representing the first endocrine and neuroendocrine processing enzymes molecularly characterized in mammalian tissues. Polymerase chain reaction amplification of a cDNA synthesized from human insulinoma total RNA, using degenerate oligonucleotides encoding the consensus sequence surrounding the active site residues Asp" (oligo S; Fig. 1) and His" (oligo AS; Fig. 1) in kexin and related subtilisins, gave a 150-bp probe. The latter was used to screen a human insulinoma library and to isolate a full-length cDNA encoding a novel convertase called PC2 (8). Independently, PCR applied to cDNA obtained from mouse pituitary total RNA using oligonucleotides encoding the sequence around the catalytically important Asn* and the active site Ser u of human furin (oligos I and II, Fig. 1), allowed the isolation of a 260-bp probe (6, 7). Screening mouse pituitary and mouse insulinoma libraries

[1] PROPROTEIN CONVERTASES

5

with this probe led to the isolation of full-length cDNA clones encoding mouse PC2 (6) and also another convertase that was named PC1 [(6, 7); also called PC3 in Smeekens et al. (9)].

Polymerase Chain Reaction Procedure The PCR methodology used (5, 6) consisted first of reverse transcribing about 1-5/zg of total RNA obtained from tissues of interest (e.g., pituitary or cell lines) and then performing 30 cycles of PCR amplification using 100 pmol of each primer (e.g., oligos I and II; Fig. 1), 2.5 units of Taq DNA polymerase in 10 mM Tris (pH 8.3), 50 mM KCI, 1.5 mM MgC12, and 200 ~M dNTPs. The original cycling PCR program used consisted of successive 1-min incubations at temperatures of 94, 53, and 72~ in a Perkin-Elmer (Norwalk, CT) model 480 cycler. The amplified products were digested with restriction enzymes, for which sites were already encoded at the 5' ends of the chosen oligonucleotides. The digested products were then purified on a preparative 2% (w/v) agarose gel, size selected, and then subcloned in a vector of choice. This cumbersome cloning procedure has now been replaced by a simpler version, whereby the amplified products are directly ligated in the PCRII vector (Stratagene, La Jolla, CA) without restriction enzyme digestion. This is possible because the Taq polymerase always adds an extra A nucleotide at the 5' ends of the amplified cDNA and, hence, the use of a vector with T overhangs permits a rapid subcloning procedure. We recommend this protocol because it saves time and also circumvents the problem of having to add, at the 5' ends of the primers used in the PCR reaction, a restriction site that may also be present within the amplified segment.

D i s c o v e r y of P C 4 a n d PC5 Analysis of the deduced sequence homology between mammalian convertases PC1, PC2, and furin revealed that other segments are also conserved. In an effort to isolate other convertases, we developed a procedure that allowed us to identify three more members of this subtilisin/kexin-like family called PC4 (10) and PC5 (11) as well as rodent homologs of human PACE4 (12). As shown from the homology of the sequences of the six known convertases (Fig. 2), highly conserved segments are also found in regions other than those encompassing the active site Ser" and the catalytically important Asn ~ We have chosen a set of two degenerate oligonucleotides, one preceding the catalytically important asparagine (sense oligo IV) and the other following the active site serine (antisense oligo Ill) (Figs. 2 and 3). We found

6

I

M O L E C U L A R APPROACHES mPCI

Consensus

KeRsKRsVqk fdRkKRgyRd KRRtKRdVyq rRRvKRslv, KRRvKRqVR. KkRtKRdydl KRR-KR-VR-

dsalDL.FND ineiDinmND dPt ...... D vPt ...... D sdpQaLYFND sraQstYFND -P-QDLYFND

PmWnqQWYLq dTrmtaalpk PIFtkQWYLf nTgqadgtpg PkFpqQWYL ..... sgvtqr PwFskQWYM ..... nkeieq PiWsnmWYLH CgDknsrcrS PkWpsmWYMH CsDnthpcqS P .... QWYLH CTD ...... S

LDLhVipvWe LDLNVaeAWe .DLNVkAAWa .DLNIlkvWn .EMNVqAAWk .DMNIegAWk LDLNV-AAW-

mPCl mPC2 hfurin rPC4-A hPACE4 rPC5 Consensus

kGiTGKGVVI IGYTGKGVtI qGYTGhGIVV qGITGrGVVV rGYTGKnVVV rGYTGKnIVV -GYTGKGVVV

TVLDDGIEWN gIMDDGIDYI sILDDGIEkN sILDDGIEkd TILDDGIERN TILDDGIERt TILDDGIERN

HtDiyANYDP HPDLAyNYna HPDLAgNYDP HPDLwANYDP HPDLApNYDs HPDLmqNYDa HPDLAANYDP

eASYDfNDND dASYDfssND gASFDvNDqD IASYDfNDyD yASYDvNgND IASCDvNgND -ASYD-NDND

hDPFPRYdlt PyPYPRYtdd PDPePRYtqm PDPqPRYtpn yDPsPRYdAS IDPmPRYdAS PDP-PRY-AS

mPCl mPC2

rPC4-A hPACE4 rPC5 Consensus

NENKHGTRCA wfNsHGTRCA NDNrHGTRCA dENrHGTRCA NENKHGTRCA NENKHGTRCA NENKHGTRCA

GEIAmqANNh GEVsAaAsNn GEVAAvANNg GEVsATANNg GEVAAsANNs GEVAATANNs GEVAATANN-

kCGVGVAYNs iCGVGVAYNs vCGVGVAYNA fCGaGVAFNA yCiVGIAYNA hCtVGIAFNA -CGVGVAYNA

KVGGIRMLDG KVaGIRMLDq rIGGVRMLDG rIGGVRMLDG KIGGIRMLDG KIGGVRMLDG KIGG-RMLDG

i.VTDaIEAs pfmTDIIEAs E.VTDaVDAr a. ITDIVEAq D.VTDVVEAk D.VTDmVEAk D-VTDIVEA-

mPCI mPC2 hfurin rPC4-A hPACE4 rPC5 Consensus

SigFNPgHVd SishmPQIId SIgLNPnHIh SIsLqPQHIh SlgirPnyId SvsYNPQHVh S--LNPQHI-

IYSASWGPnD IYSASWGPtD IYSASWGPDD IYSASWGPED IYSASWGPDD IYSASWGPDD ~ D D

DGKTVEGPGR nGKTVDGPre DGKTVhGPaR DGrTVDGPGI DGKTVDGPGR DGKTVDGPap DGKTVDGPGR

LaQkAFEyGV LtlQAmadGV LaeeAFfrGV LtQeAFrrGV LakQAFEyGI LtrQAFEnGV L-QQAFE-GV

KqGRQGkGSI nKGRgGkGSI sqGRgGLGSI tKGRQGLGtl KKGRQGLGSI rmGRrGLGSV KKGRQGLGSI

mPCI mPC2

FVWASGNGGR YVWASGdGG. FVWASGNGGR FIWASGNGGI FVWASGNGGR FVWASGNGGR ~ R ~II~

qgDNCdCDGY syDdCNCDGY ehDsCNCDGY hyDNCNCDGY egDyCsCDGY skDhCsCDGY --DNCNCDGY

TdSIYTISIS asSmWTISIn TNSIYTISIS TNSIhTISVg TNSIYTISVS TNSIYTISIS TNSIYTISIS

SAsqQGIsPW SAindGRtal SATqfGnvPW StTrQGRvPW SATenGykPW StaesGkkPW SAT-QGR-PW

YaEkCSSTLA YdEsCSSTLA YsEaCSSTLA YsEaCaSTFt YIEeCaSTLA YIEeCSSTLA Y-E-CSSTLA

TsYSSGDYtD sTFSnGrkrn TTYSSGnqnE TTFSSGvvtD TTYSSGaFyE TTYSSGEsyD TTYSSG---D

qr..ItsaDL peagVaTTDL kq..IVTTDL pq..IVTTDL rk..IVTTDL kk..IITTDL .... IVTTDL

hndCTEtHTG TSASAPLAAG ygnCTlrHsG TSAaAPeAAG RQkCTEsHTG TSASAPLAAG hhqCTDkHTG TSASAPLAAG RQRCTDgHTG TSvSAPMvAG RQRCTDnHTG TSASAPMAAG RQRCTD-HT~t__~G (- I (-

IfALALEANP VfALALEANI IIALtLEANk mIALALEANP IIALALEANs IIALALEANP IIALALEANP

nLTWRDMQHL dLTWRDMQHL nLTWRDMQHL ILTWRDLQHL qLTWRDvQHL fLTWRDvQHv - L ~ (-III(-

VVWTSeydpL tViTSkrnqL VVqTSkPAHL VVRaSRPAqL IVkTSRPAHL IVRTSRagHL VVRTSRPAHL

asN.pgWKkN hdevhqWrrN NAN..DWatN qAe..DWriN kAs..DWKvN NAN..DWKtN NAN--DWK-N

GaGLmVnsrF GvGLefnHLF GvGrKVSHsW GvGrqVSHhY GaGhKVSHFY aaGFKVSHLY G-GLKVSHLY

GFGLLnAkAL GYGvLDAGAM GYGLLDAGAM GYGLLDAGIL GFGLvDAeAL GFGLMDAeAM G-GLLDAGA-

mPCl mPC2 hfurin rPC4-A hPACE4 rPC5 Consensus

VDLAdpRTwr VkMAkdW..k VaLAqnWT.. VDLArvWl.. VveAkKWT.. VmeAeKWT.. VDLA-KWT--

nVPekkeCVV TVPerfhCVg TVapQrKCII ptkpQkKCtI aVPsQhmCVa TVPqQhvCVe TVP-Q-KCVI

kdnnfEPral gsvq.nPekI dilt.EPkdI rvvh.tPtpI asdk.rPrsI stdr.qiktI ..... EP--I

kangEVivei PPtgklvlTl gkrlEVRKT, iPrmlVpKn, PlvqvlRtTa rPnsaVRsiy PP--EVRKT-

pTrACEgqEN kTnACEgkEN vTaclgepnh vTvcCDgsrr iTSACaehsd kaSgCsdnpN -TSACE--EN

mPC1 mPC2 hfurin rPC4-A hPACE4 rPC5 Consensus

a. IksLEHVQ .FVRYLEHVQ ..ItrLEHaQ rLIRsLEHVQ qrVvYLEHVv hhVnYLEHVv ---RYLEHVQ

feaTIeYsRR GDLhVtLTSa aviTVnatRR GDLnInMTSP aRITISYnRR GDLAIhLvSP VqlslSYsRR GDLeIFLTSP VRtsIShpRR GDLqIYLvSP VRiTIthpRR GDLAIYLTSP VR-TISY-~YLTSP (-V~

vGTstvLLAe MGTkSiLLsr MGTRStLLAa MGTRStLvAi sGTkSqLLAk sGTRSqLLAn MGTRS-LLA-

Rer.DtSpnG RPrdDdSkvG RPh.DySaDG RPL.DiSgqG RIL.DlSnEG RIF.DhSmEG RPL-D-S-EG

mPC3

hfurin rPC4-A hPACE4 rPC5

hfurln

hfurin

rPC4-A hPACE4 rPC5

Consensus mPCI mPC2

hfurin

rPC4-A

hPACE4

rPC5 Consensus mPCl mPC2 hfurin rPC4-A hPACE4

rPC5 Consensus

VI

e

IV~

[1]

PROPROTEIN Ile

mPCI mPC2 hPC2 hFur mFur rFur rPC5 mPC5 rPC4 mPC4 hPACE4

Tyr Ser Ala

ATT

TAC

- -C

.....

--C

--C --C - -C

--C

AGT

GCA

C

- -C

Consensus

ATC

Ser

AGC

Trp G l y P r o

TGG

C . . . . . C --C --T G-.....

C

GGC

--C --C

T

TAC

AG-

C

SAnme

C C

C

-C-

C - -T

A

C

C

C

G AG-

T

TGG

Oliaonucleotide

GGC

CC

IV

Thr Arg

Cys A l a

G l y Glu

ACA

GGA

GAA

--C

--C

C --T

TGT

hPace4

--C --C ...... G

--T --C

--G --G

----G

Consensus

T C TAG C A - G G - A C - -GC A ACA

rPC5

RGD

S*nse

--G C-G C-C C-T C-C

..... C --C --T ..... G ..... C

TGT

A A GC- GGG C

Oliaonucleotide

VI

-T

C-G

- -A

Met

ATG

C-G C-G C-G

ACC

TGG

AA - G- G A C C G

Gln

CAA

.....

G G

Leu CTG

His

CAT C

G G G .... G G .... G C .... G C-A --G G-C --G

--C --C ---

A -T G

T CA- CTG

G

G CAA

----G-T G-T -------

--C --C --C

C

<-- <-- e - <-- <-- <-- <-- <-- <-Antisense Oliaonucleotide III

Gly

rPC2 rFurin rPC4

GAC

--G

GGA

GCA

Asp

C G

CAT

AGA

Arg

AGA

- -A

His

rPCI

Trp

TGG

A A

--C --C

GC-

Thr

ACC

C

C --T - -T

CCT

- -C

- -C --C --C

7

CONVERTASES

GAA

Arg

Arg

Gly

Asp

CGT A-G --C --C --C --C

AGA

GGA

GAC

C-T C-C C C-C

--T --G --G

Ala CTT CAT G A-C ..... G GC..... G G-C C --G ..... G G-G

T A T G CGC CGA GGA GAC G C T

4-- ~

~-

Antisense

~

Leu

T GCT

CTG

CAG

C A

C

~r- 4-- ~--

Oliaonucleotide

Ile GTC A-A-A-A-A--

G

ATC

4-V

FIG. 3 Rationale for the consensus oligonucleotide sequences III-u The alignment of the nucleotide sequences of the six convertases in the segments preceding the Asn ~ (oligo IV) and following the Set u (oligo Ill), as well as those surrounding the His i (oligo VI) and RGD sequence (oligo u is shown.

that this pair of oligonucleotides, when combined with PCR amplification, allowed us to amplify a cDNA fragment of about 450 bp for any of the six convertases known. These conserved IIeTyrSerAlaSerTrpGlyPro and ThrTrpArgAspMetGlnHisLeu sequences, which did not include the active site residues A s n ~ and Ser i , are separated by a relatively constant number

FIG. 2 Comparison of the amino acid sequences of the six known convertases, from the autocatalytic cleavage site ( $ ) up to the RGD sequence. The active site residues Asp i, His i, and Ser m are emphasized, as well as the catalytically important Ash ~ The six oligonucleotides chosen in the RT-PCR reactions on the basis of the consensus alignment sequence are shown (oligos I to VI). In this representation, identical amino acids are in capital letters, whereas differing residues are shown in lower case.

8

I MOLECULAR APPROACHES

of amino acids in all the subtilisin-like proteinases studied (about 152 amino acids; see Fig. 2). These criteria were useful in eliminating false positives on DNA sequencing of the subcloned 450-bp PCR-amplified fragments, as it was possible to eliminate unrelated structures if one or both of these sites were absent in the sequenced fragment. This approach allowed us to identify, by RT-PCR amplification of total RNA isolated from rat and mouse testis, two novel subtilisin/kexin-like structures that were called PC4 and PC5 (1, 10, l 1). These 450-bp probes were then used to screen cDNA libraries obtained from testis and adrenocortical Y1 cells to isolate the corresponding full-length cDNA sequences of PC4 (10) and PC5 (11). In both cases, it was difficult to isolate the complete cDNA sequence from all the libraries tested. For example, for PC5, we isolated from the Y1 cell cDNA library a clone of only 2.2 kb, which was missing the exon encoding the catalytically important Asn ~ To complete the sequence we made use of the conserved ArgArgGly AspLeu sequence found in all convertases (Fig. 2). Accordingly, using a sense oligonucleotide bearing the conserved active site His" in the sequence HisGlyThrArgCysAlaGlyGlu (Fig. 3, oligo VI) and an antisense degenerate oligonucleotide containing the RGD sequence (oligo V), we were able to deduce the structure of the missing mouse PC5 fragment (11). Similarly, using the same RGD oligonucleotide (oligo V), we also isolated a fragment of rat PACE4 cDNA, which is now being used in the laboratory to perform comparative in situ hybridizations between PC5 and PACE4 in the rat central nervous system (CNS) and in peripheral organs.

Improvements in Polymerase Chain Reaction Methodology Some improvements in PCR methodology were introduced during this work and are worth mentioning. These include the addition of 5-8% (v/v) dimethyl sulfoxide (DMSO) in the PCR reaction tube to reduce the number of nonspecific cDNAs amplified. In addition, we advocate the use of hot start procedures in the PCR reaction. Typically, we found it convenient to include, in the PCR amplification tube, l0 mM Tris (pH 8.3), 50 mM KC1, 1.5 mM, MgC12, 200 ~M dNTPs, 10-100 ng of the cDNA to be amplified (usually obtained following RT-PCR) and 5% (v/v) DMSO, the whole volume being adjusted to 75/~1 with sterile water. The DNA in this tube is first denatured by heating at 94~ for 5 min and then kept at 80~ for 2 min, together with another tube that contains 100 pmol of each of the two PCR primers and 2.5 units of Taq DNA polymerase in a final volume of 25 ~l. At the end of the 2 min, the 25 ~l is rapidly transferred to the 75/~l-containing tube by sliding the liquid along the side wall of the tube, and the PCR amplification cycles are allowed to begin. By this procedure, the primers and DNA are never

[1]

9

PROPROTEIN CONVERTASES

bp

I

I

I

I

I

I

1018 510 394 344 298

FIG. 4 Polymerase chain reaction amplification of all six convertases, using oligos Ill and IV. The PCR amplification was performed with a Perkin-Elmer model 980, with about l0 ng of each full-length cDNA of either mPC1, mPC2, hfurin, hPACE4, rPC4, or mPC5. The migration of molecular size standards (bp) is also shown. The program consisted of a hot start with 30 cycles of amplification in the presence of 5% (v/v) DMSO at the denaturation, annealing, and extension temperatures of 94~ (20 sec), 53~ (1 min), and 72~ (1 min), respectively.

together at a temperature more than 5~ below the theoretical melting temperature of the complex, that is, [4(G + C) + 2(A + T)]~ The addition of DMSO and use of the hot start procedure drastically diminish nonspecific amplification products. It was also useful in some cases to use a "touchdown PCR" amplification program, in which the annealing is performed in the first cycle at the highest temperature possible (e.g., 60~ and then, in the subsequent l0 cycles, the annealing temperature is gradually diminished by about 0.5~ to reach 55~ which is then kept constant for the next 20 cycles. This procedure improved background and also allowed us in some cases to amplify rare mRNA products [e.g., PC5 (11)]. In Fig. 4, we show a typical example of the use of the above-described PCR procedure and oligos Ill and IV to amplify similar 450-bp segments of any one of the six known convertases [mouse PC1 (mPC1), mouse PC2 (mPC2), human furin (hfurin), rat PC4 (rPC4), human PACE4 (hPACE4), and mouse PC5 (mPC5)], starting with about l0 ng of their respective cDNAs. Notice that the amplification of hPACE4 and mPC5 was much less efficient that that of either mPC1, mPC2, hfurin, or rPC4. A likely explanation for this observation is the presence of either an A (in mPC5) or a G (in hPACE4) at the third 3' end nucleotide as compared to a C in the consensus sequence of the sense oligo IV (see figure 3). This example serves as a model to show how small sequence variations close to the 3' ends of the oligonucleotides used for PCR can have a dramatic effect on the ability to amplify a transcript. Therefore, it would not be surprising if the RT-PCR methods used to amplify sequences related to the subtilisin/kexin family of convertases " m i s s " certain

10

I MOLECULAR APPROACHES

transcripts that in some species have a slightly different sequence from the one used in the degenerate consensus oligonucleotides.

Cloning of Distantly Related Convertases" Aplysia PC2 and Xenopus PC 1 The technique of PCR amplification, using the combination of oligos III and IV, also allowed us to clone distantly related members of this family of convertases such as the mollusk Aplysia californica PC2 convertase (13). In this particular case we used the primers III and IV to amplify the fragment from DNA obtained from a hZAPII cDNA library of the nervous system of Aplysia. Here again, the sequenced 450-bp fragment served as a probe to screen a cDNA library of bag cells and to isolate the full-length clone of A. californica PC2 (13). In another approach, based on the knowledge that in the intermediate lobe of the rodent pituitary PC1 and PC2 were colocalized (6, 7) and coregulated (14) with proopiomelanocortin (POMC), pituitary mRNAs were isolated from Xenopus laevis frogs that had been dark adapted for 15 days, causing an increase in the production of a-melanocyte-stimulating hormone (a-MSH) and a darkening of their skin. Applying RT-PCR to pituitary mRNAs along with oligos I and u allowed us to isolate a 543-bp segment containing the active sites residues His i and Ser i as well as the catalytically important Asn~ of X. laevis PC1 (Fig. 5). This sequence exhibits about 92 and 83% identity to mouse PC1 at the amino acid and nucleotide levels, respectively. It is interesting that, using a subtractive library approach, only Xenopus PC2 was identified, and not PC1 (15). This attests to the versatility and power of the RT-PCR methodology to isolate rare mRNA transcripts.

Production of Fusion Proteins for Antibody Production Once a convertase has been identified and its structure determined, it becomes necessary to express it in different cell lines and to analyze its biosynthesis, posttranslational modifications, enzymatic characteristics, and tissue distribution. For the purpose of determining tissue distribution, Northern blots, in situ hybridization histochemistry, and immunocytochemical procedures have been used to define the locations of the mRNA and the protein. For biosynthetic studies, radiolabeling of the expressed cellular convertase in different cell lines and protein immunoprecipitation are used to define the intracellular organelle where the zymogen autoactivation step occurs (16).

l1

[1] PROPROTEIN CONVERTASES HisGlyThrArgCysAlaGlyGluIleAlaMetIleAlaAsnAsnAsnLysCysGlyVal CATGKL%AC.KAGATGT~.AGK~TGAAATTGCGATGATTGCCAACAACAATAAATGTGGCGTC

60

GlyValAlaTyrAsnAlaLysValGlyGlyIleArgMetLeuAspGlyValValThrAsp GGAGTTGCATATAATGCAAAAGTTGGGGGTATACGGATGCTGGATGGTGTTGTGACAGAT

120

AlaLysGluAlaArgThrIleGlyPheAsnProGlnHisValHisIleTyrSerAlaSer GCCAAAGAAGCAAGAACCATTGGCTTCAATCCACAACATGTGCACATATACAGTGCTAGC

180

TrpGlyProAsnAspAspGlyLysThrValGluGlyProGlyArgLeuAlaGlnLysAla TGGGGACCTAATGATGATGGCAAAACCGTAGAAGGACCTGGCAGACTGGCTCAGAAAGCT

240

PheGluTyrGlyIleLysGlnGlyArgAsnGlyLysGlySerIlePheValTrpAlaSer TTTGAGTATGGCATCAAACAGGGGCGAAATGGAAAAGGTTCCATATTCGTTTGGGCATCT

300

GlyAmmGlyGlyArgGlnGlyAspAsnCysAspCysAspGlyTyrThrAspSerIleTyr GGAAATGGAGGCCGACAGGGTGATAACTGTGACTGTGATGGATACACAGACAGCATTTAC

360

ThrIleSerIleSerSerAlaSerGlnGlnGlyLeuSerProTrpTyrAlaGluLysCys ACCATCTCTATCAGCAGTGCTTCCCAGCAAGGTTTGTCTCCCTGGTATGCGGAGAAATGC

420

SerSerThrLeuAlaThrAlaTyrSerSerGlyAspTyrThrAspGlnArgIleIleSer TCTTCCACCCTGGCTACAGCATACAGCAGTGGCGACTACACCGATCAGAGAATTATAAGC

480

AlaAspLeuHisAsnGluCysThrGluThrHisThrGlyThrSerAlaSerAlaProLeuAla GCTGACCTACACAATGAGTGCACGGAGACGCACACTGGCACCTCGGCTTCA(~ACCCCT(~ICT <--<--<--4--<--4--<--<--

543

FIG. 5 Deduced partial amino acid sequence of Xenopus PC1. The positions of the sense (4) and antisense (~) primers used for the RT-PCR amplification of Xenopus total RNA isolated from pituitaries of dark-adapted flogs are shown by arrows. The active site residues His" and Sera are emphasized, as well as the catalytically important Asne.

All of these studies require the production of either specific cRNA probes or antibodies against various segments of the convertase of interest. The production of cRNA probes for in situ hybridization and the choice of probes are detailed in [2] in this volume. Briefly mentioned here is the fusion protein method we have chosen for the production of specific antisera against the C-terminal segments of PC1 and PC2. These fusion protein antibodies were used for immunocytochemical investigations both at the light and electron microscopy levels and for biosynthetic studies of these convertases (16). To amplify template cDNAs of mPC1 and mPC2, we used two pairs of oligonucleotides encompassing the borders of segment 629-726 of mPC1 (7) and segment 529-637 of mPC2 (6). The sequences of the (BamHI)sense/antisense-(EcoR1) oligonucleotides were GTGGATCCGATGAG CAGCAGGTACAAGGAACT / CAGAATTCTTAATTCTCCTCATTTAG GAT for mPC1 and CCGGATCCGGCACCAAGTCCATTTTGCTG/CG GAATTCGTAGTTCTTTCTCAGGATACT for mPC2. Following PCR amplification and purification on a 2% (w/v) agarose gel, the amplified 300-bp fragments were digested with the restriction enzymes BamHI and EcoRI prior

12

I

MOLECULAR APPROACHES

to their ligation into the pGEX-2T vector [encoding bacterial glutathinone Stransferase (GST)] digested with the same enzymes (Pharmacia, Piscataway, NJ). Transformation into Escherichia coli bacteria and selection of recombinant clones allowed the amplification of the required cDNA. Nucleotide sequence analysis of a number of clones allowed the selection of a clone in each case devoid of nucleotide changes due to Taq polymerase errors (which in our hands amounts to an average of 1 error for every 400 nucleotides per clone). The selected bacterial recombinants were grown in the presence of 25 /xM isopropyl-fl-D-thiogalactopyranoside (IPTG) for 3-6 hr, and the bacteria were collected by centrifugation. Because the recombinant products were sequestered in the bacterial inclusion bodies as insoluble proteins, we treated the bacteria with lysozyme (100/zg/ml) and then incubated this mixture with 0.2% (w/v) N-lauroylsarcosine for 15 min, followed by another 15-min incubation in 2% (w/v) Triton X-100 and a 20-min incubation in the presence of 1.25 mM MgC12 and 1 mM CaC12. The resulting extract was centrifuged at 50,000 g for 2 hr and the supernatant was removed and diluted twofold in phosphate-buffered saline. The fusion proteins were then affinity purified on a glutathione-agarose column, according to the protocol of the manufacturer (Pharmacia). The yield of the glutathione S-transferase-fusion proteins was between 15 and 35 mg/liter of bacterial culture. The inserted 100 amino acids were then excised from the fusion protein by thrombin digestion and purified by reversed-phase high-performance liquid chromatography (HPLC) on a C~8 column. Their identity was verified by amino acid analysis and protein sequencing. Rabbits were then injected, using the purified fusion proteins as immunogens. This method allowed us to obtain highly specific C-terminal PC1 and PC2 antibodies with titers of about 1" 10,000 (17). A similar approach was also used to generate antibodies against fusion proteins of GST with PC4 and PC5 segments. However, one of the major drawbacks of this technique is the need to choose the segment that does not contain thrombin cleavage sites. Alternatively, one may use another vector requiring a more specific enzyme for the excision of the insert. For example, pGEX3X could be used and the insert excised with factor Xa, which has the recognition sequence IleGluGlyArg $ (Pharmacia). The generated antibodies can then be used to define the issue and subcellular localization of the convertases in order to also define their colocalization with a suspected precursor substrate, as was done for PC1 and PC2 in the pituitary (18).

Coexpression of Convertase with Suspected Substrate One of the criteria that must be fulfilled for a protein to be classified as a proprotein convertase is its ability to cleave precursors at single, specific basic residues or pairs of basic residues. Therefore, once the amino acid

[1] PROPROTEIN CONVERTASES

13

sequence of the convertase is deduced, it becomes imperative to express this proteinase and to assess its cleavage selectivity toward a number of potential substrates that have been demonstrated to colocalize with the convertase in vivo. In this respect the use of either cellular transfection or infection techniques to coexpress enzyme and substrate has been recorded in the literature. In our laboratory, as well as in that of G. Thomas (see [4] in this volume), vaccinia virus recombinants of each identified convertase have been used to study their cleavage specificity (1). So far, we have coexpressed convertases PC1, PC2, and furin with about seven precursors including POMC, proenkephalin, prodynorphin, prosomatostatin, pro-parathyroid hormone (pro-PTH), pro-7B2, and prorenin. The overall results demonstrate that these enzymes are indeed highly specific convertases, cleaving precursors at specific and sometimes distinct pairs of basic residues. Furthermore, in the case of prodynorphin, cleavage after a single arginine residue has also been demonstrated for both PC1 and PC2. Aside from establishing that these enzymes are specific convertases, the work undertaken also demonstrated that some bonds can be cleaved only in cells containing secretory granules, whereas others can occur within the trans-Golgi network in both constitutive and regulated cells. Therefore, to assess accurately the ability of a given convertase to cleave a precursor, we must perform coexpression studies in both types of cells. A typical case is human renin, which can be cleaved by PC 1, but not by PC2 or furin, and only in cells containing secretory granules. Another example involves prosomatostatin, which is processed into somatostatin-14 (SS-14) by PC1 in both constitutive and regulated cells; but PC2 cleaves it to SS-14 only in regulated cells. The general conclusions drawn from all of the present studies showed the following:

1. Furin is mostly involved in the processing of precursors exiting from the cells by the constitutive pathway, whereas PC1 and PC2 cleave proproteins routed to the regulated pathway. However, in the case of precursors in which the cleavage site contains an Arg-X-(Lys/Arg)Arg $ sequence, these proteins may be processed by furin even though they may end up in secretory granules (e.g., pro-7B2 or pro-PTH), suggesting that in these kinds of precursors cleavage may precede cellular sorting into the constitutive or regulated pathway. 2. Each enzyme is capable of cleaving certain precursors at any combination of pairs of basic amino acids (e.g., LysArg, ArgArg, ArgLys, and Lys Lys), and it is the sequence around the pairs that defines which enzyme will be the convertase chosen in vivo. Furthermore, furin, PC1, and PC2 are able to cleave after single arginine residues in certain proproproteins, implying

14

I

MOLECULAR APPROACHES

that it is possible that some monobasic and dibasic convertases may be of the subtilisin/kexin type. 3. In general, we find that PC1 and furin cleave precursors to generate relatively larger fragments than those produced by PC2. The latter enzyme usually releases small peptides such as a-MSH, Met- and Leu-enkephalin, and dynorphins A and B. However, the combination of enzymes such as PC1 and PC2 may be necessary for the efficient processing of precursors into their final tissue-specific products. 4. Although coexpression of precursor and substrate in different cell lines may indicate the ability of a convertase to cleave a given precursor, it is by no means proof that such processing occurs in vivo in every tissue where the substrate is expressed. A typical example is human prorenin, which can be efficiently processed by PC 1; however, this convertase does not colocalize with renin within the kidney juxtaglomerular cells. Therefore, before accepting the role of a given convertase in the processing of a precursor, careful colocalization studies should complement the cleavage selectivity data obtained from cellular coexpression experiments. To this must also be added other corroborative evidence, including comparative developmental expression and coregulation of the expression of the precursor substrate and its presumed convertase(s).

Conclusions There has been an explosion of information on the nature and properties of a new family of prohormone convertases of the subtilisin/kexin type. So far, only six mammalian members have been described but already diversity generated by the differential splicing of the genes encoding some of these convertases is beginning to emerge, such as in the case of PC4, PACE 4, and PC5. The ability to devise methods to screen for the presence of these convertases in various tissues and pathological cases may allow us to study their predicted functions in proliferative diseases, neurological pathologies, and viral infections. Even though other convertases may be discovered in the future, it is clear that with the available set of enzymes we may begin to probe the complex cellular problem of precursor processing. We hope that the years to come will bring about fruitful clinical and pharmacological applications of these convertases in the diagnosis and possible treatment of specific pathologies.

References 1. N.G. Seidah, R. Day, M. Marcinkiewicz, S. Benjannet, and M. Chr6tien, Enzyme 45, 271 (1991).

[1] PROPROTEIN CONVERTASES

.

10. ll. 12. 13. 14. 15. 16. 17. 18.

15

R. S. Fuller, R. E. Sterne, and J. Thorner, Annu. Rev. Biochem. 50, 345 (1988). K. Mizuno, T. Nakamura, T. Oshima, S. Tanaka, and H. Matsuo, Biochem. Biophys. Res. Commun. 156, 246 (1988). A. J. M. Roebroek, J. A. Schalken, J. A. M. Leunissen, C. Onnekink, H. P. J. Gloemers, and W. J. M. Van de Ven, EMBO J. 5, 2197 (1986). W. J. M. Van de Ven, J. Voorberg, R. Fontijn, H. Pannekoek, A. M. W. van den Ouweland, H. L. P. van Duijnhoven, A. J. M. Roebroek, and R. J. Siezen, Mol. Biol. Rep. 14, 265 (1990). N. G. Seidah, L. Gaspar, P. Mion, M. Maracinkiewicz, M. Mbikay, and M. Chr6tien, D N A Cell Biol. 9, 415 (1990). N. G. Seidah, M. Marcinkiewicz, S. Benjannet, L. Gaspar, G. Beaubien, M. G. Mattei, C. Lazure, M. Mbikay, and M. Chr6tien, Mol. Endocrinol. 5, l l l (1991). S. P. Smeekens and D. F. Steiner, J. Biol. Chem. 265, 2997 (1990). S. P. Smeekens, A. S. Avruch, J. LaMendola, S. J. Chan, and D. F. Steiner, Proc. Natl. Acad. Sci. U.S.A. 88, 340 (1991). N. G. Seidah, R. Day, J. Hamelin, A. Gaspar, M. W. Collard, and M. Chr6tien, Mol. Endocrinol. 6, 1559 (1992). J. Lusson, D. Vieau, J. Hamelin, R. Day, M. Chr6tien, and N. G. Seidah, Proc. Natl. Acad. Sci. U.S.A. 90, 6691 (1993). M. C. Kiefer, J. E. Tucker, R. Joh, K. E. Landsberg, D. Saltman, and P. J. Barr, D N A Cell Biol. 10, 757 (1991). T. Ouimet, A. Mammarbachi, T. Cloutier, N. G. Seidah, and V. CasteUucci, FEBS Lett. 330, 343 (1993). R. Day, M. K. H. Sch~ifer, S. J. Watson, M. Chr6tien, and N. G. Seidah, Mol. Endocrinol. 6, 485 (1992). J. A. M. Braks, K. W. C. Guldemond, M. C. H. M. van Riel, A. J. M. Coenen, and G. J. M. Martens, FEBS Lett. 305, 45 (1992). S. Benjannet, N. Rondeau, L. Paquet, A. Boudreault, C. Lazure, M. Chr6tien, and N. G. Seidah, Biochem. J. 294, 735 (1993). A. Boudreault, N. Rondeau, N. G. Seidah, M. Chr6tien, and C. Lazure, J. Biol. Chem. (submitted for publication) (1994). M. Marcinkiewicz, R. Day, N. G. Seidah, and M. Chr6tien, Proc. Natl. Acad. Sci. U.S.A. 90, 4922 (1993).

[2]

In Situ Hybridization Techniques to Map

Processing Enzymes Martin K.-H. Sch~ifer and Robert Day

Introduction Great progress has been made in the molecular identification of the enzymes involved in posttranslational processing of neuropeptides. The vast majority of neuropeptides derive from large precursor molecules, which are exposed to a cascade of enzymatic processing events shortly after their synthesis and during their intracellular transport in a tissue-specific fashion. These steps include the initial partial proteolysis by the family of prohormone convertases (1), the removal of the basic amino acid residues by carboxypeptidase E (2), and modifications such as a-amidation at the glycine-extended C terminus by enzymes derived from the peptidylglycine ct-amidating monooxygenase (PAM) gene (3), sulfatation, and acetylation, among others. The molecular characterization and the availability of sequence information have enabled us very early in our studies to investigate the tissue- and cell-specific distribution of these enzymes. The use of the now well-established method of in situ hybridization histochemistry has made it possible to investigate with cellular resolution the gene expression of processing enzymes at the mRNA level. This approach has rapidly generated qualitative and quantitative data~almost simultaneously with the publication of the primary amino acid sequences~on the tissue-specific expression of many of these enzymes, before specific antibodies were available (4-7). Furthermore, in situ hybridization studies have provided important complementary data to the biochemical in vitro studies to elucidate the role of prohormone convertases in the tissue- and substrate-specific cleavage pattern. This has been particularly true for the prohormone convertases PC 1 and PC2, which play an important role in the tissue-specific processing of the proopiomelanocortin (POMC) precursor in the pituitary gland (8). Biochemical results, which demonstrated that PC1 generated POMC fragments found in the rat anterior lobe and that PC2 produced a cleavage pattern observed in the intermediate lobe (9), have been supported by in situ hybridization studies showing the presence of PC 1 mRNA in anterior lobe corticotrophs, but the absence of PC2 (10). Thus, knowing the cleavage properties of a prohormone convertase one can quickly predict from the tissue-specific distribution pattern based on in situ hybridization studies which processed forms of the precursor of interest should be 16

Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

[2] I N SITU HYBRIDIZATION TECHNIQUES

17

present in specific areas or cells. Furthermore, a unique tissue distribution of a processing enzyme may help the biochemist to design experiments to elucidate the enzyme specific properties. In this chapter we give a practical perspective on the in situ hybridization procedure, which has become a standard method in many laboratories, not only in those with histological facilities. We do not emphasize the theoretical aspects and the molecular biological framework of nucleic acid hybridization, which have been discussed in great detail elsewhere (11-15); rather, we focus on the description of specific procedures, which are used in our laboratories to investigate the tissue- and cell-specific expression of neuropeptide-processing enzymes.

Selection, Design, and Synthesis of Probes Probe Selection For the successful localization of the enzyme mRNA of interest, a selection of the type of probe used in the hybridization experiment is a critical first step. The three types of probes typically used are oligonucleotides, cDNA, and cRNA probes. Each class of probe offers certain advantages and disadvantages, which are summarized in Table I. The abundance of the mRNA in question, alternative spliced mRNA forms, and sequence homologies to other members of an enzyme family are important factors that require consideration. Riboprobes In our mapping studies we have relied mainly on complementary, singlestranded RNA probes (cRNA), which were generated from specially designed transcription vectors containing viral RNA polymerase promotors and multiple cloning sites for the insertion of cDNA fragments of interest. These cDNA fragments were obtained from cDNA clones or via the reverse transcriptaseassisted polymerase chain reaction (RT-PCR) as applied to RNA amplification (see [1] in this volume). Particularly for the localization of low-abundance messages such as furin and other prohormone convertases, riboprobes offer distinct advantages over oligonucleotides. Long single-stranded transcripts up to several kilobases can be generated to extremely high specific activity (105 Ci/mmol), which results in at least a 10- to 100-fold higher sensitivity of detection in comparison to oligonucleotides. In addition, RNA-RNA hybrids form highly stable duplexes and show high specificity because they are resistant to RNase A digestion, which allows the removal of the unhybrid-

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TABLE I

Comparison of Hybridization Probes i

Probes

Advantages

Oligonucleotides

Acquisition easy (custom synthesis) Good tissue penetration

cDNA

cRNA

Easy combination of multiple probes ("cocktail") High freedom of design High specific activity Signal amplification with overlapping strands Good for cross-species hybridization High specific activity Single stranded Highest sensitivity Very stable hybrids Suitable for quantitation (incorporation of label can be exactly calculated) Enzymatic removal (RNase) of non-specific bound probe

Disadvantages Dependent on published DNA sequence information Low sensitivity (due to low specific activity) Low hybrid stability

Double strand reannealing Variations in probe length (random priming) Sophisticated mol. biol. setup Instable probes RNase free environment Probe "stickiness" Intolerant to mismatches Bad for cross species hybridization

ized probe by posthybridization treatment with RNase A. However, this feature can turn into a disadvantage when using riboprobes for cross-species hybridization, because even small numbers of mismatches can affect the hybrid stability dramatically, as they present a target for the action of RNase. Finally, riboprobes are suitable for quantitative estimates of probe-mRNA hybrid concentration in a tissue section, which are based on the uniform labeling reaction, the specific activity of the label, and the nucleotide sequence of the transcribed cDNA insert.

Oligonucleotides If cloning facilities are not established in the laboratory, oligonucleotides provide an acceptable alternative. They can be easily generated from any published nucleotide sequence. Most laboratories have access to an automated nucleotide synthesis facility and there are enough commercial companies providing fast, reliable, and relatively inexpensive custom service. The disadvantage of oligonucleotides is their low hybrid stability, low specific activity, and therefore lower sensitivity. The latter is critical to a successful experiment, particularly when looking at relatively low-abundance messages such as the prohormone convertases (6, 8, 10). Oligonucleotide probes have been successfully applied to the localization of carboxypeptidase E mRNA

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in the central nervous system (CNS) (16). In this study three different oligonucleotides complementary to the rat carboxy-peptidose E (CPE) mRNA were combined as a "probe cocktail" to overcome the problem of lower sensitivity of oligonucleotides, an approach that has previously been established for the localization of protein kinases in the CNS (17).

Design of RNA Probes For distribution studies in the central nervous system the rat is generally the species of choice among neuroscientists. Therefore, at the beginning of our studies, cDNA fragments specific to the rat sequences of the processing enzymes were obtained. In some cases the rat cDNA clone was not available for a specific enzyme, but the sequence information in another species (e.g., mouse) was known. Therefore, the RT-PCR technique was used to produce cDNA from isolated reverse-transcribed mRNA using oligonucleotides as specific primers (see [1] in this volume). These PCR-amplified cDNA fragments were then subcloned into the appropriate transcription vectors. For subcloning and general molecular biological techniques we followed previously described procedures (14, 18). After construction of the plasmids the constructs were transformed into bacteria. Amplification of single colonies in bacterial culture and isolation of the plasmids were carried out according to standard procedures (18) to generate enough template for probe synthesis.

Synthesis and Labeling of Riboprobes RNA probes are labeled during their synthesis by in vitro transcription of linearized plasmid constructs, using a prokaryotic RNA polymerase (19). Prior to transcription the circular plasmid construct is digested by a restriction enzyme downstream (3') of the polymerase promotor to yield an RNA transcript of the desired length from the linearized DNA template. Restriction enzymes producing 3' overhangs should not be used, because SP6, T7, and T3 polymerases may transcribe from the opposite strand (20). The RNA polymerase will recognize its promoter, which is located on the 5' end of the insert, and transcribe single-stranded RNA molecules from the cDNA in the 5'-to-3' direction, until the end of the linearized plasmid is reached. Radioactive-labeled, nonradioactive-labeled, and unlabeled ribonucleotides are incorporated if a concentration above 13/zM in the reaction mixture is maintained. Probes of extremely high specific activity can be produced when one or more radiolabeled nucleotides are present in the reaction mix. The consistency of the transcription reaction generates full-length transcripts

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with uniform labeling and fixed probe length, which are essential for accurate quantitation of mRNA transcripts detected in tissue.

In Vitro Transcription For the novice, it is best to obtain an in vitro transcription kit [Promega (Madison, WI) or Boehringer Mannheim (Indianapolis, IN)] that contains all necessary reagents and enzymes, excluding the radioactive label. We follow the general protocol described below for in vitro transcription of 35S-labeled cRNA probes with high specific activity. In a sterile, small Eppendorf tube 200 pmol of [35S]UTP (New England Nuclear, Boston, MA) is aliquoted and dried down in a Speed-Vac (Savant, Hicksville, NY). The radioactive pellet is resuspended to a final reaction volume of 10/zl containing the following: 4.5/zl RNase-free water Transcription buffer (Boehringer Mannheim), 10x" 1.0/zl 0.4 M Tris-HC1 (pH 7.5), 60 mM MgC12, 20 mM spermidine, 0.1 mM NaC1 1.0/xl Linearized DNA template (1/zg) NTP-UTP (ATP, GTP, CTP), 5 mM 1.0/xl Dithiothreitol (DTT; Sigma, St. Louis, MO), 100 mM 1.0/xl RNase inhibitor (Promega), 20 U 0.5/zl RNA polymerase (SP6, T7, or T3) (Promega or Boeh- 1.0/xl ringer Mannheim) The reaction is incubated for 60 to 90 min at 37~

Nonradioactive Labeling o f R N A Probes For the synthesis of nonradioactive labeled probes, biotin, fluorescein isothiocyanate (FITC), or digoxigenin (Boehringer Mannheim) is used as the reporter molecule. In particular, the digoxigenin system from Boehringer Mannheim has been successfully applied to the detection or relatively lowabundance messages in the CNS, such as nerve growth factor receptor, using immunological detection systems (21). For the labeling of cRNA probes with digoxigenin the same transcription protocol is used as described above, with the following changes. 1. Omission of the [35S]UTP 2. Replacement of the NTP-UTP mix with a digoxigenin (DIG)-NTP mix

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from Boehringer Mannheim (10 mM ATP, 10 mM GTP, 10 mM CTP, 6.6 mM UTP, and 3.3 mM DIG-UTP) 3. Extention of reaction time to 2 hr at 37~

Purification o f Probes

At the end of the transcription reaction 1/xl of RNase-free DNase I (Boehringer Mannheim is added for 15 min at 37~ to remove the DNA template. Immediately after its completion 80/xl of water and 10/xl of 100 mM DTT are added to the transcription reaction. The labeled probe in the reaction mix can now be purified from the unincorporated radioactive label by (a) ethanol precipitation as described below or (b) by purification over a Sephadex G-50 column or spin columns (Pharmacia, Piscataway, NJ) according to the protocol provided by the manufacturer.

Probe Hydrolysis

Riboprobes greater than 500 nucleotides in length are generally subjected to a mild alkaline hydrolysis to reduce their size, which improves their tissue penetration considerably and therefore gives stronger hybridization signals. The following protocol, previously described (22), is followed: dissolve labeled probe in 50 tzl of RNase-free water. Add 50 tzl of sodium carbonate buffer, pH 10.2 (80 mM NaHCO3-120 mM NazCO3) , and incubate at 60~ for the time t calculated using Eq. (1): t = (L o - L f ) : ( k • L o • Lf)

(1)

where t is the incubation time (in minutes), L 0 is the initial probe length (in kilobases), Lf is the final probe length (in kilobases) (0.2 to 0.4 kb), and k is the hydrolysis rate constant (0.11 scissions/kilobase/min at 60~ (For example, the 1244-nucleotide rat furin cRNA probe was subjected to hydrolysis for 15 rain to reduce the length to 400 nucleotides.) Neutralize the hydrolysis reaction by adding 3/xl of 3 M sodium acetate (pH 6.0) and 5/xl of 10% (v/v) glacial acetic acid. Precipitate by adding 2.5 vol of 100% ice-cold ethanol for 30 rain at -20~ and spin for 15 rain a microfuge at 15,000 rpm. Remove the supernatant to the radioactive waste and wash the pellet with 70% ethanol. Dry and resuspend the pellet in 50/xl of 10 mM Tris-HC1-1 mM ethylenediaminetetraacetic acid (EDTA; pH 8.0) containing 100 mM DTT. The size of the hydrolyzed probes can be determined on a denaturing formaldehyde-2%

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(w/v) agarose gel or a 4% (w/v) polyacrylamide sequencing gel (8 M urea). Count a 1-/xl aliquot in a/3 counter. Store the probe undiluted or diluted in the hybridization buffer at -70~ until use. Detailed examples for synthesis of riboprobes specific for furin and the prohormone convertases PC1 and PC2 are given below. Furin

A 1228-nucleotide cDNA clone obtained from a hgtl 1 rat liver library was subcloned into a transcription vector pSP72 (Promega), using standard molecular biological procedures described in detail elsewhere (18). The clone corresponded to the previously described rat furin cDNA from position 1111 to 2338 (23). The plasmid was linearized with NcoI. Transcription of this plasmid yielded a 320-nucleotide cRNA probe in antisense orientation. Another furin probe corresponded to nucleotides 691-1935 of the rat cDNA clone and had to be exposed to limited alkaline hydrolysis for 15 min as described above to decrease the probe length to 300-400 nucleotides to allow better tissue penetration. Using [35S]UTP as radioactive label, specific activity (SA) was calculated to be 600,000 Ci/mmol. This probe was used to examine the furin mRNA distribution in the rat brain (see Fig. 1). PC1

The rat PC1 cDNA was obtained by RT-PCR. Total RNA from rat anterior pituitary was reverse transcribed. For PCR amplification oligonucleotides comprising mouse PC1 (mPC1)-specific sequences were used" nucleotides 715-737 (forward primer: 5' GGT GAA ATT GCC ATG CAA GCA AAT 3') and mPC1 nucleotides 1206-1186 (reverse primer: 5' GGTTCTCTGTG CAGTCATTGTG 3'). The amplified 492-bp segment subcloned into the transcription vector Bluescript KS + (Stratagene, La Jolla, CA) was completely sequenced and showed a 97% identity to the corresponding mPC 1 sequence. Radiolabeled cRNA had a specific activity of 250,000 Ci/mmol. PC2

Two rat-specific PC2 probes were generated by PCR amplification of a cDNA from a rat striatum library (Stratagene) and rat neurointermediate lobe total RNA, respectively. The first probe represents a 450-bp cDNA obtained using mPC2 nucleotides 878-900 (forward primer: 5' GAC ATC TAC AGT GCA AGC TGG GG 3') and mPC2 nucleotides 1326-1301 (reverse primer: 5' AGA TGT TGC ATG TCT CTC CAG GTC AG 3') (specific activity, 240,000 Ci/ mmol). The second probe represents a 3'-specific probe, 425 bp in length, using mPC2 nucleitdes 1584-1604 (forward primer: 5' AAA ACT TCG TCC GCT ACC TGG 3') and PC2 nucleotides 2008-1988 (reverse primer: 5' CTA GTT CTT TCTCAG GAT ACT 3').

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PC5

An 837-bp fragment (nucleotides 1089-1925) was subcloned into PCRII vector by the T/A cloning procedure (Invitrogen, San Diego, CA). The specific activity of the partially hydrolyzed probe was 270,000 Ci/mmol. PACE4

A 534-bp fragment corresponding to nucleotides 1153-1687 of the published sequence was subcloned into PCRII (Invitrogen) according to the manufacturer protocol (specific activity, 300,000 Ci/mmol). PAM

The PAM probe corresponded to nucleotides 164-1197 of the rat cDNA clone (24, 25) (specific activity, 300,000 Ci/mmol). CPE

The CPE probe was complementary to nucleotides 1080-1470 of the PCRgenerated rat cDNA clone (2) (specific activity, 95,000 Ci/mmol).

Labeling of Oligonucleotides Oligonucleotides are labeled by adding radioactive or nonradioactive labeled nucleotides to either their 5' or 3' ends (26). Labeling of oligonucleotides can also be performed directly during their synthesis by incorporation of fluorescent or biotin-labeled nucleotides, which is still a relatively expensive procedure. We prefer the 3' end labeling (tailing reaction) with radiolabeled or nonradioactive-labeled nucleotides, using terminal deoxynucleotidyltransferase (tdT) (15, 27). In a standard labeling reaction distilled water, 5 • tailing buffer [500 mM potassium cacodylate (pH 7.2), 10 mM COC12, 1.0 mM DTT], oligonucleotide (final concentration, 0.1 /zM), [35S]dATP (final concentration, 1/zM; New England Nuclear), and tdT [75-100 units; Mannheim or Bethesda Research Laboratories (Gaithersburg, MD)] are added to a final volume of 50/xl in an Eppendorf vial. Incubation is carried out at 37~ for 30 min (batches at various time points should be checked to assess activity and number of nucleotides added). The reaction is stopped by the addition of 380/zl of TE (10 mM Tris-HC1, 1 mM EDTA, pH 7.6), 20 ~1 of 5 M NaC1, and 1 pA of tRNA (20 mg/ml). The sample is extracted twice with phenol-chloroform-isoamyl alcohol (75:29:1, v/v). The aqueous phase is transferred to a new tube and precipitated with 2.5 vol of ethanol on dry ice for 30 min. After centrifugation the pellet is rinsed in 70% ice-cold ethanol. The pellet is dried to remove the remaining alcohol and dissolved in 50/xl

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of TE and 1/zl of 5 M DTT. To calculate specific activity, 1/xl of the labeled oligonucleotide solution is counted in a scintillation counter (~4C spectrum).

Tissue Preparation By working with a variety of different pretreated tissues from animals and humans, including frozen tissue postfixed on the slide, immersion and perfusion-fixed tissues, and tissue embedded in paraffin or plastic, we found that fresh-frozen cryostat sections postfixed on the slide gave optimal results in our distribution studies. It should be pointed out that the optimal hybridization procedure is always a compromise between sufficient morphological preservation of the tissue to identify the labeled cells and RNA detectability. Tissue samples are removed from anesthesized animals, mounted with O.C.T. (Lab-Tek; Miles, Elkhart, IL) in proper orientation on plastic molds, and immediately frozen either in isopentane cooled to - 3 0 to -50~ or in liquid nitrogen (if samples are small enough to crack). The block is remounted on a block holder on dry ice, transferred to the cryostat chamber to equilibrate to the cutting temperature, and cut at the desired thickness (15-/xm sections at - 12 to - 16~ Sections are picked up from the knife holder with gelatinor polylysine-coated glass slides and stored at -80~ until further processing.

Subbing of Slides Gelatin Coating New glass slides are placed into slide racks and soaked for 30 min in hot tap water with a glassware detergent. Soap is washed off under running hot tap water for 30 min, followed by several changes in distilled water. In a dust-free area slides are drained and immersed in the subbing solution for 2 min. Subbing solution is always prepared fresh by diluting 5 g of gelatin in 500 ml of distilled water, which is warmed to 37~ to which 0.5 g of chromium potassium sulfate (chromalum) is added under stirring. The solution is filtered and used warm. Dipped slides are dried overnight at temperatures not exceeding 37~

Lysine Coating New glass slides are cleaned as described above. Twenty-five milligrams of poly(L-lysine) (P-1399; Sigma) is dissolved in 500 ml of 10 mM Tris, pH 8.0. Slides are incubated in this solution for 10 min and then dried overnight at 50~

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Fixation Tissue must be preserved to maintain sufficient morphology and to retain the RNA in the tissue. Rapid fixation is important to prevent the degradation of RNA by intrinsic RNases. A variety of different fixatives have been used in hybridization experiments (formaldehyde, glutaraldehyde, Zenker's, Bouin's, acetic acid-ethanol, Carnoy' s, and Karnovsky) (28). Formaldehyde fixation has proved to be the most reliable method of tissue preservation in terms of RNA retention and reasonably maintained morphology. However, other fixatives have been used successfully when pretreatment conditions were adequately adjusted. In our procedure fresh-frozen, 10- to 20-/zm-thick cryostat sections are immersed in freshly prepared 4% paraformaldehyde buffered with 10 mM phosphate-buffered saline (PBS), pH 7.4, and incubated for 60 min at 4~

Preparation of Formaldehyde Solution Prepare 10• PBS solution (100 mM: 13.8 g of NazHPO 4 9H20, 3.12 g of NaHzPO4, 75.97 g in 1000 ml) and adjust to pH 7.4. To prepare 1 liter of paraformaldehyde solution warm 500 ml of distilled water to 65~ and dissolve 40 g of paraformaldehyde under vigorous stirring. Slowly add drops of 6 M NaOH until the solution has cleared. Add 500 ml of 2x PBS and filter the solution through a Whatman (Clifton, NJ) No. 2 filter. Adjust the pH to 7.4 and store at 4~ The temperature of the formaldehyde solution should not exceed 68~ and should not be stored too long before use, because it will polymerize.

Tissue Storage Frozen unfixed tissues should be stored at -80~ In our experience hybridization signals can be obtained from pretreated fixed tissue sections stored desiccated at 4 or -20~ for several weeks, and from fresh tissue stored at -80~ for over 1 year.

Prehybridization Treatments Prehybridization treatments of tissue sections serve two purposes" they (a) improve accessibility of the RNA in the tissue and (b) reduce nonspecific binding of the probe.

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Denaturing To make RNA more available for hybridization slides are incubated in 0.2 N HC1 for 5 min at room temperature and rinsed in distilled water for 5 min.

Proteinase K Treatment Tissue digestion with proteases is an alternative method to the above, which we routinely use in our protocol. A 10-mg/ml proteinase K (PK) stock solution is prepared in PK buffer [0.1 M Tris-HC1, 50 mM EDTA (pH 8.0)]. For tissue digestion 25/zl of stock solution is diluted in 500 ml of PK buffer 30 min before use to yield a final concentration of 0.1/zg/ml. Digestion is carried out at room temperature for 10 min. The optimal concentration of PK and the digestion time depend on the section thickness, tissue fixation, and embedding and should be determined empirically. If tissue is very susceptible to PK digestion, this step is replaced by a 10min incubation in PBS containing 0.4% (v/v) Triton X-100.

Acetylation The acetylation step blocks nonspecific binding of the probe to positively charged free amino groups. Slides are placed in 0.1 M triethanolamine (TEA) (T-1377; Sigma), pH 8.0, for 1 min and transferred to a fresh batch of TEA containing 0.25% (v/v) acetic anhydride (A6404; Sigma) and incubated under vigorous stirring for 10 min at room temperature. Slides are removed and rinsed in 2x SSC (1 • SSC is 0.15 M NaC1 plus 0.015 M sodium citrate).

Dehydration and Delipidation Slides are dehydrated through a graded series of ethanol (50, 70, 95, and 100%), immersed in chloroform for 5 min, rinsed in 100% ethanol, 95% ethanol and then air dried. We typically apply the probe diluted in hybridization buffer immediately after the prehybridization. However, tissue sections can be stored for several days at room temperature and in a desiccated form for several months at -20~ prior to hybridization.

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Hybridization Hybridization conditions must be chosen in such a way that the probe can interact optimally with the RNA of interest in the tissue. Several factors can influence the efficiency of the hybridization reaction, and are described in detail elsewhere (12, 14, 29-31). These include hybridization temperature, probe concentration, composition of the hybridization buffer, and hybridization time. Hybridization temperature is usually chosen at 20 to 25~ below the theoretical Tm (predicted temperature at which 50% of the formed hybrids are dissociated), determined by Eq. (2) (valid for probes longer than 50 nucleotides):

Tm = 81.5~ + 16.61(logM) + 0.41(% GC) - 820/L - 0 . 6 ( % F) 1.4(% mismatch) -

(2)

where M is the concentration of monovalent ion in the hybridization buffer, GC is the number of guanidine and cytosine bases, L is the length of probe, and F is formamide. Increasing formamide and decreasing salt concentrations have a destabilizing effect on hybrid formation, that is, they increase the stringency of the hybridization conditions.

Preparation of Hybridization Mix Final concentrations are 75% formamide (50% for oligonucleotides), 3 • SSC, 50 mM sodium phosphate (pH 7.4), 1 mM EDTA, 1 x Denhardt's solution [0.2% (w/v) bovine serum albumin (BSA), 0.02% (w/v) Ficol1400, and 0.02% (w/v) polyvinylpyrrolidone], yeast tRNA (0.1 mg/ml), and sheared salmon sperm DNA (0.1 mg/ml. Riboprobes are diluted in the hybridization mix to a final concentration of 5 x 106 to 5 x 107 dpm/ml hybridization solution and dithiothreitol (DTT) is added to the final concentration of 10 mM. The addition of DTT is important when using 35S-labeled probes to prevent the formation of covalent disulfide bonds, which causes irreversible nonspecific binding of the probe to sulfated groups in the tissue. This solution can be stored at -80~ for several weeks. We have found that the degradation of the radiolabeled probes in the hybridization solution is greatly reduced.

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Hybridization Conditions To each slide 30/zl of hybridization solution is added, followed by coverslipping and sealing with rubber cement. Slides are placed in a Nunc (Roskilde, Denmark) culture dish with moistened paper towels to maintain humidity. Alternatively, soaking the paper towels in 50% formamide solution the sealing step can be omitted. The containers are transferred to the hybridization oven and hybridized for 16 to 24 hr at 55 to 60~ For oligonucleotide hybridization a lower temperature of 37 to 42~ is chosen.

Posthybridization Treatments Posthybridization procedures consist of a sequence of washes with decreasing salt concentrations to remove nonhybridized and nonspecifically bound probe from the tissue sections. When riboprobes are used, an incubation step in RNase buffer is inserted to digest single-stranded, nonhybridized probe, which dramatically reduces background.

Procedure for cRNA Probes Slides are removed from the humid chambers and immersed in a beaker containing 2 • SSC. The rubber cement is gently removed with forceps until the coverslips come loose. Slides are collected in a slide rack in 2x SSC and then transferred to another dish containing 2• SSC and incubated for 10 min. Then sections are immersed in RNase buffer [RNase A (20/~g/ml) and RNase T1 (1 U/ml) in 10 mM Tris-HC1 (pH 7.6), 0.5 M NaC1, 1 mM EDTA] for 30 min at 37~ After RNase digestion slides are washed sequentially in 2 • 1 • and 0.5 x SSC for 10 min each, followed by a 1-hr wash in 0.1 x SSC at the hybridization temperature (60~ Sections are quickly rinsed in distilled water, dehydrated in 50, 70, and 95% ethanol, and air dried.

Procedure for Oligonucleotides After hybridization coverslips are removed as described above and sections are washed in 2• SSC. Then slides are washed twice (30 rain each) in 2• SSC 50% formamide at 42~ Two washes in 0.5 x SSC at room temperature follow. After the final wash slides are briefly dipped in water to remove excess salt and then air dried.

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D e t e c t i o n of H y b r i d i z a t i o n Signal Radioactive hybridization signals may be detected by film or nuclear emulsion. Films are especially suited for screening tissues for enzyme expression, carrying out rapid semiquantitative area analysis, and estimating the duration of autoradiogram exposure. Nuclear emulsions allow the study of individual cells or heterogeneous cell populations and are used for simultaneous detection of two or more mRNAs in combination with nonradioactive procedures.

X-Ray Film The dry slides are fixed on cardboard and placed in an X-ray cassette. A sheet of X-ray film is laid over the sections. For rapid results we generally use Kodak (Rochester, NY) XAR-5 film, which offers high sensitivity but poor resolution. Film is exposed for 24 to 96 hr and then developed. For mapping studies well-resolved images are obtained with Amersham Hyperfilm/3-max (Arlington Heights, IL) and Du Pont Cronex 4 film (Wilmington, DE), which are of finer grain. Photographic prints can be made directly from the X-ray image.

Liquid Emulsion Autoradiography For microscopic analysis, the hybridized sections are coated with nuclear emulsion (Kodak, Rochester, NY or Ilford, London, UK). All procedures are performed in a darkroom illuminated only with a safe light (Kodak Safelight No. 2). The working area should be as light tight as possible, because nuclear emulsion is sensitive to light.

Preparation of New Emulsion A new batch of Kodak NTB2 emulsion (112 ml) is melted in a prewarmed waterbath at 42~ Before dilution a few native slides are dipped to check the quality of the emulsion. The completely melted emulsion is then diluted 1 : 1 (v/v) with 600 mM ammonium acetate (prewarmed to 42~ mixed with a magnetic stirring bar, and aliquoted into 20-ml plastic scintillation vials. The vials are wrapped twice in aluminum foil and placed in a light-tight box, which is wrapped in two black plastic bags and stored at 4~ Slides double labeled with 35S and DIG followed by an alkaline phosphatase reaction are dipped in Ilford K5 emulsion. This emulsion does not tolerate repeated melting, which causes increases in background. Therefore the

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amount of gel needed for one dipping session should be taken from the unmelted container and diluted 1:1 (v/v) at 42~ with water in the dipping container.

Dipping Slides An aliquot of the emulsion is melted in the water bath at 42~ The melted emulsion is poured into a dipping chamber (Electron Microscopic Sciences, Fort Washington, PA). After a few minutes blank slides are coated with emulsion to remove remaining air bubbles. Then the hybridized slides, which have been arranged into racks, are dipped by immersing them vertically, with a steady slow motion, into the emulsion. This procedure is repeated once. Excess emulsion is allowed to drip back into the emulsion and the bottom edge of the slide is touched by a paper towel. Slides are placed upright into a scintillation vial box to dry completely (several hours to overnight). Slides are then transferred to black slide boxes that contain desiccant capsules and sealed with electrical tape, wrapped in foil, and stored at 4~ for various exposure times, using the X-ray exposure as a guideline.

Developing All photographic solutions are cooled in the dark room to 14~ Slide boxes are allowed to warm to room temperature to avoid the formation of condensation. Slides are carefully placed into carriers and developed in Kodak D19 developer with water for 4 min, rinsed in water for 30 sec, and fixed in Kodak fixer for 5 min. Developed autoradiograms are washed in running cold water for 15 min.

Counterstaining and Mounting To identify specific cell types and tissues a relatively weak counterstaining must be carried out, which does not mask the autoradiographic signal (silver grains). Sections of nervous tissue are usually counterstained with Cresyl Violet solution [0.5% (w/v) in sodium acetate, pH 3.5] for 5 min. Other tissues are stained with hematoxylin-eosin (HE) or Giemsa staining [5% (v/v) for 2 min]. However, HE counterstaining can cause the emulsion to peel off and can mask the autoradiographic signal owing to its high staining intensity. The slides are dehydrated through graded series of ethanols (70, 95, and 100%) and incubated in xylene twice and coverslipped with DPX (Fluka, Buchs, Switzerland) or Permount (Fisher Scientific, Pittsburgh, PA).

Photography Microscopic examination and documentation are performed with a Zeiss (Thornwood, NY) Axiophot microscope. For black-and-white photography we use Ilford 400 film, which has sufficient speed for dark-field photography.

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For color slide documentation a Kodak 64 tungsten slide film or Fuji RTP is used.

Nonradioactive Detection The detection of nonradioactively labeled probes depends on the marker molecule used. We describe here the detection procedure for probes labeled with the antigen digoxigenin (DIG; Boehringer, Mannheim), which has been successfully used in colocalization studies of prohormone convertases with substrates such as proopiomelanocortin (POMC) in the pituitary and melaninconcentrating hormone (MCH) in the hypothalamus (see Fig. 4).

Immunological Detection of Digoxigenin-Labeled Probes The procedures for detecting DIG-labeled riboprobes and oligonucleotides are identical and based on procedures previously described (15, 21, 32). After the final wash of the posthybridization treatment slides are incubated in a solution of 2x SSC, 0.05% (w/v) Triton X-100, and 2% (v/v) normal sheep serum (NSS) (S-2382; Sigma) overnight. The sections are then washed in buffer 1 [100 mM Tris-HC1 (pH 7.6), 150 mM NaC1] twice for 10 min each, followed by a 5-hr incubation at room temperature with the alkaline phosphatase-conjugated anti-digoxigenin antibody diluted 1 : 1000 in buffer 1 [adjusted to 0.3% (w/v) Triton X-100, 1% (v/v) NSS]. A thorough wash is carried out in buffer 1 for 10 min. Then the tissue is equilibrated in buffer 2 [100 mM Tris (pH 9.5), 50 mM MgCI2, 100 mM NaC1, 1 mM levamisole) for 10 min. The final enzymatic reaction is carried out by applying the chromogen solution consisting of 45/~1 of nitroblue tetrazolium (NBT; 75 mg/ml in 70% dimethy|formamide) and 35 /~1 of 5-bromo-4-chloro-3-indolyl phosphate (BCIP; 50 mg/ml in dimethylformamide) in 10 ml of buffer 2. Incubation is performed in humid chambers in the dark for several hours until a visible, robust blue precipitate has developed. Development times depend on the abundance of the RNA under investigation, but should not be carried out for longer than 2 days because high background and nonspecific precipitation of the dye solution will occur. The color reaction is stopped in 10 mM Tris-HCl (pH 8.0), 1 mM EDTA and sections are rapidly dehydrated and mounted in glycerol for preliminary inspection or permanently coverslipped using Aquamount (Polysciences, Warrington, PA), because the color precipitate is not stable in xylene-based embedding media.

Specificity Controls In any in situ hybridization experiment, the proper controls are required to exclude the possibility of false-positive hybridization signals. Below, controls

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are listed that we routinely use in our experiments. Rationales in designing controls have been described previously (11, 14).

Positive Hybridization Controls Whenever possible, hybridization of a probe to tissue where the mRNA of interest is abundant should be included in the experiment as a positive control. For example, pituitary or brain sections are always included in our mapping studies ofprohormone convertases in novel tissues. When available, a second probe complementary to a different region of the mRNA under investigation should give the same labeling pattern. This control has been used in distribution studies of PAM gene expression in the CNS (24). However, if alternative splicing events occur, one must be careful when using this control, particularly with oligonucleotides, that the target sequences for both probes are present in the investigated mRNA form.

Negative Controls RNase Pretreatment Tissue sections are treated with RNase buffer as described above prior to hybridization, which should abolish all specific signal. However, this control demonstrates only that the probe has single-stranded RNA as target.

Sense Strand Probes Hybridization is carried out with message sense RNA probes transcribed from the same transcription vector as the antisense probe, but in the opposite direction. Sense strand probes have the same GC content and about the same length as the antisense form, thereby controlling for nonspecific stickiness of the antisense probe under investigation.

Temperature Sensitivity RNA-RNA hybrids melt across a temperature range that can be predicted with considerable accuracy from the hybridization kinetics dependent on the structure of the cRNA-mRNA hybrid, in particular probe length and GC content. Therefore temperature modulation provides an excellent tool with which to test stringency sensitivity. This stringency sensitivity is assayed by gradually increasing the posthybridization wash stringency (e.g., increasing the temperature and formamide concentration, and lowering the salt concentration). RNA hybrids should tolerate temperatures less than 15~

[2] I N SITU HYBRIDIZATION TECHNIQUES

33

below the Tm without major loss of signal intensity; at 15~ above the Tm the majority of the hybrids should be dissociated, causing a dramatic loss of signal. Excessive loss of hybridization signal at low wash temperatures suggests mismatches between probe and tissue mRNA. A low Tm may thus indicate some homology of probe-mRNA over short stretches of their sequence, which mandates closer examination of the in situ conditions employed.

Double-Labeling in

Situ

Hybridization

Double-labeling techniques are used to examine the presence of two or more mRNAs in the same cell or tissue. This is particularly important for the characterization of prohormone convertases, to identify specific sets of enzymes that may act on the precursor of interest. Another goal of hybridization colocalization studies is the identification of substrates for a specific enzyme. This can be done by simultaneous demonstration of the mRNA encoding the peptide precursor and the enzyme mRNA in the same cell. To perform these studies the different mRNA species must be visualized by detection methods giving different reaction signals during microscopic examination. The combination of radiolabeled RNA probes with nonradioactively labeled probes (e.g., digoxigenin) is the procedure with the greatest sensitivity at present. The following strategy has been employed in our laboratory. The mRNA that is lower in abundance (in most cases the enzyme) is detected with the radiolabeled probe, and the more highly abundant mRNA is detected with the DIG-labeled probe. Both probes are diluted in the same hybridization buffer (35S-labeled probe at a concentration of 1.5 to 3 x 10 6 dpm/30/xl, the DIG-labeled probe at 0.5/xg/ml). Hybridization procedures are carried out as described above. The last posthybridization wash is followed by the immunological detection of the DIG-labeled hybrids as detailed above. After the final step the double-hybridized, air-dried sections are dipped in Ilford emulsion if alkaline phosphatase has been used as reporter enzyme. If peroxidase-conjugated antibodies were used to detect the DIG hybrids (which is less sensitive) both the Ilford and Kodak emulsions can be used. After the appropriate exposure time autoradiograms are developed as described and coverslipped. A typical example of double in situ hybridization (ISH) is shown in Fig. 4.

Quantitation Strategies The detection of mRNA by autoradiographic techniques in combination with computer-aided image analysis methods allows a qualitative, semiquantita-

34

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M O L E C U L A R APPROACHES

tive, and quantitative measurement of mRNA concentration. In general, mRNA levels can be considered a measure of the biosynthetic activity, with some restrictions (e.g., RNA degradation and translation efficiency). Comparing relative or absolute levels of expression for two mRNAs is particularly important for prohormone convertases, where it has been shown that the relative levels of two or more PCs may determine the posttranslational processing pattern of a precursor. Quantitative measures are obtained either from (a) densitometry of digitized images from X-ray films or emulsioncoated autoradiograms or (b) grain counting of microscopic images, which has been described in detail elsewhere (33). We focus here on densitometric analysis of X-ray films and autoradiograms using computer-assisted image analysis. We have used both a Macintosh system based on software (Image 1.47) developed by W. Rasband at the National Institutes of Health (NIH, Bethesda, MD) and a PC-based system (M4 developed by Imaging Research, Inc., Toronto, Ontario, Canada).

Preparation of Radioactive Standards A requirement for any quantitative or semiquantitative analysis is the availability of a set of radioactive standards and the construction of a standard curve to relate optical densities (ODs) to radioactivity. For the comparison of relative amounts of signals from X-ray films, ~4C-based polymer standards (/3 decay profile similar to 35S) are commercially available (Amersham; ARC, St. Louis, MO). We make standards from brain paste labeled with L[35S]methionine (Amersham). Fresh whole brains are homogenized. Increasing amounts of L-[35S]methionine are thoroughly mixed with the brain paste to give a concentration range of 1000 (1 • 10 6 dpm in the hottest standard), molded in plastic tubes, and frozen in isopentane as described above. Cryostat sections are cut at the same thickness as the tissue and mounted on subbed slides. Usually 8 to 12 standards across the chosen concentration range are constructed.

Standard Curve Construction Standard curves are generated by establishing a relationship betwen OD measures and radioactivity. In our procedure, images from one set of standard sections are captured with a video camera from the light box or the microscope at the same magnification used for the sampling. Optical densities from each standard image are measured and a table of OD averages of the standard images is established. Radioactivity from another set of standards

[2] I N SITU HYBRIDIZATION TECHNIQUES

35

is determined by counting adjacent cut sections immersed in scintillation fluid in a/3 counter. Counted disintegrations per minute per standard section are converted to total disintegrations (disintegrations per minute corrected for exposure time and fractional volume), which are input into the table. The calibration mode of our image analysis software allow us to perform multiregression analysis on the relation between OD and disintegration data. Calibration curves with the best fits are usually second order or logarithmic fits owing to the limited linear range of film and emulsion, respectively. To avoid time-consuming standard construction for every in situ experiment, the 35S-labeled standards, which have a relatively short half-life, can be crosscalibrated to ~4C plastic standards.

Sampling For analysis, the region of interest is sampled by outlining with a cursor the area of positive hybridization signal. The background signal is subtracted by (a) sampling an area with no signal on the same sections or (b) sampling from the same region on the adjacent section hybridized with the sense strand probe. If a standard curve has been established, OD values are automatically transformed into disintegrations. To express data as "copy numbers detected," disintegrations are converted to curies (Ci) (for 35S, 1/zCi = 2.2 • 106 dpm). Using the specific activity of the probe (Ci/mmol), total curies can be converted to moles and to copy number by multiplying by Avogadro's number (6.023 • 1023 molecules/mol). The measured amounts of mRNA reflect only the number of RNA molecules detected per cell or region, and usually represent an underestimation of the total amount of the examined mRNA present in a cell. Therefore, one should always consider these data as semiquantitative, because the measurements are largely dependent on the hybridization conditions and kinetics used in a particular experiment such as tissue and signal penetration, fidelity of probes, accessibility of the mRNA, fixation, and histological environment.

Localization of mRNAs

Encoding Processing Enzymes

We have applied the in situ hybridization procedures outlined above to studies investigating the tissue-specific distribution of various processing enzymes, with particular focus on their expression in the nervous system and in endocrine tissues. Using specific cRNA probes labeled with [35S]UTP, we were able to identify tissue-specific expression of sets of enzymes, which

36

I MOLECULAR APPROACHES

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F]o. ] Comparative distribution of mRNA transcripts encoding the prohormone conve~ases PC I, PC2, PC5, PACE4, and s in the rat forebrain (approximately !.8 mm caudal ofbregma). Dark-field autoradiograms of coronal sections show clusters of silver grains (white) after hybridization with specific [35S]UTP-labeled riboprobes. Note the region-specific differences in the labeling pattern for each enzyme mRNA,

[2] I N S I T U HYBRIDIZATION TECHNIQUES

37

are probably responsible for the region-specific differences of the posttranslational processing of neuropeptide precursors (6, 10, 34). For example, the growing family of the prohormone convertases is widely distributed throughout the CNS, but its members exhibit distinct individual distribution patterns, (see Fig. 1). Interesting results were obtained from the X-ray analysis of hybridized autoradiograms with various PC probes. Whereas PC1 and PC2 were expressed predominantly in neurons, the distribution of furin and PACE 4 was widespread, but with differences in expression levels in certain areas such as the choroid plexus for furin and the habenulae for PACE4 (Fig. 1). Without sophisticated image analysis these regional differences in mRNA abundance can be evaluated by the human eye, using subjective ratings. The eye is quite reliable in making reproducible subjective rating scales. However, the visual impression can be misleading, if the specific activities of the probes used are not in the same range. For example, the level of furin mRNA is overestimated by visual comparison because the specific activity of the furin probe (600,000 Ci/mmol) used was three times as high as that of the PC 1 and PC2 probes (ca. 250,000 Ci/mmol). Therefore, a semiquantitative approach is always preferable if the number and labeled cells in the sampling region are taken into account. To determine the cellular source the hybridization signal, represented by white clusters of silver grains in the dark-field illumination of the autoradiograms, microscopic analysis of counterstained sections should be performed in bright field, as shown in Fig. 2. Examination of rat brain sections hybridized with cRNA probes specific for PC1, PC2, and furin revealed that in rat cortex PC1 and PC2 probes give a positive hybridization signal over neurons, identified by their bigger and lighter stained nucleus (Fig. 2A and B), in contrast to hybridization with the furin probe, which labeled both neurons and nonneuronal cells, most likely glial cells with smaller and darker stained nuclei (Fig. 2C). Counterstained sections hybridized with a furin sense probe exhibited a random distribution of silver grains without cluster formation, the typical picture of a negative in situ control. Processing enzymes act on a variety of neuropeptides, enzymes, and other substrates. For many questions concerned with substrate specificity of

particularly in the hippocampal formation (HC). The summary diagram depicts subjective rating of relative mRNA abundance in selected areas. Exposure time to Kodak XAR-5 X-ray film was 4 days. ch pl, Choroid plexus; CPU, caudate-putamen; CTX, cortex; Hb, habenulae; HC, hippocampal formation; PVA, paraventricular thalamic nucleus; PVN, paraventricular hypothalamic nucleus; SON, supraoptic nucleus; TH, thalamus; V, third ventricle.

38

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APPROACHES

MOLECULAR

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FIG. 2 Bright-field autoradiograms of prohormone convertase mRNAs in rat cortex counterstained with Cresyl Violet. High-power photomicrograhps of the cerebral cortex show the mRNA expression of PC1 (A), PC2 (B), and furin (C), using insitu hybridization histochemistry with [35S]UTP-labeled riboprobes. The large arrows indicate neurons (larger and lighter stained nucleus) and the small arrows indicate nonneuronal cells (e.g., glia), which have smaller and darker stained nuclei. As a negative control a representative section was hybridized with a sense strand furin cRNA probe (D). The few, randomly distributed grains present background levels. PC 1 and PC2 sense strand probes yielded identical results. Tissue was counterstained with Cresyl Violet (0.5%, v/v). Bars: 10/~m.

enzymes and their possible role in the processing of a specific substrate it is important to identify coexpression patterns of enzymes and substrates at the cellular level. We have performed studies along these lines to examine the cell-specific distribution of prohormone convertases in endocrine cells of the pituitary gland, focusing on the role of PC1 and PC2 in the processing of the opioid precursor POMC in the anterior and intermediate lobe (10). As

[2] I N SITU HYBRIDIZATION TECHNIQUES

39

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FIG. 3 Localization of prohorrnone convertases in the rat pituitary gland. Darkfield rnicrographs were taken from emulsion-coated autoradiograrns hybridized with [35S]UTP-labeled riboprobes specific for PC1 (A), PC2 (B), furin (C), and PC5 (D) rnRNA. Numerous anterior lobe (AL) cells expressed PC1 rnRNA and fewer cells expressed PC2 rnRNA (see text). In the intermediate lobe (IL), all cells expressed high levels of PC2 rnRNA (B) and lower levels of PC1 rnRNA. Furin rnRNA (C) was ubiquitously distributed in all three lobes, including the neural lobe (Ns PC5 mRNA showed the highest expression levels in the AL. Exposure time, 3 weeks. Bar: 100/~rn.

40

I MOLECULAR APPROACHES D-

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FIG. 4 Bright-field photomicrographs depicting mRNA colocalization of POMC and MCH with the prohormone convertases PC1 and PC2 in emulsion-dipped autoradiograms. Sections from rat anterior pituitary (A and B) were simultaneously hybridized with a DIG-labeled POMC cRNA probe and radioactive probes complementary to PC1 mRNA (A) and PC2 (B). (A) Note the dark-stained corticotrophs positive for POMC, showing some grains (PC 1) over their perikarya (arrow) and other hormonecontaining cells expressing PC1 mRNA (arrowhead). (B) Following double hybridization with a PC2 cRNA probe, corticotrophs are labeled only with POMC mRNA (arrowhead) and do not show clusters of silver grains, which appear over other endocrine cells (arrows). (C and D) In the lateral hypothalamus large neurons expressing the MCH gene are visualized with a DIG-labeled riboprobe (courtesy of R. C. Thompson, Ann Arbor, MI), using interference contrast optics. (C) Using doublelabeling in situ hybridization (see text), radioactive labeled PC1 cRNA probes hybrid-

[2] I N S I T U HYBRIDIZATION TECHNIQUES

41

shown in Fig. 3, several mRNAs encoding processing enzymes including PC1, PC2, PC5, and furin are expressed by heterogeneous cell populations in the pituitary gland. In regard to POMC processing initial in situ studies demonstrated a difference in PC 1/PC2 expression ratios in the anterior pituitary in comparison to the intermediate lobe, corresponding to the known differences in POMC processing in these two lobes. PC2 showed a severalfold higher mRNA abundance than PC1 in the intermediate lobe (Fig. 2B). To determine the level of PC1 and PC2 expression in the anterior lobe corticotrophs we carried out double-labeling experiments to detect PC transcripts within the POMC cell population, using a combination of radioactive- and nonradioactive-labeled probes. Colocalization of POMC/PC1 and POMC/ PC2 transcripts, respectively, was examined in anterior lobe corticotrophs using a DIG-labeled POMC cRNA probe (courtesy of J. Eberwine, Philadelphia, PA) and radiolabeled PC cRNA probes (Fig. 4A and B). Whereas PC1 mRNA was expressed in almost all anterior lobe cells including the corticotrophs, as shown by double-labeling in situ hybridization, PC2 mRNA was not detectable in corticotrophs, but was present in anterior lobe cells of different phenotypes (Fig. 4B). This cell-specific expression pattern of PC 1/PC2 is in agreement with the tissue-specific processing pattern of POMC to larger fragments in corticotrophs in comparison to the complete processing in the intermediate lobe due to the high levels of PC2 (9). In the rat hypothalamus, we have demonstrated at the mRNA level the overlapping distribution of PC2 and MCH, which is processed to several different neuropeptides (34). Using the double-labeling hybridization technique (in collaboration with M. Otto and E. Weihe, Mainz, Germany), almost all MCH neurons, identified with a DIG-labeled cRNA probe probe (courtesy of R. C. Thompson, Ann Arbor, MI), expressed PC2 mRNA when cohybridized with a radiolabeled PC2 cRNA probe (Fig. 4B). Cohybridization experiments using PC 1 and MCH riboprobes revealed that PC 1 mRNA is expressed at relatively high levels by a cell population in the lateral hypothalamus, unlike MCH neurons (Fig. 4A). While the involvement of PC1 and PC2 in the processing of MCH awaits in vitro examination, it can be proposed from our in situ hybridization studies that PC2, and not PC1, plays an important role in the posttranslational processing of MCH.

ize to neurons in the lateral hypothalamus (arrowhead), distinct from MCH mRNAexpressing neurons (arrow). (D) In contrast, cohybridization with an [35S]UTP-labeled PC2 cRNA parobe produced double labeling of almost all MCH neurons (arrow). Exposure times, 3 weeks. Bars: (A and B) 10 txm; (C and D) 25 ixm.

42

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Conclusions In situ hybridization techniques have been consistently improved over the

last decade, enabling us to study the expression of low-abundance messages at the cellular level, such as RNA transcripts encoding processing enzymes. With the assistance of polymerase chain reaction (PCR) technology and molecular biological procedures, cRNA probes labeled with radioisotopes have proved to be the most sensitive tools, reaching a detection limit of a few RNA molecules per cell, compromised only by background labeling. Applying this technology to mapping studies of processing enzymes, the determination of region-specific expression of enzyme sets has given us new insights into the tissue-specific differences of posttranslational processing and elucidated the role of some of these enzymes for the processing of specific precursors. We were able to identify prohormone convertases associated predominantly with the secretory pathway of secretion in neurons and endocrine cells, and enzymes ubiquitously expressed. An important lesson we have learned from our studies is that not only the presence or absence of a prohormone convertase but the relative expression ratios are crucial for the tissue- and cell-specific processing. Using nonradioactive hybridization techniques we have shown that these studies can now be carried out at high anatomical resolution at the single-cell level. In the coming years the initial mapping studies will be extended to a quantitative level to determine expression ratios for several enzymes in homogeneous and heterogeneous cell populations in situ and in vitro. Improvement of the sensitivity and quantifiability of the nonradioactive detection techniques will allow simultaneous visualization of multiple messages, providing new data on specific colocalization patterns of enzyme-enzyme and enzyme-substrate combinations. Detailed molecular characterization of the transcriptional regulation of enzyme genes will allow us to address the role of alternative spliced enzyme mRNA forms and to design more specific probes to examine the possible role of tissue-specific splicing events, which is beginning to be studied for the PAM gene. Finally, with novel enzymes being cloned and isolated at an accelerating rate, the histological studies will continue to make important contributions to the characterization of cleavage and substrate specificity of processing enzymes.

Acknowledgments This work was supported by the German Research Foundation (WE 910/2-3) and the Medical Research Council of Canada (MT-11268). R.D. is a scholar of the Fonds de la Recherche en Sant6 du Qu6bec (FRSQ).

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References N. G. Seidah and M. Chr6tien, Trends Endocrinol. Metab. 3, 133 (1992). 2. L. D. Fricker, J. P. Adelman, J. Douglass, R. C. Thompson, R. P. von Strandmann, and J. Hutton, Mol. Endocrinol. 3(4), 666 (1989). B. A. Eipper and R. E. Mains, Annu. Rev. Physiol. 50, 333 (1988). N. G. Seidah, R. Day, M. Marcinkiewicz, S. Benjannet, and M. Chr6tien, Enzyme 45(5-6), 271 (1991). N. G. Seidah, R. Day, J. Hamelin, A. Gaspar, M. W. Collard, and M. Chr6tien, Mol. Endocrinol. 6(10), 1559 (1992). M. K.-H. Sch~ifer, R. Day, W. E. Cullinan, M. Chr6tien, N. G. Seidah, and S. J. Watson, J. Neurosci. 13(3), 1258 (1993). R. Day, M. K.-H. Sch~ifer, W. E. Cullinan, S. J. Watson, M. Chr6tien, and N. G. Seidah, Neurosci. Lett. 149, 27 (1993). N. G. Seidah, R. Day, S. Benjannet, N. Rondeau, A. Boudreault, T. Reudelhuber, M. K.-H. Schfifer, S. J. Watson, and M. Chr6tien, NIDA Res. Monogr. 126, 132 (1992). S. Benjannet, N. Rondeau, R. Day, M. Chr6tien, and N. G. Seidah, Proc. Natl. Acad. Sci. U.S.A. 88(9), 3564 (1991). 10. R. Day, M. K.-H. SchS.fer, S. J. Watson, M. Chr6tien, and N. G. Seidah, Mol. Endocrinol. 6(3), (1992). 11. M. K.-H. Schfifer, J. P. Herman, and S. J. Watson, in "Imaging of Drug Action in the Brain" (E. D. London, ed.), p. 337. CRC Press, Boca Raton, FL, 1993. 12. R. J. Britten and E. H. Davidson, in "Nucleic Acid Hybridization: A Practical Approach" (B. D. Hames and S. J. Higgins, eds.), p. 3. IRL Press, Oxford, 1985. 13. K. L. Valentino, J. H. Eberwine, and J. D. Barchas, "In Situ Hybridization: Applications to Neurobiology." Oxford, New York, 1987. 14. S. J. Watson, P. D. Patel, S. Burke, J. P. Herman, M. K.-H. Schfifer, and S. Kwak, in "In Situ Hybridization and Related Techniques to Study Cell-Specific Gene Expression in the Nervous System (Society for Neuroscience Short Course Syllabus)" (J. L. Roberts, ed.), p. 4. Society for Neuroscience, Washington, DC, 1988. 15. W. S. Young, III, in "Handbook of Chemical Neuroanatomy" (A. Bj6rklund et al., eds.), Vol. 8, p. 481. Elsevier, Amsterdam, 1990. 16. M. W. MacCumber, S. H. Snyder, and C. A. Ross, J. Neurosci. 10(8), 2850 (1990). 17. W. S. I. Young, J. Chem. Anat. 1, 177 (1988). 18. J. Sambrook, E. F. Fritsch, and T. Maniatis, "Molecular Cloning: A Laboratory Manual." Cold Spring Harbor Lab., Cold Spring Harbor, NY, 1989. 19. D. A. Melton, P. A. Krieg, M. R. Rebagliati, T. Maniatis, K. Zinn, and M. R. Green, Nucleic Acids Res. 12, 7035 (1984). 20. E. T. Schenborn and R. C. Mierendorf, Jr., Nucleic Acids Res. 13, 6223 (1985). 21. J. E. Springer, E. Robbins, B. J. Gwag, M. E. Lewis, and F. Baldino, Jr., J. Histochem. Cytochem. 39, 231 (1991). 22. K. H. Cox, D. V. DeLeon, L. M. Angerer, and R. C. Angerer, Dev. Biol. 101, 485 (1984). o

.

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I MOLECULARAPPROACHES 23. Y. Misumi, Y. Sohda, and Y. Ihehara, Nucleic Acids Res. 18, 22 (1990). 24. M. K.-H. Schafer, D. A. Stoffers, B. A. Eipper, and S. J. Watson, I. Neurosci. 12(1), 222 (1992). 25. B. A. Eipper, L. P. Park, I. M. Dickerson, H. T. Keutmann, E. A. Thiele, H. Rodriguez, P. R. Schofield, and R. E. Mains, Mol. Endocrinol. 1(11), 777 (1987). 26. M. E. Lewis, R. G. Krause, II, and J. M. Roberts-Lewis, Synapse 2, 308 (1988). 27. M. E. Lewis, T. G. Sherman, and S. J. Watson, Peptides (N. Y.) 6, 75 (1985). 28. R. H. Singer, J. B. Lawrence, and C. Villnave, Biotechniques 4(3), 230 (1986). 29. L. M. Angerer, M. H. Stoler, and R. C. Angerer, in "In Situ Hybridization: Applications to Neurobiology" (K. L. Valentino, J. H. Eberwine, and J. D. Barchas, eds.), p. 42. Oxford Univ. Press, New York, 1987. 30. R. J. Britten, D. E. Graham, and B. R. Neufeld, in "Methods in Enzymology" (L. Grossman and K. Moldave, eds.), Vol. 29E, p. 363. Academic Press, New York, 1974. 31. J. Casey and N. Davidson, Nucleic Acids Res. 4, 1539 (1977). 32. F. Baldino, Jr. and M. E. Lewis, Methods Neurosci. 1, 282 (1989). 33. J. T. McCabe, R. A. DeSharnais, and D. W. Pfaff, in "Methods in Enzymology" (P. M. Conn, ed.), Vol. 168, p. 823. Academic Press, San Diego, 1989. 34. W. E. Cullinan, N. C. Day, M. K.-H. Sch/ifer, R. Day, N. G. Seidah, M. Chr6tien, H. Akil, and S. J. Watson, Enzyme 45(5-6), 285 (1991).

[3]

Analysis of Ontogeny of Processing Enzyme Gene Expression and Regulation Min Zheng and John E. Pintar

Introduction Studies of proteolytic processing enzymes, especially endoproteases, have long been hampered by their presumed low abundance in vivo. Indeed, it was once estimated that to complete proteolytic processing of 1000 precursor molecules in an hour, a secretory granule would need only a single copy of an endoprotease with a turnover number of 50 precursors per minute (1). As a result, our knowledge has been confined to determining the enzymatic properties of potential proteases and the cleavage sites present on proprotein substrates, with little known about the ontogeny and regulation of the proteolytic processing enzymes. As detailed by other contributions in this volume, the cloning of processing enzyme genes has allowed rapid progress to be made in defining their structure and function. Furthermore, the availability of identified cDNA sequences has made it possible to apply sensitive methods such as in situ hybridization and RNase protection assays to study their developmental expression. In situ hybridization identifies gene expression within a spatial context, in contrast to other molecular and biochemical methods that usually require tissue isolation and homogenization. This is particularly advantageous to the study of gene expression in development because heterogeneous tissues with changing properties are compacted in miniature. Because the processing enzymes had been expected to be expressed (and are expressed) at lower levels compared to their substrates, characterization of their expression requires methods sensitive enough to detect a few copies of mRNA per cell. In situ hybridization, if performed properly, can achieve such sensitivity (2). On the other hand, once the spatial and temporal distributions of the processing enzymes are defined, a more quantitative measure of the precise level of gene expression is a prerequisite for the investigation of regulation of gene expression; this is now afforded by RNase protection assays. To dissect the molecular mechanisms of gene expression, an in vitro system is usually employed, which includes isolation and propagation of a particular tissue or cell type of interest. Introduction of a cell culture system allows the precise control of the extracellular environment. Various physiologically relevant exogenous factors can be applied and the response of target gene Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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expression assessed. This approach, combined with the study of proprotein gene expression, helps to evaluate the distinctive contribution of different processing enzymes in the maturation of proproteins and neuropeptide precursors in development. This chapter focuses on techniques in the analysis of processing enzyme gene expression and regulation in development. In situ hybridization is introduced first. Emphasis has been placed on tissue and probe preparation for in situ hybridization, as well as application of colocalization of processing enzymes with their potential substrates. The second part discusses the methods pertaining to the regulation of gene expression. This includes the coupling of a primary culture system with the RNase protection assay, using prohormone convertase gene expression in postnatal pituitary as a model system. Practical applications of these techniques as well as the analysis of typical results are given following the introduction of each method.

In Situ H y b r i d i z a t i o n In situ hybridization takes advantage of the sequence-specific hybridization of applied complementary nucleotide probes to endogenous nucleic acids preserved in tissue sections. Strict complementarity of base pairing (hydrogen bonding) between the probe and immobilized target nucleic acid together with appropriate hybridization and posthybridization washes ensure a high specificity of the signals obtained. Spatial positions of target nucleic acids are subsequently visualized by various mechanisms depending on the nature of probe labeling. The in situ hybridization method was first developed in the late 1960s for detecting amplified ribosomal DNA sequences in cell nuclei (3, 4), but more recently has been most widely used to detect the presence of specific mRNA species, which indicates the distinct nature and functional state of different cell types. In applying this technique to study gene expression in embryogenesis, sequential stages of embryos are isolated and tissue sections are prepared. Hybridization of these sections with nucleotide probes thus allows the spatial distribution of cell types containing mRNA species of interest to be determined, as well as their temporal changes in development.

Experimental Animals All studies are conducted in accordance with the principles and procedures outlined in the NIH Guide for the Care and Use of Laboratory Animals. For in situ hybridization, timed pregnant Sprague-Dawley rats (Carom, Inc., Wayne, NJ) are sacrificed by decapitation. For embryos younger than embry-

[3]

ONTOGENY OF PROCESSING ENZYME EXPRESSION

47

onic day 9 (e9) the whole uteri are fixed by immersion in 4% (w/v) paraformaldehyde prepared in phosphate-buffered saline (PBS) for about 4-6 hr and equilibrated in a 20% (w/v) sucrose solution. The tissues are subsequently embedded in OCT compound (Miles, Inc., Elkhart, IL) for cryostat sectioning. Embryos older than e9 and postnatal rat pituitaries are quickly dissected and are either fixed prior to embedding (e9-e 12 embryos) or freshly frozen embedded in OCT compound (over el2). When possible, embryos are staged according to the morphology of their limb buds in reference to criteria established by Wanek et al. (5; see below for details).

Tissue Preparation Two procedures are used to ensure adherence of tissue sections to the slides. In the first, the slides are pretreated sequentially with gelatin and poly-Llysine. Slides are washed in 0.2 M HC1 and rinsed with acetone. Slides are subbed in 0.15% (w/v) gelatin for 5 min, followed by an overnight drying at 50~ Slides are then immersed in 20% (w/v) poly-IAysine (Mr >300,000, Sigma, St. Louis, MO) for 10 min, followed by additional overnight drying at 50~ A simplified slide subbing procedure has been used, in which both treatments with gelatin and poly-IAysine are replaced by a 30-sec dip in a siliconizing silane solution [2% (v/v) 7-aminopropyltriethoxysilane prepared in acetone; Pierce, Rockford, IL). This is followed by two additional rinses in acetone and one rinse in diethylpyrocarbonate-treated H20 (DEPC-H20). Thus, the whole process of treating a batch of slides can be completed within 1 hr, with the slides subsequently allowed to dry at room temperature overnight. In our experience, slides coated with silane offer equivalent efficiency of tissue adhesion as gelatin and poly-IAysine subbing. Coated slides can be stored for several months in a dust-free environment. Both sagittal and transverse sections (8-12 ~m thick) of the embryonic tissues are prepared in a cryostat (Hacker Instruments, Inc., Fairfield, NJ) for in situ hybridization experiments. Tissue sections are thaw mounted on slides and stored at -80~ until use.

Preparation o f Probes Both sense and antisense [35S]UTP-labeled cRNAs are prepared using an in vitro transcription system (Promega, Madison, WI). The transcription mixture in a total volume of 10/xl is assembled at room temperature, containing the transcription buffer [40 mM Tris (pH 7.5), 6 mM MgC12,2 mM spermidine, and 10 mM NaC1], 10 mM dithiothreitol (DTT), 40 units of ribonuclease

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inhibitor (RNasin), 160/zCi of speed-vacuum dried [35S]UTP (specific activity, 1000-1500 Ci/mmol; DuPont-New England Nuclear, Boston, MA), 0.5 mM each of unlabeled nucleotide triphosphates (ATP, GTP, and CTP), 1/zg of linearized DNA template, and 20 units of bacteriophage RNA polymerase. The transcription reaction is allowed to proceed at 37~ for 60 min. The DNA template is removed by digestion with 1 unit of RNase-free DNase at 37~ for 20 min. Size-exclusion chromatography is used to remove unincorporated nucleotides, by passing probes through a Sephadex G-50 RNase-free spin column (Boehringer-Mannheim, Indianapolis, IN) after phenol-chloroform extraction. The specific activity of recovered cRNA is measured by scintillation counting. Riboprobes are aliquoted and stored at -80~ The sense probes are used as a negative control.

In Situ Hybridization Protocol In situ hybridization experiments are performed following protocols previously described (6), with certain modifications. The following is a summary of the procedure currently in use. Tissue slides are retrieved from the -80~ freezer and fixed in 4% (w/v) paraformaldehyde at room temperature for 10 min. The slides are rinsed in DEPC-H20 for 5 min, dehydrated in ethanol, and air dried for 10 min. Slides are subsequently rehydrated and treated with acetic anhydride to suppress nonspecific electrostatic binding of probes to the tissue and slide coating (7). In this procedure, after a brief pretreatment of slides with 0.1 M triethanolamine-0.05 M acetic acid, slides are rinsed vigorously in freshly prepared 0.2% (v/v) acetic anhydride diluted with triethanolamine. After a further 10-min incubation, slides are rinsed in 0.2x SSC (1.0• SSC is 0.15 M NaC1 plus 0.015 M sodium citrate), dehydrated, and air dried. To be able to apply different probes to multiple tissue sections on a slide and help to retain the applied solution via capillary tension, a rubber cement ring is carefully applied, circling each section. The prehybridization is performed at room temperature for 1-3 hr in a buffer containing 50% (v/v) deionized formamide (Boehringer-Mannheim), 10 mM Tris (pH 7.5), 1 mM ethylenediaminetetraacetic acid (EDTA), 600 mM NaC1, Denhardt's solution [0.02% (w/v) Ficoll, 0.02% (w/v) polyvinylpyrrolidone, and 0.02% (w/v) bovine serum albumin], and heat-denatured heterologous nucleic acids [0.05% (w/v) yeast total RNA, 0.005% (w/v) yeast tRNA, and 0.05% (w/v) salmon sperm DNA]. The riboprobes are heated at 85~ for 5 min and then chilled on ice (to disrupt the possible formation of RNA secondary structure). These are diluted to an activity of---4 x 10 7 dpm/ml (confirmed by scintillation counting) in the hybridization buffer. The hybridization buffer is similar to the buffer used for prehybridization, except that 10% (w/v) dextran sulfate

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(Mr 8000; Sigma), 10 mM DTT, and 0.1% (w/v) sodium dodecyl sulfate (SDS) are added. Prehybridization solution is replaced by 25-60/~1 of the hybridization buffer, depending on the size of the tissue section. The slides are placed in a moist chamber and hybridization is carried at 50~ for 12 hr (overnight). Slides are then washed at 50~ for 30 min in a solution containing 50% (v/v) deionized formamide, 1 x SSC, and 10 mM DTT. This is followed by a rinse in 0.5 x SSC and treatment at room temperature for 30 min with RNase A [0.1 mg/ml, diluted in 10 mM Tris (pH 8), 1 mM EDTA, 500 mM NaC1] to degrade nonhybridizing single-stranded probes (8). The final wash is performed with 0.2x SSC at 50-60~ for 2 hr with gentle stirring. Slides are dehydrated in an ethanol series and subjected to X-ray autoradiography. This is followed by nuclear emulsion autoradiography for higher resolution of signal detection. Slides are coated with a 1:1 (v/v) dilution of Kodak NTB2 emulsion (Eastman Kodak, Rochester, NY) and exposed at 4~ in the dark. Usually at least a threefold longer time of exposure for emulsioncoated slides is needed compared to a sufficient X-ray film exposure. Finally, dipped slides are developed with Kodak D19 developer and stained with hematoxylin-eosin (HE). The slides are examined in Wild and Leitz microscopes (Leica, Heerbrugg, Switzerland) under bright-field, dark-field, and epipolarized illumination.

Discussion o f Methodology Rat embryonic development proceeds relatively rapidly. Both tissue morphology and the functional status of specific cells change significantly during organogenesis. The precise determination of developmental stage of embryo, therefore, is of paramount importance in the study of expression of genes involved in developmental processes. Although the experimental animals used to obtain embryos are "timed-pregnant" and gestational stages inferred from observed copulation plug date, sometimes embryos are obtained that are different from their expected age because plugs were missed. Furthermore, because of differences in fertilization time of individual oocytes, individual embryos within the same litter are at different developmental stages and indeed can differ in developmental age by as much as 24 hr. In establishing a more precise staging system, we have adopted a procedure of Wanek et al. (5), which is based on evaluating the gross external morphology of the mouse limb at different developmental stages. Corresponding ages for rat embryos of equivalent age are deduced from the listed mouse stages by adding 1-1 89days. The rat embryo is rapidly isolated from the uterus and the morphological characteristics of both forelimb and hindlimb are evaluated under a dissecting microscope (with the hindlimb usually 1 day behind the forelimb in development). If necessary, embryonic limbs are dissected and

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placed in a petri dish containing PBS solution for better viewing. Gross morphology of the limbs is compared to a chart detailing the morphological characteristics of the limb at various developmental stages and the appropriate gestational stage is assigned to the embryo. Sophisticated stereotypic development of the limb ensures that this staging system can effectively encompass a large developmental window (e 10-e ! 8). The application of limb bud staging system ensures a consistency in determining rapidly the precise age of each embryo. In general, e12-e18 embryos are embedded in OCT compound (directly fresh frozen). We have found that sections derived from fresh-frozen embryos consistently give a higher signal-to-noise ratio than those from fixed embryos. This is particularly advantageous in the detection of low-abundance transcripts such as processing enzyme mRNAs and has therefore been used in most experiments reported here. In contrast, prefixed tissues compromise the signal labeling intensity for better morphology. Fixation of the embryo helps to identify regions and cell types of gene expression, such as those of early postimplantation staged embryos, with greater precision and often in conjunction with immunocytochemistry. We have, therefore, used paraformaldehyde prefixation only for embryos younger than el2, although freshfrozen sections have also been prepared. Owing to the small size of the uterus in these stages and the accessibility of its intrauterine cavity when decidua are isolated individually, intracardial perfusion of adult rat is not necessary. Although cDNA and oligonucleotide probes have been used, the most common probes for in situ hybridization remain radiolabeled cRNAs (riboprobes), which can readily be labeled to high specific activities. To ensure success and consistency during in situ hybridization experiments, the quality of newly synthesized probes should always be evaluated via gel electrophoresis prior to use. Probes of high quality usually result after an efficient transcription reaction (over 108 dpm/labeling reaction, using 1 /zg of plasmid template) and appear as a major band of appropriate size when electrophoresed on a denaturing polyacrylamide gel. Probes of relatively short length (riboprobes varying in length from 320 to 372 bp are used in the present study), can be used directly for hybridization experiments. Substantially longer probes should be shortened after synthesis by alkaline hydrolysis to allow their efficient penetration into the tissue sections (2). The extent of hydrolysis should be carefully controlled and the average length of probe monitored by electrophoresis. Alternatively, sections can be pretreated with proteinase to facilitate probe penetration. Although the possibility of RNase contamination may exist, inefficient transcription is usually due to the precipitation of DNA template by the spermidine present in the stock concentration of transcription buffer (9). To avoid this problem, all the components of the

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reaction may be added separately to the side wall of an Eppendorf tube and mixed by a brief spin on a microcentrifuge. Additional pipetting of the mixture is recommended to ensure thorough dispersion of all the components, especially the bacteriophage polymerase. To compare informatively the relative abundance of different processing enzyme gene mRNAs in development, probes of similar length and specific activity should be used. Colocalization of a processing enzyme with a proprotein is indicative of a possible enzyme-substrate relationship, whereas the coexistence of distinctive types of processing enzymes suggests their possible collaboration in the completion of posttranslational modification processes. Although such information can sometimes be deduced by comparing the expression of different genes on serial sections, unequivocal demonstration of coexpression of two genes requires their covisualization on a single tissues section. One way to achieve this goal is to apply two different cRNA probes to the same section, which are subsequently distinguished by virtue of the distinct nature of probe labeling. Besides the commonly used [35S]UTP-labeled probes, we have also used digoxigenin-UTP to synthesize cRNA probes, using commercially available reagents (Genius RNA labeling kit; Boehringer Mannheim). Digoxigenin-UTP-labeled probes are subsequently detected on the tissue sections by an enzyme immunoassay. Owing to the similar nature of nucleotide hybridization with both kinds of cRNA probes, we have been able to apply both probes simultaneously, with minimum change in the basic in situ hybridization protocol. In our experience, the hybrids formed by the [35S]UTP-labeled probes are stable during the subsequent color visualization of digoxigenin-UTP-labeled probes. Prehybridization is performed in a buffer containing 50% (v/v) deionized formamide, 5 • SSC, Denhardt's solution, 0.025% (w/v) yeast tRNA, 10% (w/v) dextran sulfate, 0.01% (w/v) Nlauroylsarcosine, and 0.02% (w/v) SDS. The hybridization buffer is similar to the prehybridization buffer, except for the addition of 10 mM DTT. Digoxigenin-UTP-labeled probe is empirically diluted to the final concentration of 1-100 pg/ml in the hybridization buffer, depending on the abundance of the transcripts to be detected; higher transcript concentrations, although required for the detection of low-abundance mRNA, can also result in higher nonspecific background. After hybridization, slides are washed under identical stringency conditions and digested with RNase A as described above. Slides are then treated with nonfat dry milk (1 mg/ml), which is dissolved in a buffer containing 100 mM Tris (pH 7.5) and 150 mM NaCI (buffer I). This is followed by incubation with a 1 : 1000 (v/v) dilution in buffer I of alkaline phosphatase-conjugated anti-digoxigenin antibody (Fab fragment; Boehringer Mannheim) at room temperature for 1-3 hr. If the signals are low, lengthening the incubation time usually enhances the antibody binding and should be performed at 4~ Slides are then washed in the buffer I for

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10 min twice and equilibrated with buffer II [100 mM Tris (pH 9.5), 100 mM NaC1, and 50 mM MgCI2]. Digoxigenin-UTP-labeled probe is visualized by a 2-hr to overnight incubation of slides in chromogen solution in the dark at room temperature. The chromogen solution is freshly prepared in buffer II and consists of 150 ~g of nitroblue tetrazolium [NBT, stock solution 75 mg/ ml, prepared in 70% (v/v) dimethylformamide] per milliliter, 100/~g of 5bromo-4-chloro-3-indolylphosphate (X-phosphate; stock solution 50 mg/ml, prepared in straight dimethylformamide) per milliliter, and 25 ~g of L[- ]-2,3,5,6-tetrahydro-6-phenylimidazo[2,1-b]thiazole (levamisole; Sigma) per milliliter. The color reaction is terminated by incubating slides in a solution consisting of 10 mM Tris (pH 8), 1 mM EDTA, and 100 mM NaC1. Slides are then dehydrated in an ethanol series. After exposure to X-ray film, slides are emulsion dipped in Ilford L4 emulsion (Polysciences, Inc., Warrington, PA), developed, and mounted in Permount without counterstain. Digoxigenin-UTP labeling usually appears as bluish deposits, which form high contrast to the silver grains derived from [35S]UTP-labeled probes on the same section (see Fig. 2c and d). Besides its greater speed compared to using conventional radiolabeled probes (lengthy emulsion dipping is not needed), digoxigenin-UTP labeling in situ hybridization offers satisfactory sensitivity in mapping gene expression in development. Using the same preparation of digoxigenin-UTP-labeled proopiomelanocortin (POMC) probe as well as [35S]UTP-labeled POMC probe, for instance, we have obtained comparable results in detecting prenatal POMC gene expression just following the closure of Rathke's pouch. In addition, there is an increasing number of reports that have used "whole-mount in situ hybridization" to detect specific mRNAs in vertebrate embryos at early postimplantation ages (10).

Examples Expression of Furin and Carboxypeptidase E in Early Rat Embryogenesis Several mammalian endoproteases have been successfully identified, including furin, PC1, PC2, PC4, and PACE4 (11, 12). Among these, furin has been shown to cleave preferentially those proproteins maturing from the constitutive secretory pathway. The proteolytic cleavage of proproteins by furin exposes the dibasic residues at the C termini, which are usually absent from the mature protein and thus are removed by subsequent exoproteolytic cleavage. The only known enzyme capable of performing such a trimming reaction is carboxypeptidase E [CPE (13)]. It is thus possible that furin and CPE may functionally collaborate to complete the proteolytic processing of common proproteins in embryogenesis and, if so, would be expected to be

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expressed in both a spatially and temporally correlated manner. To compare the expression domain of these two genes in development, we have performed in situ hybridization experiments on serial transverse sections of el2 rat embryo, with furin and CPE probes applied on adjacent sections. The results are shown in Fig. 1 (14). Positive labeling appears as a bright region owing to the reflection of autoradiographic grains under dark-field illumination. At this stage, in agreement with its general involvement of constitutive proprotein processing, furin mRNA is expressed at low but detectable levels in nearly all tissues and structures throughout the embryo. Severalfold higherthan-basal levels of expression, however, are observed in heart and liver primordium, as well as in the body wall surrounding the umbilical vein (Fig. l c, f, and i). In contrast, the developing nervous system noticeably lacks detectable levels of furin gene expression. In comparison, CPE mRNA is prominently expressed throughout the mantle layer of the neural tube, as well as in cranial and peripheral ganglia. In addition, a significant level of CPE expression is also observed in the embryonic heart, in the epithelium of the mandibular component of first bronchial arch, and the mesentery surrounding the gut (Fig. l d, g, and j). Although the expression domains of both genes overlap, with the embryonic heart being a clear example, several regions where furin is expressed at high levels lack CPE expression, such as the body wall surrounding the umbilical vein (compared Fig. li and j) and the liver primordium (data not shown). Applying both furin and CPE probes in parallel on adjacent tissue sections has thus enabled direct comparison of their spatial expression at this stage. Careful comparison of their expression on serial transverse sections shows that the expressions of furin and CPE only partially overlap. This indicates that if furin is functional at this stage, additional types of carboxypeptidase(s) may be recruited for exoproteolytic processing in such regions as liver primordium. The molecular identity of such a processing enzyme remains to be elucidated.

PC2 Mediation of Developmentally Regulated Proopiomelanocortin Proteolytic Processing in Anterior Rat Pituitary In the rat pituitary, the polyprotein precursor proopiomelanocortin (POMC) is synthesized by all parenchymal cells in the intermediate lobe (IL) and by 5-10% of cells in the anterior lobe (AL) [(15), see also Fig. 2a]. Distinctive posttranslational modifications, however, occur in these two cell populations. Proopiomelanocortin undergoes limited proteolytic cleavages in the AL, generating mainly adrenocorticotropic hormone (ACTH) and/3-1ipotropin (/3-LPH). In contrast, further cleavages inside both ACTH and/3-LPH occur in the IL, thus generating smaller peptide products such as c~-melanocytestimulating hormone (c~-MSH), y-LPH, and/3-endorphin (/3-EP) (15). PC1 and PC2 have been shown to be expressed distinctly in pituitary: specifically,

Hb

b, c, d

a

b

d

C

e

. w

Uv

h

.j

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PC1 is expressed in both the AL and IL, whereas PC2 is mainly expressed in the IL [(16), see also Fig. 2b]. These patterns, when correlated with results of gene transfer experiments (17, 18), indicate that the distinct expression of PC1 and PC2 is responsible for the observed lobe-specific proteolytic processing of POMC. Therefore, POMC is processed to a limited extent by PC 1 in the AL, whereas collaboration of both PC 1 and PC2 in the IL results in more extensive POMC processing. Interestingly, fetal and early postnatal AL have been shown to process POMC more extensively, generating a peptide profile similar to that of the IL (19, 20). This developmental plasticity could be mediated by the presence of additional proteinaceous modulators, alterations of the processing microenvironment, or regulation of the expression of distinct processing enzymes. To address this question, we examined the expression of PC1 and PC2 on serial ages of postnatal pituitary. In contrast to the findings in adult, the AL contains significant levels of PC2 mRNA during these stages, reaching a peak at postnatal day 15 (p15) (Fig. 2d). In situ hybridization was performed with digoxigenin-UTP-labeled POMC probe and [35S]UTP-labeled PC2 probe on the same pituitary section, using protocols described above. This allows covisualization of both probes. The POMC transcripts are revealed as areas of blue deposits. The cytoplasm of POMC-expressing cells in both the IL and AL are clearly delineated, whereas the nuclear regions are largely devoid of labeling (Fig: 2c). This indicates that the majority of the POMC transcripts are processed to their mature form and exported out of the nuclei. The extremely fine deposit of nonradioactive labeling products allows the resolution of this method to rival that of immunocytochemistry. [35]UTP-labeled PC2 hybrids appear as autoradiographic grains. Aggregates of the grains are scattered in both the AL and IL, indicating that PC2 is expressed in both lobes at this stage (Fig. 2d). The majority of the POMC-expressing cells in the IL and some of the cells in the AL

FIG. 1 Comparison of furin and CPE gene expression in the 12-day rat embryo. (a) Line drawing of an e 12 rat embryo, indicating the approximate planes of section. Bright-field (b, e, and h) and dark-field (c, d, f, g, i, and j) micrographs of corresponding adjacent sections are shown. Adjacent sections are hybridized with furin cRNA (c, f, and i) and CPE cRNA (d, g, and j), thus allowing the comparison of their spatial patterns of expression. Furin is expressed at a low level in most tissues, but is undetectable in the neural tube. In contrast, CPE is expressed in the newly differentiated neurons in the diencephalon (Di), hindbrain (Hb), trigeminal ganglia (Tg), and spinal cord (Sc). Both furin and CPE are expressed in embryonic heart (H). Note that furin is expressed at a significant level in the body wall surrounding the umbilical vein (Uv), whereas CPE is expressed at the adjacent mesentery (Ms) surrounding the midgut. Exposure time for autoradiography is 6 weeks. Magnification: x8.5.

i~~,~ ~I ', ~i........... ~,

b

Ir

~

jr

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possess both types of labeling products, indicating that POMC and PC2 transcripts can coexist. This is in contrast to the lack of apparent PC2 expression in the AL in adult pituitary. This suggests that enhanced expression of PC2 in AL POMC-expressing cells may be responsible for the developmental switch in the POMC proteolytic processing pattern.

RNase Protection Assay The RNase protection assay (solution hybridization, RNase mapping) detects the presence and abundance of mRNA transcripts by virtue of hybridization with the complementary ribonucleotide probes in an aqueous phase. The probes annealing to the homologous sequence on the transcripts are protected from subsequent ribonuclease digestion and are detected by gel electrophoresis. This technique was pioneered in the early 1980s by Zinn et al. (21) and has been refined for studying the neuroendocrine systems (22). Over the years the RNase protection assay has been perfected to be one of the most sensitive methods in evaluating levels of gene expression. In the following sections we describe the methods utilizing the RNase protection assay in combination with a short-term tissue culture system, which can be used to study the regulation of processing enzyme gene expression in rat pituitary development.

Fit:;. 2 Comparison of POMC and PC2 expression in adult pituitary (a and b) and postnatal day 15 (p15) pituitary (c and d). Adjacent frontal sections of adult pituitary are hybridized with POMC (a) and PC2 (b), respectively, using [35S]UTP-labeled riboprobes. Micrographs are taken under dark-field illumination. Note that the anterior lobe (AL) expresses only a low level of PC2 compared to the intermediate lobe (IL), whereas the posterior lobe (PL) does not express PC2. Exposure times are 4 days (a) and 6 weeks (b). Magnification: x8.4. (c and d) Colocalization ofdigoxigenin-UTPlabeled POMC transcripts with [35S]UTP-labeled PC2 transcripts in the p l5 rat pituitary. POMC-expressing cells are outlined by areas of chromagen deposits in the cytoplasm [(c), bright-field illumination; section without counterstain]. Nearly all cells in the IL and scattered cells in the AL express POMC. In (d), the same section has been photographed under dark-field illumination and PC2-expressing cells are identified by aggregates of fine autoradiographic grains. Note that a considerably higher level of PC2 is expressed in the AL at this stage compared to that in adult [compare (b) and (d)]. Some aggregates of autoradiographic grains in both the IL and AL overlap with the chromagen deposits (arrows), indicating coexpression of PC2 with POMC. Note that there is some chromagen interference with optimal photography of autoradiograph grains. The slide was incubated for 3 hr in the chromogen reaction, followed by 6 weeks of autoradiographic exposure. Magnification: x 35.

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Primary Tissue Culture Rat pituitaries of various postnatal ages are rapidly isolated, and the anterior lobe (AL) and neurointermediate lobe (NIL, intermediate lobe and posterior lobe) of each are separated under a dissecting microscope. Six ALs or NILs are pooled per sample and collected in 1 ml of ice-cold medium containing Dulbecco's modified Eagles' medium (D-MEM; GIBCO-BRL, Gaithersburg, MD), 1% (v/v) heat-inactivated fetal bovine serum, and kanamycin (50/zg/ ml). Each sample is washed in 1 ml of this medium for 5 min and transferred to 24-well plates (16-mm well diameter) containing 0.5 ml of medium with or without a desired regulatory factor. Samples are incubated at 37~ for 1-24 hr in a chamber supplied with a 95% air/5% CO2 mixture.

RNA Isolation Pituitary samples either freshly isolated or harvested from the primary culture are collected and homogenized in 200/zl of ice-cold buffer containing 10 mM Tris (pH 8), 3 mM CaC12, 2 mM MgC12, 0.15% (v/v) Triton X-100, and 0.3 M sucrose. Homogenization is carried out in a Dounce (Wheaton, Millville, NJ) all-glass homogenizer. Typically 25 strokes are sufficient to disrupt the cell membrane integrity. Homogenates are then layered onto a 300-/zl cushion of the same buffer, except that 0.3 M sucrose is replaced by 0.4 M sucrose. Intact nuclei are separated from the cytoplasmic fraction by centrifugation at 1000 g for 10 min at 4~ Cytoplasmic RNA is purified from the supernatant by digestion with proteinase K (100/zg/ml) at 42~ for 1 hr in the presence of 0.1 vol of digestion buffer, which contains 10 mM Tris (pH 8), 5 mM EDTA, and 1% (w/v) SDS. This is followed by phenol-chloroform extraction and 2-propanol precipitation. Pelleted cell nuclei are resuspended in 60/xl of buffer containing 50 mM Tris (pH 8), 0.1 mM EDTA, 5 mM MgC12, and 40% (v/v) glycerol. Nuclear samples are then treated with 30 units of RNasefree DNase, followed by digestion with proteinase K (100/xg/ml) at 42~ for 30 min in the presence of digestion buffer. Nuclear RNA is purified by phenol-chloroform extraction and ethanol precipitation. Both nuclear and cytoplasmic RNA are resuspended in 5-10/xl of TE [10 mM Tris (pH 7.6), 1 mM EDTA] and the combined total RNA is measured by ultraviolet (UV) absorbance at 260 nm.

Preparation of Probes [a-32p]UTP-labeled runoff transcripts are synthesized from cDNA templates, using an in vitro transcription system (Promega). The plasmid clone is linear-

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ized inside the cDNA template at a desired site downstream of the bacteriophage promoter. Transcription is initiated by adding the appropriate bacteriophage RNA polymerase and terminated by the removal of DNA templates by adding 1 unit of RNase-free DNase. Twenty micrograms of carrier tRNA is added and probes are extracted with phenol-chloroform. To remove incomplete runoff transcripts that may contribute to smearing background in the hybridization, riboprobes are electrophoresed on a 5% (w/v) denaturing polyacrylamide gel. Gel strips corresponding to the full-length probes are excised and eluted in a buffer containing 10 mM Tris (pH 7.5), 1 mM EDTA, 0.1% (w/v) SDS, and 500 mM ammonium acetate. The labeled RNA is isolated by precipitation with 2-propanol in the presence of 2.5 M ammonium acetate and resuspended at saturating levels in the hybridization buffer (see below) to a final concentration of 2 x 10 7 dpm/ml (for detecting abundant transcripts, such as POMC) or 5 x 10 6 dpm/ml (for detecting processing enzymes).

RNase Protection Assay Protocol The RNase protection assay protocol is based on those previously reported (22-25). The hybridization mixture in the volume of 30/zl is freshly prepared, which contains 80% deionized formamide, 40 mM piperazine-N,N'-bis(2ethanesulfonic acid) (PIPES; Sigma), 0.4 M NaC1, and 1 mM EDTA. Five microliters of each RNA sample (1.5-4/zg, diluted in TE) and diluted probes (5-20 x 106 dpm/ml) are added. The hybridization mixture is heat denatured at 85~ for 5 min and hybridization is allowed to proceed at 45~ for 12 hr (overnight). To remove single-stranded probes, samples are digested with RNase A (40/zg/ml) and RNase T1 (2 /xg/ml) at 30~ for 1 hr in a buffer containing 100 mM Tris (pH 8), 5 mM EDTA, and 300 mM NaC1. The digestion is terminated by adding 20/xl of 10% (w/v) SDS, 20/xg of proteinase K, and incubating at 37~ for 20 min. Samples are phenol-chloroform extracted and ethanol precipitated, with 5 /xg of carrier tRNA added. The pellet is resuspended in 10/xl of gel loading buffer [TE containing 1% (w/v) bromphenol blue and 0.1% (w/v) xylene cyanol], heated at 70~ for 5 min, and electrophoresed on a 5% (w/v) nondenaturing polyacrylamide gel. The gel is heat dried under vacuum and exposed to Kodak X-Omat film (Eastman Kodak) with a calcium tungstate intensifying screen at -80~ for 2 hr to 3 days, depending on the abundance of the transcripts to be detected and level of background. The gel strips corresponding to protected bands by different probes are excised and radioactivity measured by liquid scintillation counting.

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Data Analysis A cRNA probe for the ubiquitously expressed gene encoding cyclophilin has been included in all experiments performed. Cyclophilin gene expression has been shown to be inert to a variety of exogenous treatments (26). Since small variations of total tissue RNA applied in each reaction usually occur, the level of cyclophilin RNA is assayed simultaneously to serve as an internal standard, reflecting the actual amount of total RNA in each sample. The level of t h e expression of gene of interest can then be normalized to the amount of cyclophilin RNA. Results obtained from a treated group are then converted to the percentage of those of a control group. These are subject to one-way analysis of variance, followed by a Student's t test to evaluate the difference in processing enzyme RNA level between treated and control untreated groups.

Discussion o f Methodology Preparation of probes for RNase protection assay is essentially the same as for in situ hybridization, except that [a-32p]UTP is used. A specific activity of over 1 x 10 9 dpm//~g of RNA probes, representing incorporation of over 60%, is typically achieved in a single reaction. In linearizing the DNA template for synthesizing the runoff transcripts, restriction enzymes generating a 5' overhang or blunt end should be used. This can avoid possible synthesis of sense strand sequences, which can initiate at a 3' overhang sequence (27). In our procedure, however, the choice of restriction enzyme for DNA template linearization is not necessarily restricted, because only full-length transcripts initiated from the promoter are isolated from the polyacrylamide gel. Although in this case the transcription efficiency may be compromised by ectopic initiation of transcription, the yield of riboprobe is still sufficient for multiple rounds of RNase protection assays. For RNA purification, we have used a protocol [modified from Autelitano et al. (23-25)] in which nuclear and cytoplasmic RNAs are isolated separately. Separation of these two species of RNA, in conjunction with the use of intron-exon splice junction probes, allows the effect of exogenous factors on nuclear and cytoplasmic RNA levels be evaluated separately [(23, 25), also see Fig. 3]. If sufficient RNA species are detected in a subsequent RNase protection assay, changes in heteronuclear RNA (hnRNA) level are usually easier to observe than those of mRNA after short-term treatment, as the more abundant mRNA requires a longer time of treatment to exhibit significant change. If nuclear and cytoplasmic RNAs are to be isolated separately, care must be taken to avoid excessive tissue homogenization that

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would disrupt nuclear integrity, and to avoid transferring any of the nuclear pellet into cytoplasmic fraction. In the absence of DNase digestion, a low level of contamination of DNA in the cytoplasmic fraction may significantly inflate the overall 0D260reading of the cytoplasmic RNA sample. A common problem encountered with the RNase protection assay is the appearance of multiple shorter band species, protected by the probe, that can accompany the "appropriate-sized" band. This is most often a concern when a lengthy X-ray exposure time is required such as when detecting lowabundance transcripts, such as an endopeptidase. This problem is primarily due to the hybridization and protection from RNase digestion of shortened radiolabeled cRNA or partially degraded RNA transcripts. Care, therefore, must be exercised to avoid ribonuclease contamination prior to hybridization. In addition, incomplete runoff transcripts in the probe labeling reaction will contribute significantly to the hybridization background but can be readily removed by isolating only the full-length transcripts from a denaturing polyacrylamide gel. Even purified full-length probes, however, will eventually break down to smaller species owing to autoradiolysis. Therefore, the best hybridization results are obtained when probes are used on the day they are prepared or within 2-3 days when stored at -80~ Probes stored for longer than 1 week should not be used. If the high background problem persists, other steps may be modified empirically to optimize the procedure for a particular probe. These include (a) determining the minimum but sufficient amount of probe to be used, (b) varying the hybridization temperature, (c) varying the amount of ribonucleases used, digestion temperature, or digestion time, and finally, (d) redesigning a shorter length version of the probe.

Example: Hormonal Regulation of Processing Enzyme Gene Expression in Developing Rat Pituitary Proopiomelanocortin synthesized in pituitary is under complex hormonal regulation, most of which is exerted at least in some part at the transcriptional level (23). Corticotropin-releasing factor (CRF), for instance, elicits an increase in both biosynthesis and release of POMC-derived peptides, with an accompanying change in POMC mRNA levels (28). Activation of the pituitary-adrenal axis, mediated principally by ACTH, results in elevated levels of circulating glucocorticoids, which in turn inhibits the biosynthesis and release of POMC peptides in the AL, but not the NIL. Previous work in the laboratory has shown that the transcriptional responsiveness of POMC to CRF is established by as early as el5 (25). Interestingly, in the NIL a transient inhibitory effect of glucocorticoids on POMC transcription exists at late prenatal and early postnatal ages, which is lost by pl0. As a first step

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AL

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2

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PC2 (136 bp)

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,~

POMC hnRNA(160 bp)

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FIG. 3 Distribution of POMC and PC transcripts in different adult rat pituitary lobes as determined by RNase protection assay. Autoradiogram of RNA samples from neurointermediate lobe (NIL) (2 /zg; lanes 1-4) and AL (3 /zg; lanes 5-8) after hybridization with POMC exon 1/intron A splice junction probe (lanes 1 and 5), PC 1 probe (lanes 2, 4, 6, and 8), PC2 probe (lanes 3, 4, 7, and 8), and RNase A/T1 digestion. Rat cyclophilin cRNA is also included in each lane as a standard to monitor the amount of total RNA applied. The position and length of each protected transcript are shown (arrows indicate the positions of protected PC2 transcripts). Use of POMC splice junction probe allows separate detection of hnRNA and mRNA, by virtue of the different-sized transcripts protected [see Scott and Pintar (25) for details]. Note that multiple probes protecting different lengths of specific mRNA can be incubated with a single RNA sample and subsequently separated by gel electrophoresis (compare lanes 2, 3, 6, and 7 with lanes 4 and 8). The autoradiographic film was exposed for 2 days at -80~ with an intensifying screen.

to investigate the possible mechanisms of how POMC-expressing cells in pituitary cope with altered demand of POMC processing under hormonal regulation, we have established the RNase protection assay to quantify the expression of processing enzymes PC1 and PC2 in the pituitary. Figure 3 shows that in adult pituitary, PC1 probe protects the corresponding transcripts in the appropriate 200-bp band, which are detected in RNA samples derived from both the NIL and AL. In contrast, the protected PC2 transcripts (136 bp) mainly appear in the NIL. This is in agreement with the abovedescribed results on lobe-specific expression of PC1 and PC2 RNA by in situ hybridization experiments. Although both PC1 and PC2 transcripts are apparently expressed in less abundance than POMC, the sensitivity of the assay allows a reliable quantitation of their expression level. This makes it possible to apply this method in conjunction with the short-term tissue culture

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procedure to determine whether the level of expression of PC1 and PC2 mRNA changes in concert with POMC in response to hormonal influence, and whether these responses are established in development.

Conclusions As a sensitive and reliable tool, in situ hybridization allows the visualization of the spatial pattern of gene expression. This technique can be readily applied to the study of the temporal pattern of processing enzyme gene expression in embryogenesis. This is aided by devising a reliable developmental staging system, allowing the determination of the precise age of each embryo under study. To assess the possible contribution of processing enzymes to the maturation process of an individual proprotein, both can be covisualized at the cellular level by virtue of dual labeling of probes for the processing enzyme and the potential substrate. On the other hand, to study the regulation of processing enzyme gene expression, isolation of the tissue of interest away from the in vivo context is usually advantageous. Various regulatory factors can be applied and their effects assayed by a quantitative measure, such as an RNase protection assay. Three particular examples are given to illustrate how an appropriate application of these techniques can advance our understanding of molecular mechanisms of proprotein processing in development.

Acknowledgments The authors are pleased to thank Dr. Nabil Seidah for providing the PC1, PC2, and furin probes, and Dr. Lloyd Fricker for providing the CPE probe used in this work. We also wish to thank Drs. James Roberts, Roderick Scott, Randal Streck, Joseph Cerro, and Teresa Wood for technical advice and assistance during these studies. This work was supported by Research Grants HD-18592 and DA-08622 from the National Institutes of Health to J.E.P.

References 1. R. E. Mains, E. I. Cullen, V. May, and B. A. Eipper, Ann. N. Y. Acad. Sci. 493, 278 (1987). 2. K. H. Cox, D. V. DeLeon, L. M. Angerer, and R. C. Angerer, Dev. Biol. 101, 485 (1984). 3. J. G. Gall and M. Pardue, Proc. Natl. Acad. Sci. U.S.A. 63, 378 (1969).

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I MOLECULARAPPROACHES H. A. John, M. L. Birnstiel, and K. W. Jones, Nature (London) 223, 582 (1969). 5. N. Wanek, K. Muneoka, D. G. Holler-Dinsmore, R. Burton, and S. V. Bryant, J. Exp. Zool. 249, 41 (1989). D. I. Lugo, J. L. Roberts, and J. E. Pintar, Mol. Endocrinol. 3, 1313 (1989). 7. S. Hayashi, I. C. Gillam, A. D. Delaney, and G. M. Tener, J. Histochem. Cytochem. 26, 677 (1978). L. H. Tecott, J. H. Eberwine, J. D. Barchas, and K. L. Valentino, in "In Situ Hybridization: Applications to Neurobiology" (K. L. Valentino, J. H. Eberwine, and J. D. Barchas, eds.), p. 3. Oxford Univ. Press, New York, 1987. P. A. Krieg and D. A. Melton, in "Methods in Enzymology" (R. Wu, ed.), Vol. 155, p. 397. Academic Press, Orlando, FL, 1987. 10. D. G. Wilkinson, in "In Situ Hybridization: A Practical Approach" (D. G. Wilkinson ed.). IRL Press, Oxford, 1992, p. 75. 11. P. J. Barr, Cell (Cambridge, Mass.) 66, 1 (1991). 12. W. J. M. Van de Ven, A. J. M. Roebroek, and H. L. P. Van Duijnhoven, CRC Crit. Rev. Oncogen. 4, 115 (1993). 13. L. D. Fricker, Annu. Rev. Physiol. 50, 309 (1988). 14. M. Zheng, R. D. Streck, R. E. M. Scott, N. G. Seidah, and J. E. Pintar, J. Neurosci., 14, 4656 (1994). 15. B. A. Eipper and R. E. Mains, Endocrin. Rev. 1, 1 (1980). 16. R. Day, M. K.-H. Schafer, S. J. Watson, M. Chr6tien, and N. G. Seidah, Mol. Endocrinol. 6, 485 (1992). 17. S. Benjannet, N. Rondeau, R. Day, M. Chretien, and N. G. Seidah, Proc. Natl. Acad. Sci. U.S.A. 88, 3564 (1991). 18. L. Thomas, R. Leduc, B. A. Thorne, S. P. Smeekens, D. F. Steiner, and G. Thomas, Proc. Natl. Acad. Sci. U.S.A. 88, 5297 (1991). 19. S. M. Sato and R. E. Mains, Endocrinology 117, 773 (1985). 20. J. E. Pintar and D. I. Lugo, Annu. N. Y. Acad. Sci. 512, 318 (1987). 21. K. Zinn, D. DiMaio, and T. Maniatis, Cell (Cambridge, Mass.) 34, 865 (1983). 22. M. Blum, in "Methods in Enzymology" (P. Conn, ed), Vol. 168, p. 618. Academic Press, San Diego, 1989. 23. D. J. Autelitano, M. Blum, M. Lopingco, R. G. Allen, and J. L. Roberts, Neuroendocrinology 51, 123 (1990). 24. R. E. M. Scott, D. J. Autelitano, D. I. Lugo, M. Blum, J. L. Roberts, and J. E. Pintar, Mol. Endocrinol. 4, 812 (1990). 25. R. E. M. Scott and J. E. Pintar, Mol. Endocrinol. 7, 585 (1993). 26. P. E. Danielson, S. Forss-Petter, M. A. Brow, L. Calavetta, J. Douglass, R. J. Milner, and J. G. Sutcliffe, DNA 7, 261 (1988). 27. E. T. Schenborn and R. C. Mierendorf, Jr., Nucleic Acids Res. 13, 6223 (1985). 28. J. L. Roberts, N. Levin, D. Lorang, J. R. Lundblad, S. Dermer, and M. Blum, Handb. Exp. Pharmacol. 104, 347 (1992). ,

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[4]

Use of Vaccinia Virus Vectors to Study Neuropeptide Processing Judy K. VanSlyke, Laurel Thomas, and Gary Thomas

Introduction Advances in cell biology have been greatly assisted by the application of vaccinia virus (VV) as an eukaryotic expression vector. Recombinant VV have been used successfully to express foreign proteins from viral, bacterial, protozoan, fungal, and metazoan sources (1, 2). Polypeptides of cellular origin include voltage-gated ion channels, G proteins, cell surface receptors, growth factors, neuropeptides, and proteinases from the regulated and constitutive secretory pathways of eukaryotic cells (3-11). Especially in the study of prohormone processing, VV recombinants have been useful in answering complex biological questions. Recombinants have been used to study the fates of prohormones and prohormone convertases in heterologous cell types (12). Large quantities of the prohormone and proprotein convertase are often produced and this feature has been exploited for purification of these molecules for in vitro analysis (13-16). Recombinant VV have been used in concert to reconstitute proprotein processing pathways in suitable mammalian cells in order to mimic the processes that occur in both neuroendocrine, endocrine, and nonendocrine cells (9-11, 17-20). These vectors have also been used in the study of targeting and localization of proproteins and proteinases in cellular compartments (8, 21-23). A gene transfer method to facilitate the study of the cell biology and enzymology of neuropeptide processing must meet several criteria. First, variability inherent to the expression system should be minimal. Second, reasonably high levels of expression are required to facilitate biochemical characterization of processed peptides (preferably >0.1 pmol/106 cells). Third, a study of tissue-specific processing requires the ability to express the prohormone in a variety of distinct cell types. Methods for gene transfer into mammalian cells can be divided into two general categories: those that result in stable integration of the foreign gene into the genome of heterologous cells, and those that produce transient expression. Each method has unique advantages and disadvantages (12). Expression systems based on random integration of foreign DNA into the host genome are generally subject to clonal variations. These may result from differences in expression of foreign protein as well as phenotypic drift Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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frequently associated with transformed cell lines. Isolation of stable transformants is both a labor-intensive and time-consuming process. Conversely, transient transfection systems do not suffer from clonal variation and results can be obtained expediently. Plasmid transfection efficiencies, however, vary with cell type and frequently low levels of expressed protein are obtained in these types of preparations. Typically, transfection approaches are limited to only a select cell type. By contrast, vaccinia virus does satisfy each of the above criteria. The construction of a vaccinia recombinant is rapid and not difficult. For example, purified recombinant virus can be prepared in less than 2 weeks. The vaccinia genome can accommodate large and/or multiple DNA inserts (24). Foreign genes are inserted by homologous recombination, precluding clonal variations. High levels of foreign protein can be obtained routinely by cloning the cDNAs along with vaccinia promoters (on the order of picomoles per 10 6 cells, using the VV 7.5K promoter) (25, 26). Infection with vaccinia is efficient. Between 80 and 100% of the cells in a population can be made to express the foreign protein. The broad host range of vaccinia allows information to be shuttled readily and rapidly between a variety of mammalian cell types and species. Importantly, experiments can be performed with primary cultures as well as established cell lines. Because cells can be infected simultaneously with multiple vaccinia recombinants, interactions among a group of foreign proteins can be studied in a defined cell system. Finally, vaccinia expression experiments can be performed within 24 hr. The development of gene transfer strategies has greatly enhanced the elucidation of the function of many molecules in higher eukaryotic cells. The goal of this chapter is to describe one gene transfer approach, recombinant vaccinia virus, in understanding the cell biology and enzymology of neuropeptide processing.

Vaccinia Virus Life Cycle The reasons why VV is a favorable expression vector become apparent when one considers the unique features surrounding the viral life cycle and genetic makeup. Vaccinia virus, a member of the orthopoxvirus family, is a doublestranded DNA (dsDNA) virus that replicates solely within the cytoplasm of an infected host cell (27). The 192-kbp genome encodes approximately 198 polypeptides that are expressed in a regulated fashion throughout the virus life cycle (28). After the virus enters the cell, partial uncoating of the virion takes place and early gene expression begins. The enzymes necessary for transcription (29) are packaged inside the virus particle and carried into the host cell during infection. Early gene expression produces the enzymes

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necessary for DNA replication and intermediate gene expression. As the life cycle proceeds, a second and complete uncoating event takes place, releasing the genomic material. As DNA synthesis begins, early gene expression ceases and intermediate genes are transcribed and translated. The polypeptides synthesized from these open reading frames include the transcription factors necessary for late gene expression. Subsequently, late genes are transcribed and translated, producing the constituents necessary for the virus particle formation.The infectious life cycle of vaccinia virus results in the formation of mature virions within the cytoplasm, referred to as intracellular enveloped virus (IEV), and extracellular enveloped forms (EEV) outside the host cell. Both forms of the virus are infectious, but EEV is thought to be the chief cause of cell-to-cell spread. The IEV form, however, is the one usually manipulated for expression vector work and the strains used in most laboratories produce large amounts of this virus (27).

Strategies for Making Vaccinia Virus Expression Vectors Recombinant DNA technology can be applied so that recombinant VV vectors can be tailored specifically for the desired application. For instance, VV recombinants can be constructed that will produce large amounts of a desired protein during viral infection (30, 31) or express a foreign gene only when an appropriate inducer is present (32, 33). Generally, a strategy for making a VV expression vector begins with selection of a recombinant plasmid that contains an appropriate viral promoter and genomic sequences that will allow homologous recombination to take place with the viral genome. [An example of such a plasmid, pZVneo (34), is represented in the top part of Fig. 1.] Generally, the promoter from the 7.5K gene of VV has been used in most expression vectors because it directs the efficient production of the gene of interest throughout the viral life cycle (35). Another important aspect to consider before making a recombinant VV is the method(s) for selecting the new virus away from the parental wild type. Insertional inactivation of a viral gene results in a loss of function that in some cases can be used as a means for selecting a recombinant. For instance, inactivation of the thymidine kinase (TK) gene can be selected for by growing virus in the presence of bromodeoxyuridine (BUdR), a lethal metabolite, in the presence of an active TK enzyme (36). Similarly, insertion into the hemagglutinin (HA) gene can be screened for by using an agglutination assay of erythrocytes (37). Most commonly the positive recombinant is selected out of a background of wild-type VV by screening thousands of plaques with a radioactively labeled DNA probe. With the development of liposomes for delivering plas-

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mid DNA into an infected host cell, the initial step in producing a recombinant virus (marker transfer) is easier and the transfer of DNA is more efficient (38). More powerful methods for selecting recombinants have been developed (2, 35). Genes that confer resistance to antibiotics or other drugs or have a reporter function have been used to select recombinant VV. To use this mechanism of selection to make a recombinant virus that expresses a foreign gene, dual cassette plasmids have been constructed and used (Fig. 1). Plasmids that contain two open reading frames, one being the gene for expression behind the VV 7.5K promoter and the other being the resistance or reporter gene under the direction of another vaccinia promoter, are flanked by portions of a VV gene. The promoters are abutted so that transcription is driven in opposing directions. These plasmids have been used successfully in constructing recombinant VV; selectable markers such as the neomycin transferase (39), guanine phosphoribosyltransferase (40), and fl-galactosidase genes (41) have improved the process of isolating the recombinants from parental virus. Described in detail below is a method for making a VV expression vector using the neomycin resistance gene in the recombinant plasmid pZVneo and selection with the antibiotic G418 sulfate (Geneticin; GIBCO-Bethesda Research Laboratories, Gaithersburg, MD). A schematic diagram of this procedure is represented in Fig. 1; however, procedures for growing, isolating, and manipulating vaccinia virus will be described initially to familiarize the reader with basic techniques that will be used throughout the process of isolating a recombinant virus.

Vaccinia Virus Methodology

General Vaccinia Virus Maintenance It is important to remember that vaccinia virus, although relatively harmless, is still a human pathogen. However, with a few safety precautions, potential hazards can be easily avoided. Virus can be killed with bleach, surfaces that have been near virus (such as counter tops and pipetting instruments) should be disinfected with Lysol, and disposable items should be autoclave sterilized. When working with virus, the researcher should always wear gloves, a laboratory coat, and protective eyewear. Most research institutions have guidelines for working with an infectious agent. The National Institutes of Health (NIH, Bethesda, MD) recommends routine vaccinations for laboratory personnel and safety level 2 procedures must be followed (42). With these guidelines in mind, the following protocols can be employed to maintain and utilize vaccinia virus in the laboratory.

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Infection of Cells with Vaccinia Virus Although stock preparations of vaccinia virus may vary, the basic protocol for infecting cells is the same. The general procedure is outlined below, but the variations will be described when appropriate. 1. Count the cells on a replicate plate: The cells are trypsinized from the plate, diluted with medium, and counted under the microscope with a hemocytometer. 2. Calculate the amount of virus to use to give a desired multiplicity of infection (MOI), which is recorded in plaque-forming units (PFU) per cell: With the known cell count, the titer of the virus (described below), the amount of virus to use can be calculated as follows: (No. of cells/plate)(PFU/cell)(103/A/ml) = virus (/A)/plate Viral titer (PFU/ml) 3. Prepare viral inoculum: A virus stock is thawed and sonicated (use a bath sonicator) for two 8- to 10-second intervals to disrupt aggregates (be careful not to overheat; place on ice intermittently). The appropriate amount of virus is diluted in PBS + MB [phosphate-buffered saline plus 1 mM MgC12 plus 0.1% (w/v) bovine serum albumin]. The thawed virus stock is stored at 4~ for 4-8 weeks and should be sonicated briefly before each use. 4. Infect cells with diluted virus: The cells are washed with PBS+M (phosphate-buffered saline plus 1 mM MgCI2), warmed to 37~ and overlaid with viral inoculum (just enough to cover the plate, i.e., 0.5 ml for 35- or 60-mm plates, 1.5 ml for 100-mm plates, and 5 ml for 150-mm plates). The cells are kept at room temperature for 30 min (or in some cases the cells are placed at 31~ for 2 hr); rocking at least once during the incubation is advisable to ensure that the cells remain covered with liquid. 5. Replace the medium: The viral inoculum is removed and the cells are washed once or twice with PBS+M and overlaid with culture medium, prewarmed to 37~

Large-Scale Preparation of Partially Purified Vaccinia Virus Most manipulations of vaccinia virus are executed from a central stock of partially purified virus. This form of the virus can be diluted easily and prepared for inoculating cells. To make a sizable quantity of virus, four confluent 150-mm plates of BSC-40 cells are routinely used for growth of the virus. 1. Infect plates at an MOI of 0.01 PFU/cell: Viral inoculum is prepared

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in 5 ml of PBS +MB per plate. The cells are watched between day 2 and 3 postinfection to ensure that they are harvested when approximately all of the cells have rounded up, but are still attached to the plate. 2. Harvest the infected cells: The cells should detach easily when gently triturated with the surrounding medium, using a serological pipette, but if they are still attached to the plate they can be scraped with a sterile rubber policeman. This procedure should be done gently to avoid cell rupture. 3. Pellet and wash the infected cells: The cells suspended in medium are pelleted in 50-ml conical tubes at 200 g for 5 min. The medium is removed and discarded, the cells are resuspended in 10 ml (per pellet) of PBS+M, collected in one tube, and centrifuged again. All of the following procedures are performed at 0-4~ 4. Hypotonically swell and disrupt the cells: After the wash solution is removed and discarded, the cells are resuspended in 10 ml of cold 10 mM Tris-HC1, pH 9, placed on ice for 10 to 30 min, and resuspended occasionally. This step causes the cells to swell and makes them more susceptible to mechanical rupture. The suspension is placed in a sterile Dounce (Wheaton, MiUville, NJ) homogenizer, with a tight pestle, and the cells are disrupted with 25 strokes, keeping the apparatus on ice the entire time. 5. Pellet the nuclei: After the cells are ruptured, the suspension is placed in a 15-ml conical tube and centrifuged at 800 g for 5 min at 4~ to pellet the nuclei. The resulting supernatant should be milky in appearance with a soft, white pellet at the bottom. The supernatant is transferred to a new tube, without disturbing the pellet. The pellet is then resuspended in 2 ml of 10 mM Tris-HC1, pH 9, and subjected to 25 additional pestle strokes. The mixture is recentrifuged and the supernatant removed and added to the portion collected from the first spin. 6. Pellet VV through a sucrose cushion: Two ultracentrifuge tubes (14 x 89 mm), each containing 6 ml of 36% (w/v) sucrose (in 10 mM TrisHC1, pH 9) and chilled to 4~ are carefully overlaid with the collected supernatant, placing 6 ml on each sucrose shelf. The tubes are balanced and placed in an SW41 rotor (prechilled) and centrifuged at 18,000 rpm for 80 min at 4~ 7. Harvest the virus pellet: The interface between sample and sucrose, which contains cell debris, is removed and discarded first so that it does not mix the pellet. The rest of the supernatant is discarded and the pellet that contains virus is resuspended in 800 ~1 of 10 mM Tris-HC1, pH 9. 8. Homogenize the virus suspension: The viscous suspension is placed in a sterile Duall homogenizer (Kontes Glass, Vineland, NJ) and the centrifuge tube is then washed with 100/~1 of 10 mM Tris, pH 9, which is added to the homogenizer. The virus mixture is homogenized with eight strokes of the pestle (the viscosity of the solution makes this difficult) to disrupt aggregates

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and returned to the ultracentrifuge tube. The homogenizer is then washed with 100/zl of 10 mM Tris, pH 9, and this is added to the partially purified virus stock. 9. Aliquot the partially purified virus preparation: The virus is dispensed into 25- to 40-/xl aliquots and stored at -70~

Titering Virus Stock To determine the titer of a particular vaccinia virus stock, confluent cell monolayers are needed for infection with various dilutions of the virus. Sixtymillimeter dishes are sufficient for determining viral titer. 1. Make a series of 10-fold dilutions of the viral stock: Ten microliters of a partially purified virus stock is placed into 990/zl of PBS + MB, sonicated, vortexed, and labeled as a 10-2-fold dilution. From this solution, virus is diluted sequentially in 10-fold increments 8 more times, vortexing between each dilution. 2. Infect confluent monolayers with the diluted virus solutions: Confluent monolayers are covered with 0.5 ml of three sequentially diluted virus solutions (e.g., 10 -7, 10 -8, and 10 -9 from a partially purified virus stock; 10 -6, 10 -7, and 10-8 from a crude virus stock). For more accurate determinations, the infections should be done in duplicate. Thirty minutes after inoculation, medium is replaced and the cells are incubated as previously described. 3. Approximately 36 hr postinfection, stain the plaques formed on the monolayers: The medium from each well or plate is removed and the cells are covered with 1 ml of a 0.5% (w/v) methylene blue solution in 50% (v/v) methanol. After about 5 min, the stain is removed and cells are gently rinsed two or more times with a 10% (v/v) solution of methanol until plaques are easily visible. 4. Count plaques and calculate the viral titer: A plate that has at least 30 plaques on it (but not more than 100) is selected for counting. After the number is determined, the titer is calculated as follows: (No. of plaques/0.5 ml)(2)(1/10 -n) = No. of plaque-forming units/ml where n is the dilution factor of the viral inoculum used on the monolayer of cells counted (step 2 above) and the number 2 corrects the value to a 1-ml volume.

Isolation of Vaccinia DNA Often it is necessary to obtain a large preparation of vaccinia virus genomic DNA. The following procedure describes isolation of VV DNA from one 100-mm plate of virally infected cells, but it can be scaled up for bigger preparations.

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1. Infect a 100-mm plate of confluent BSC-40 cells with VV at an MOI of 0.01. Virus is diluted into a 1.5-ml inoculum volume for overlaying the cells. 2. Remove the inoculum and replace with medium warmed to 37~ 3. Two days after infection, harvest the infected cells: Remove cells as described for making a large virus preparation, depending on whether they are floating or still attached to the plate. 4. Pellet the infected cells and resuspend them in 600/zl of PBS+M in a microfuge tube, using the same procedure as previously described. 5. Swell the cells by adding the following reagents to the cell suspension and place on ice for 10 min: 30/zl of 10% (v/v) Triton X-100, 1.5 ~1 of 2-mercaptoethanol (2-ME), 48/xl of 250 mM ethylenediaminetetraacetic acid (EDTA), pH 8. (Mix the reagents together first, then add to cells.) Vortex occasionally. 6. Pellet the nuclei in a microcentrifuge for 2 min at 3000 rpm. 7. Pellet the virus: The supernatant is removed (avoiding the pellet) to a new microfuge tube and centrifuged at 15,000 rpm for 10 min. 8. Gently resuspend the virus pellet in 100/zl of TE buffer, pH 8 (10 mM Tris-HCl, 1 mM EDTA, pH 8). Do not vortex the sample from this step forward. 9. Incubate the resuspended virus pellet with proteinase K. The following reagents are added to the virus suspension and gently mixed by flicking or rocking the tube: 1.5 tzl of a 10-mg/ml stock of proteinase K, 6.7/zl of 3 M NaC1, 0.3/xl of 2-mercaptoethanol, 10/xl of 10% (w/v) SDS. The solution is incubated at 50~ for 30 min and mixed occasionally. 10. Extract the mixture twice with phenol-chloroform (CHC13)-isoamyl alcohol (IAA) (made at a ratio of 25:24:1, v/v). a. An equal volume of phenol-CHC13-IAA is added to the mixture. b. The tube is shaken vigorously by hand 50 times. c. The suspension is centrifuged at 15,000 rpm for 2 min. d. The top layer is removed (slowly and avoiding the interphase) and placed in a new microfuge tube. Steps a through d are repeated with this solution. 11. Precipitate the DNA with ethanol and NaC1 twice. a. Four/zl of 3 M NaC1 and 250/zl of cold 100% ethanol are added to the solution (top layer from the last phenol extraction) and the mixture is mixed and placed at -70~ for 30 min. b. The precipitated DNA is centrifuged at 15,000 rpm for 15 min and the supernatant is removed and discarded (avoiding the pellet). c. The DNA is resuspended in 100/xl of distilled water and precipitated again with 5 ~1 of NaC1 and 250/xl of ethanol. Steps a and b are repeated.

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12. Rinse the pellet with cold 80% ethanol. a. Five hundred microliters of 80% ethanol is added to the tube with the pellet. The tube is centrifuged at 15,000 rpm for 30 sec and the supernatant is removed. b. The tube is dried completely under vacuum. c. The DNA is allowed to resuspend in 20/~1 of distilled water for at least several hours and is stored at 4~ The preparation is more viscous when there is more DNA present. 13. Determine the DNA concentration by optical density.

Making Recombinant Vaccinia Virus DNA Preparation The recombinant plasmid pZVneo contains a multiple cloning site featuring unique restriction enzyme recognition sites for insertion of the gene that is desired to be expressed. The foreign insert needs to contain a functional start and stop codon and should be oriented so that the 5' end of the open reading frame lies closer to the 7.5K promoter. Selection of recombinant plasmids can utilize the ampicillin resistance gene supplied by the Bluescript (Strategene, La Jolla, CA) backbone or the neomycin resistance gene abutted to the VV 11K promoter. Apparently the VV promoter is recognized by the Escherichia coli RNA polymerase and confers resistance to kanamycin to the bacterium housing the plasmid. This feature is used to select pZVneo away from any other contaminating plasmids (e.g., an uncut version of the plasmid utilized to isolate the insert DNA). After the insert is cloned into the plasmid and the DNA sequence checked for spontaneous mutations, a large-scale preparation of plasmid DNA is made using either CsCI gradients or polyethylene glycol (PEG) precipitation.

Marker Transfer Protocol HeLa or BSC-40 cells are prepared in advance so that they are 50 to 80% confluent in a 35-mm plate. HeLa cells are cultured in Dulbecco's modified Eagle's medium (DMEM) containing 10% (v/v) fetal calf serum (FCS) at 37~ and 5% CO2 and BSC-40 cells are grown under the same conditions except that MEM is used. 1. Determine the number of cells on a replicate plate. 2. Infect cells with wild-type virus (MOI = 0.5 or 1.0): Virus is diluted in 0.5 ml of PBS+MB and sonicated to disrupt aggregates. 3. Prepare the DNA-lipid mixture: Fifteen minutes before the end of

[4] VACCINIA VIRUS VECTORS

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inoculation time period (30 min), the lipofectin-DNA mixture is prepared by bringing 5 /~g of plasmid DNA and 1 /~g of wild-type VV DNA (both resuspended in sterile distilled H 2 0 ) up to a total volume of 50/A in water. (VV DNA is included for optimal recombination efficiency, but is not absolutely required.) In a polystyrene tube, 20 ~1 of lipofectin (Cat. No. 8292A; GIBCOBRL) is mixed with 30/A of sterile water. The DNA is then gently added to the lipofectin mix and the tube is gently flicked to mix. This mixture is left at room temperature for 15 min while the liposome-DNA complexes form. 4. Overlay infected cells with the transfection medium: The inoculum is removed and the cells are washed three times with PBS + M warmed to 37~ Nine hundred microliters of the appropriate medium (depending on the cell type) minus serum is added to the polystyrene tube, mixed gently, and overlaid on the infected cells. 5. Allow the transfection to proceed for 3 to 4 hr and then add medium supplemented with serum and G418 sulfate: The cells are placed in the incubator for 3 to 4 hr, at which time 1 ml of the appropriate medium plus 10% (v/v) fetal calf serum plus G418 sulfate (4 mg/ml) is added to the plate (without removing anything first) and mixed gently. The plate is returned to the incubator. The concentration of G418 sulfate refers to total weight. Although the active weight (recorded on the label) varies from bottle to bottle, it usually comprises ->50% of the total weight. The antibiotic is added to medium, allowed time to dissolve, and the medium is sterilized by passing through a syringe filter (0.22-/~m pore size) before applying to infected cells. 6. Harvest the cells from the marker transfer step: After 24 hr, if the cells are still adherent to the bottom of the plate, they are scraped up with a sterile rubber policeman. Cells are collected by centrifugation as previously described, the medium is removed, and the cells are washed with 2 ml of PBS+ M before centrifuging again. All procedures are performed at 4~ 7. Prepare a crude stock: Cells are resuspended in 0.5 ml of PBS+M and a crude stock is prepared by putting the suspension through three freeze/ thaw cycles with liquid N2 and a 37~ water bath. The crude virus stock should be stored at -70~ until use. At this point the crude stock will have an approximate titer of 1 • 105 PFU/ml and recombinant viruses should constitute approximately 5% of the population. A marker transfer reaction without pZVneo present can be included and carried along through the process of selecting a recombinant virus to ensure that the drug selection is working.

Drug Selection The crude viral stock is grown in BSC-40 cells in the presence of the G418 sulfate (2 mg/ml) to select for viruses that contain the neomycin resistance gene.

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1. Infect cells at an MOI of 0.01 from the crude stock made from the marker transfer reaction: A 35-mm dish with a confluent monolayer (cell count known) is infected in the same way as described before for the marker transfer procedure. 2. Overlay infected cells with medium containing G418 sulfate: MEM plus 10% (v/v) FCS plus G418 sulfate (2 mg/ml) is prepared as previously described. After the viral inoculum has been on the cells for 30 min, it is removed and the cells are washed with PBS+M warmed to 37~ which is replaced with medium containing antibiotic. 3. Harvest the cells and make a crude stock: After 2 or 3 days, if cytopathic effects of the viral infection are clearly visible (cells are rounded up), cells can be harvested in the same way as described for the marker transfer protocol. At this point the titer of the virus stock will be around 5 x 10 6 PFU/ml and between 30 and 50% of the population will be bona fide recombinants. Titering the crude stocks is recommended to ensure that the drug selection is working and to determine titers for calculating the amount to use in generating a specific multiplicity of infection, but estimated titers can be used so that the selection process is not delayed. Another round of growth in the presence of the antibiotic will increase the titer and the proportion of recombinants in the total virus population. However, if time is of the essence, this is not absolutely necessary.

Methods for Plaque Purification of Recombinant Virus An isolated plaque should represent a clonal population of virus originating from a single infectious virion. With this in mind, the next step in purification involves generation of well-isolated plaques and isolation of the recombinant virus. If G418 sulfate selection is used, plaques are isolated through agarose overlays containing antibiotic. If drug selection cannot be used, isolated plaques are screened and recovered from filter lifts prior to a final purification from agarose overlays. Both methods are described here. Isolation from under Agarose Overlays At this point, the crude stocks that have been passaged through cells in the presence of G418 sulfate can be plated on cell monolayers for plaque purification. Confluent monolayers of BSC-40 cells in 100-mm dishes are prepared. 1. Calculate the amount of viral inoculum to use: Keeping in mind both the titer of virus in the last crude stock and the percentage of the population

[4]

2.

3.

4.

5.

VACCINIA VIRUS VECTORS

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that is actually recombinants, dilutions are made so that about 50 recombinant viruses will infect 1 plate. (It is best to try plating out several different dilutions to ensure that there will be enough plaques.) Infect confluent monolayers with virus from the crude stock produced at the last drug selection step: The infection protocol is performed as already described. Prepare agarose medium overlay containing G418 sulfate. a. Agarose mixture: SeaPlaque agarose (FMC, Philadelphia, PA) is mixed with distilled water (0.12 g/5 ml) and sterilized by autoclaving for 15 min only. [A larger sterile preparation of 2.4% (w/v) agarose can be made ahead of time and stored until needed. Once autoclaved, agarose can be melted in a microwave.] b. Medium mixture [20% (v/v) FCS, G418 sulfate (4 mg/ml)]: Prepare 4 ml of 2x strength sterile MEM plus 1 ml of fetal calf serum plus 20 mg of G418 sulfate (dissolved completely). Filter sterilize and place in a 37~ water bath. c. Mix agarose and medium in a 1" 1 ratio" Once the agarose solution is cooled to about 45~ it is mixed with the above medium. After removing the viral inoculum and washing the cell monolayer, 8 ml of medium plus agarose is carefully layered over the cells. The agarose is allowed to solidify at room temperature (without rocking the plate) and placed in the 37~ incubator. After 48 hr, stain the plaques grown under agarose: Prepare sterile molten 1% (w/v) SeaPlaque agarose in PBS. After the solution is cooled to 45~ a sterile stock solution of Neutral Red is added (final concentration is 0.02%, v/v). The agarose is carefully overlaid on the preexisting agarose, beginning in the center of the plate. Again the plate is allowed to sit at room temperature for 5 min before transferring to a 30~ incubator. The plaques are usually visible within 2 hr, but selection can be done as late as the next day. They will appear small (if compared to virus grown under agarose without drug selection) but only the recombinant virus should grow in the presence of the drug. Pick plaques from under the agarose overlay: Tubes containing 0.25 ml of PBS§ are prepared for as many plaques as will be selected. A pipettor (20-200 /~1, with the dial set at 20 /~1) is used and the tip is exchanged between each selection. Virus is isolated by punching the pipette tip through the agarose directly above a well-isolated plaque and suctioning up and down gently a few times, trying to capture a little of the cell layer at the edge of plaque. The pipette tip is then placed in a tube of PBS+MB and the solution is used to rinse the tip by pipetting action. Clearly established plaques should be preferentially selected. Isolated plaques should be frozen and thawed three

78

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M O L E C U L A R APPROACHES

times and stored at -70~ immediately.

if not taken through an amplification step

Isolation from Filter Lifts Confluent monolayers of BSC-40 cells in 100-mm dishes are required for this procedure. 1. Infect monolayers with virus from the crude stocks from the marker transfer step or the drug selection step: Infect cells as previously described with a dilution that will generate around 100 plaques/plate. 2. Replace viral inoculum with medium: Remove viral inoculum, wash cells with PBS+M warmed to 37~ and overlay with MEM plus 10% (v/v) FCS. 3. After 48 hr, stain the cells with Neutral Red: Prepare a room temperature solution of PBS plus 0.01% (v/v) neutral red and add 5 ml directly to the medium on plates of infected cells. Transfer the plates to a 30~ incubator. 4. Transfer plaques to a filter. a. Prepare 100-mm dishes for storing filter lifts: Invert the dishes and place a Whatman (Clifton, NJ) 3MM filter circle in the lid. Add 1 ml of PBS + M to each filter. b. Remove medium from the culture plate. c. Carefully place a nylon filter on the monolayer of cells. Do not let it slip after contact with the plate surface. Press a Kimwipe dampened with PBS+M on any area of the filter that is dry and not absorbed to the cell monolayer. d. Lift the filter off the plate and place it plaque side up on the moistened Whatman circle. e. Immediately make a replicate filter lift: Place a replicate nitrocellulose filter on the plaque side of the nylon filter and press them together. (Again, do not move after contact has occurred.) With a paper punch, make holes in the paired filters for orientation later. With forceps, separate the two filters and place the nitrocellulose filter on the Whatman circle in the dish and place the nylon filter on paper toweling to dry. Seal the dish containing the filter with Parafilm and store at -70~ 5. Prepare nylon filter lifts for hybridization. a. Float the filters on denaturing solution for 10 min. Denaturing solution: 1.5 M NaC1, 0.5 M NaOH

[4]

VACCINIA VIRUS VECTORS

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b. Float the filters on neutralizing solution for 2 min. Repeat. Neutralizing solution: 3.0 M NaCI, 0.5 M Tris-HC1, pH 7.5 c. Wash the filters (submerge) briefly in SSC. SSC: 0.3 M NaC1, 30 mM sodium citrate d. Let the filters air dry, then bake them for 20 min at 80~ e. Incubate the filters in proteinase K buffer plus proteinase K (50 ~g/ ml) enough to cover the filters, at 50-55~ for 30 min. Proteinase K buffer: 100 mM Tris-HC1, pH 8, 150 mM NaC1, l0 mM EDTA, 0.2% (w/v) SDS. Proteinase K is added just before incubation period 6. Prehybridize the filters for 2 to 4 hr at 37~ Prehybridization solution: 50% (v/v) deionized formamide, 1 M NaC1, 10% (w/v) dextran sulfate, 1% (w/v) SDS, 25 /~g of sheared salmon sperm DNA (heated to 100~ for 10 min immediately before adding to the prehybridization solution) 7. Hybridize overnight at 37~ Add denatured radioactive probe (randomprimed DNA labeling kit; Boehringer Mannheim, Indianapolis, IN) directly to prehybridization solution. 8. Wash the filters in the following manner: a. Submerge in SSC at room temperature for 10 min. Repeat. b. Incubate in SSC plus 1% (w/v) SDS at 65~ for 30 min. 9. Dry the filters. Mark orientation marks with luminescent dye. Place under X-ray film. 10. Isolate "positive" plaques from the nitrocellulose filter. a. Line up a positive signal on the film with a plaque on the nitrocellulose filter. b. With a sterile paper punch or razor blade, excise the area containing the plaque and put it in a tube containing 200 ~1 of PBS+MB. c. Freeze/thaw the preparation three times and store at -70~ if not proceeding through the amplification step immediately.

Amplification of Plaque-Purified Vaccinia Virus Isolate When the plaques are ready for selection, a 24-well dish or 100-mm plates should be ready with confluent monolayers of BSC-40 cells. 1. Infect the 24-well plate with isolated plaques.

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a. From agarose overlay: After all the desired plaques are selected, the medium on the 24-well plate is aspirated. A 1-ml pipettor is used to homogenize the agarose plug in PBS + MB, by passing it up and down through the pipette tip multiple times, and deliver the solution to an individual well. The tip is exchanged after each inoculation and each well is inoculated with a different viral isolate. b. From filter lift: Infect 1 well of a 24-well plate with 100/xl of the frozen and thawed viral inoculum. If a larger crude stock is desired, dilute 5 to 20/xl of the viral inoculum up to 1.5 ml with PBS+MB and infect a 100-mm dish of cells. Replace with regular medium. 2. Replace viral inoculum with medium containing antibiotic. a. From a 24-well plate: Prepare MEM plus 10% (v/v) FCS plus G418 sulfate (2 mg/ml). After 30 min, the viral inoculum is removed from each well, using a Pasteur pipette with a 20- to 200-/zl pipette tip on the end. The tip is replaced after each aspiration. The cells are then overlaid with medium containing G418 sulfate, avoiding any crosscontamination between the wells. The infections are allowed to proceed for 2 days before cell harvest. b. From a 100-mm plate: Remove the viral inoculum and replace with regular medium warmed to 37~ 3. After 2 days, harvest the cells and make crude stocks: Prepare crude stocks of the virally infected cells in the 24-well plate. Aspirate the medium (using a new pipette tip each time), overlay the cells with 0.3 ml ofPBS+ M per well, and perform three freeze/thaw cycles by placing the plate alternately at -70 and 37~ Screen for desired recombination events and purity of the virus stock by methods described below.

Screening for Recombinants and Purity Multiple crude stocks are screened for the presence of recombinant VV and the purity of the stock. Four methods of doing this are discussed below. Slot (Dot)-Blot Screening A portion of the crude stocks (100/xl) is transferred to a nylon filter through a slot-blot manifold and standard Southern analysis (43) procedures are followed to screen for the presence of the insert DNA. This screening method will identify crude stocks containing recombinant virus and the intensity of the radioactive signal will indicate the extract of enrichment. It will not show if the recombinant virus stock is still contaminated with wild-type VV. DNA Isolation and Southern Analysis Purity of the crude stock can be determined by analyzing the genomic VV DNA. A large preparation of VV DNA is isolated from an infected 100-mm plate, cleaved with the appropriate restriction enzyme (usually HindlII), and

[4] VACCINIA VIRUS VECTORS

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subjected to electrophoresis in a 0.7% (w/v) agarose gel (43). Wild-type DNA is included for comparison so that the change in band patterns can be examined. The DNA is transferred to a nylon filter and subsequently probed with appropriate radiolabeled nucleic acids to determine the presence of foreign DNA and absence of wild-type viral DNA.

Immunoblot Analysis Another method of screening for the desired recombinants requires immunological reagents that recognize the foreign protein expressed from the recombinant virus in an immunoblot analysis. If this method is used, a replicate infected 24-well plate is preferable so that the cells can be harvested directly in a buffer compatible with SDS-polyacrylamide gel electrophoresis (43). This method may also indicate which stocks are at higher titers than others. However, virus purity is not discernible with this approach.

Polymerase Chain Reaction Analysis The last method outlined for screening multiple virus stocks utilizes the sensitivity and speed of the polymerase chain reaction (PCR) (43, 44). DNA is isolated from crude stocks by the method described for the VV DNA isolation procedure, beginning with the proteinase K treatment. Three primers, one complementary to the 5' end of the TK gene, one complementary to the 3' end of TK, and one complementary to the foreign DNA of interest, are used to prime the PCR reaction (Fig. 2A). The primers are oriented such that if the TK gene has not been inactivated, a small fragment will be generated. If foreign DNA had been inserted then the primer corresponding to the 3' end of the TK gene and the insert primer will amplify a somewhat larger PCR product. The product that would be made from the two TK primers across the insert is much too large to occur efficiently. Therefore, a putative recombinant viral stock could potentially generate a small product, a larger product, or both, as seen in the samples screened and analyzed by gel electrophoresis (Fig.2B). The presence of both a smaller and a larger PCR product indicates either that the crude stock still has wild-type VV or that a single-site cross-over event has occurred during homologous recombination (e.g., Fig. 2B, lane 2). However, a sample that generates only a larger PCR product represents a purified recombinant virus stock (e.g., Fig.2 B, lanes 3 and 4). The differential size is important so that the results are easily visualized on an agarose gel. The PCR products are all kept around or under 1 kb to maintain efficiency, with the band that denotes the presence of wildtype VV DNA being the smallest. This ensures the sensitivity for the presence of contaminating wild-type virus.

82

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FIG. 2 Polymerase chain analysis ofVV DNA from putative recombinant virus crude stocks. (A) Potential VV DNA templates present in a crude stock of recombinant virus are shown with the three primers (discussed in text) aligned next to their complementary regions within the DNA. The arrows indicate the direction in which DNA synthesis will occur. Filled bars below the template and primers represent the potential products generated by PCR when either VV DNA is present (**, recombinant VV DNA specific product; *, wild-type VV DNA-specific product). (B) The dsDNA products generated by PCR reactions containing plasmid DNA (pZVNEO construct) used to make the recombinant virus (lane 1), VV DNA from three different crude stocks of recombinant virus (lanes 2-4), or wt VV DNA (lane 5) are separated on a TBE-agarose gel. The positions of nucleic acid markers are indicated by size (in bp) on the left-hand side of the photograph and the two potential PCR products are marked as noted in (A) on the right.

[4] VACCINIA VIRUS VECTORS

83

Experimental Strategies to Investigate Neuropeptide Processing Using Vaccinia Recombinants To illustrate the utility and efficacy of vaccinia technology in addressing the cell biology and enzymology of neuropeptide processing, a summary of the work that utilized this gene transfer vector to elucidate many of the molecular mechanisms that govern the tissue specificity of proopiomelanocortin (POMC) processing is presented. The studies are divided into three sections, each exploiting features unique to the vaccinia expression system. First, we show the use of vaccinia technology to assess the ability of heterologous cell types to mimic the tissue-specific processing of POMC. Second, we show how vaccinia can be used to express multiple foreign genes simultaneously to permit study of their interactions. Specifically, we show the reconstitution of POMC processing by the PC2 and PC3 (also called PC1) proprotein convertases. Third, we show how the vaccinia vector can be used to express significant levels of bona fide prohormone to use as a substrate for prohormone processing in vitro.

Expression of Proopiomelanocortin in Heterologous Cells Using a Vaccinia Recombinant To identify heterologous cell types that mimic the tissue specificity of POMC processing as it occurs in the anterior (AL) and neurointermediate (IL) lobes of the pituitary (Fig. 3), a VV recombinant that directs the synthesis of mouse POMC (VV : mPOMC) was constructed and used to express the prohormone in a variety of endocrine and nonendocrine cell types (18). The cell lines studied include several nonendocrine [e.g., BSC-40 (African Green monkey kidney epithelium) and Ltk- (mouse fibroblast)] and several endocrine and neuroendocrine [e.g., G H 4 C 1 (rat somatomammotroph), NG108 (mouse neuroblastoma x rat glioma), Rin m5F (rat insulinoma)] cell lines. In addition, the expression and processing of POMC was studied in primary cultures of bovine adrenomedullary (BAM) cells. For most cell lines, infections were performed as described in General Vaccinia Virus Maintenance, above (MOI = 5). However, for some cell types (e.g., Rin m5F and BAM), the inoculum was applied to the cells for 2 hr at 31~ and then the infected cultures were refed with medium containing 2% (v/v) serum. The efficiency of infection in all cases is high; between 80 and 100% of the cells in a culture will express the foreign protein. After 16 hr the media were removed and frozen and the rinsed cells were extracted with 1 M acetic acid (pH 1.9). The level of POMC expression as measured by an adrenocorticotropin (ACTH)

84

I MOLECULAR APPROACHES N-TERM

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FIG. 3 Cell type-specific processing of mouse preproopiomelanocortin (mPOMC). Shown are the primary POMC peptides produced in pituitary anterior lobe corticotrophs and neurointermediate lobe melanotrophs. Also depicted are the primary POMC processing products produced in heterologous cells expressing mPOMC with a vaccinia vector. Cell types that express only the prohormone ("maturation deficient") include BSC-40, Ltk-, HeLa, PC12, GH4C1, and P388D1 (an interleukin 1-secreting monocyte line). Primary cultures of bovine adrenomedullary chromaffin cells (BAM) process mPOMC just as do anterior lobe corticotrophs, whereas the rat insulinoma

[4] VACCINIA VIRUS VECTORS

85

radioimmunoassay (RIA) ranged between 0.7 and 3.7 pmol of ACTH per 106 cells in cell extracts and between 5 and 22 pmol of ACTH per 106 cells in the medium samples, depending on the cell type examined. Peptide analysis showed that vaccinia-mediated expression of POMC in a number of cell lines including BSC-40, NG 108, and GH4C 1 resulted in the efficient production of bona fide POMC (see, e.g., Fig. 4A) (8, 45). Studies of stimulated secretion showed the prohormone was packaged into the regulated pathway in GH4C 1 cells (8). None of these cell types was capable of processing the prohormone; hence they were termed "maturation deficient" (for neuroendocrine peptide precursors). In contrast, expression of POMC in the rat insulinoma (Rin m5F) resulted in the efficient processing of the ACTH and/3-1ipotropin (/3-LPH) domains in the regulated pathway to sets of peptides reminiscent of neurointermediate lobe melanotrophs. Both the ACTH and/3-LPH domains were efficiently processed to corticotropin-like intermediate lobe peptide (CLIP), 7-LPH, and/3-endorphin (1-31) (8, 45). Expression of POMC in primary cultures of BAM cells resulted in the targeting of the prohormone to the regulated pathway (23). Stimulation of secretion with 3 mM BaCI 2 elicited an 11.5-fold increase in the secretion of POMC immunoreactivity (IR) in 30 min, with greater than 50% of the intracellular contents being released. Peptide analysis showed that the BAM cell endoproteases processed POMC to generate peptides present in anterior lobe corticotrophs; ACTH and/3-LPH were the primary peptide products with limited cleavage of/3-LPH to y-LPH and/3-endorphin (1-31). These studies exemplify several prominent features of the vaccinia system. First, a wide variety of cell types, both established cell lines and primary cultures, can be screened rapidly to assess their processing phenotypes. Second, sufficiently high levels of foreign protein are expressed that a complete characterization of product peptides can be performed from as little as two 150-mm plates of cells (---3 • 107 cells). Third, vaccinia does not affect the ability of the prohormone to be targeted to the regulated pathway, where it is correctly and efficiently processed.

Rin m5F processes mPOMC in a manner similar to the neurointermediate lobe melanotrophs (no detectable processing at Lys232Lys233).( | ) cleavage site; P, phosphorylation; (73), O-linked carbohydrate; ( 9 N-linked carbohydrate. Note that open symbols represent partial glycosylation whereas closed symbols denote complete glycosylation. The amino acid numbering of the prohormone begins with the initiator methionine. Data describing these gene transfer studies have been previously reported (8, 23, 45).

86

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retention time (min)

1 0 2 0 3 0 4 0 5 0 6 0 7 0

retention time (mln)

[4] VACCINIA VIRUS VECTORS

87

Reconstitution of Prohormone-Processing Pathways Unique features of vaccinia biology afford a powerful gene transfer approach to assess the ability of several candidate processing enzymes to cleave precursor proteins in the secretory pathway. This approach exploits the maturationdeficient cell lines described above as well as the ability to use VV recombinants to express multiple foreign genes simultaneously in a cell population. Vaccinia offers two strategies for coexpression of multiple foreign genes in heterologous cells. These are either coinfection with multiple vaccinia

FIG. 4 Processing of mPOMC by Kex2, PC3, and PC2 in vivo and in vitro. (A) Replicate 150-mm plates of BSC-40 cells (---2 x 107 cells) were coinfected with VV recombinants expressing VV : mPOMC (2 PFU/cell) and VV : WT, VV : KEX2, VV : mPC3, and VV :hPC2 (each at 5 PFU/cell, 7 PFU/cell total), or VV :mPC3 and VV :hPC2 together (each at 3 PFU/ceI1, 8 PFU/cell total). After 16 hr, the medium was removed and replaced with serum-free minimal essential medium-0.07% (w/v) bovine serum albumin. After an additional 2 hr, this medium was recovered, adjusted to 0.2% (v/v) trifluoroacetic acid, and applied to a C4 reversed-phase column. Retained material was eluted with a gradient of acetonitrile and 1-min fractions were collected and assayed for ACTH (+)-,/3-endorphin (O)-, or y-LPH (O)-immunoreactive material. Elution positions of mPOMC-derived peptides from the mouse corticotroph cell line AtT-20 are indicated. [From Thomas et al. (9); a description of the methods used to characterize each POMC peptide fully is presented in references (8, 9, 16, 18, 19, 23). (B) Conversion of fl-LPH to y-LPH and fl-endorphinl_31 in the regulated pathway. At 36 hr after plating, replicate plates of chromaffin cells were coinfected with VV : mPOMC (2 PFU/cell) and either VV :WT, VV : KEX2, or VV : hPC2, each at 5 PFU/cell (7 PFU/cell total). Sixteen hours after infection, cells were washed and incubated for 2 hr in medium containing cycloheximide at 50/zg/ml. The cells were again washed and incubated 45 min in Ca2+-free balanced salt solution containing cycloheximide with or without 3 mM BaC12. This medium was collected, adjusted to 0.1% (v/v) trifluoroacetic acid, and resolved on the C4 column, as described in (A). Aliquots from 1-min fractions were assayed for fl-endorphin- and y-LPHimmunoreactive material. No corresponding y-LPH IrM peptides were detected in control samples (no Ba2+; top panel). (C) Radioimmunoassay analysis of mPOMC processing products generated by insulin secretory granule endopeptidase activity. mPOMC processing by Triton X-114 aqueous phase-extracted material from insulin secretory granule lysate over 3.5 hr at 30~ was assessed as described (16). The mPOMC products were resolved by Vydac reversed-phase C4 HPLC, 5-/zl aliquots of the 1-ml fractions were assayed by specific RIA to regions of mPOMC, and the total amount of product per fraction was calculated. The elution positions of various POMC processing product standards are indicated. The RIA analysis was for flendorphin (0), y-LPH (O), and ACTH (+).

88

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recombinants or construction of vaccinia recombinants containing two (dual recombinant) or three (triple recombinant) foreign genes as vectors. Marker transfer plasmids that direct recombination into either the VV HindIII J (e.g., pZVneo), C, or A regions have been reported (35) and each has been used successfully to construct dual or triple recombinants (B. A. Thorne, L. Thomas, and G. Thomas, unpublished results, 1994). In most experiments, however, the use of multiple single recombinants is satisfactory. To begin identification of the mammalian processing endoproteases, we developed a strategy based on knowledge gained from studies in Saccharomyces cerevisiae that provided the first unequivocal identification of the gene encoding a eukaryotic processing endoprotease, KEX2 (46). In our initial studies we showed that coexpression of POMC and Kex2p in any of the maturation-deficient cell lines shown in Fig. 3 resulted in the efficient processing of the prohormone on the C-terminal side of-LysArg- doublets to yield fl-endorphin (1-31) and y-LPH (18) (see, e.g., Fig. 4A, POMC/WT vs POMC/KEX2). The studies on KEX2 showed that this endoprotease is functionally and, thus, structurally similar to the mammalian processing endoprotease(s). Indeed, a number of DNA sequences encoding a family of higher eukaryotic KEX2-1ike endoproteases has been reported (47, 48). The possibility that two of these DNAs, PC2 and PC3, are prohormone convertases became readily apparent because they (a) are KEX2-1ike, (b) are neuroendocrine specific, and (c) their relative RNA distribution in different tissues matched what our gene transfer studies predicted-both AL and BAM preferentially express PC3 RNA compared to PC2 RNA whereas both IL and Rin cells preferentially express PC2 RNA compared to PC3 RNA. Vaccinia recombinants expressing mouse PC3 and human PC2 were constructed as described earlier and were used in expression studies to examine their ability to cleave mPOMC (9). Cells were then coinfected with two or three vaccinia recombinants, using the standard inoculation protocol. The multiplicity of infection was adjusted such that any cell expressing the substrate will also express the enzyme (see caption to Fig. 4A). Depending on the cell type and endoprotease used, processed peptides are isolated from either cell extracts or from the culture medium. For example, with KEX2 sufficient quantities of processed peptide accumulate within cellular compartments to allow peptide identification directly from cell extracts. Similarly, with PC2 expressed in BAM cells, sufficient quantities of processed peptide accumulate in secretory granules to allow peptide identification from cell extracts. By contrast, with PC2 and PC3 expressed in BSC-40 cells, insufficient amounts of cellular peptides accumulate to perform biochemical characterization. Therefore peptides are isolated from the culture medium. For these experiments, secreted peptides are collected either for 16-18 hr in a

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serum- and Phenol Red-free defined medium, MCDB 202, or for 2 hr (beginning 16 hr postinfection) in MEM containing 0.07% (w/v) bovine serum albumin. When secreted material is analyzed, it is critical to perform a mixing experiment as a control to ensure that the processed peptides were generated in the cell by the candidate endoprotease rather than artifactually by extraneous proteases in the medium. Coexpression of POMC and PC3 in BSC-40 cells results in the production of peptides similar to those of anterior lobe corticotrophs; ACTH and/3LPH were the prominent processing products (Fig. 4A, POMC/PC3). Coexpression of POMC and PC2 resulted in the efficient excision of/3-endorphin (1-31) and a processing intermediate containing the ACTH and ),-LPH sequences (ACTH/3,-LPH, open arrowhead) (Fig. 4A, POMC/PC2). However, coexpression of PC3 and PC2 together with POMC in BSC-40 cells resulted in ACTH and an insulinoma-like pattern of cleavages in the/3-LPH domain, with efficient processing to ~/-LPH and /3-endorphin (1-31) (Fig. 4A, POMC/PC3+PC2). The efficient processing of/3-LPH to ~/-LPH and/3-endorphin (1-31) by PC2 in BSC-40 cells suggested that this enzyme is capable of catalyzing one insulinoma (IL) cleavage but the conversion of ACTH to a-MSH was not detected. Therefore we chose to determine whether coexpression of PC2 and POMC in cells containing a regulated pathway (BAM cells) would enable PC2-directed processing of ACTH. In addition, these studies would allow us to determine whether PC2 could cleave POMC in the regulated pathway~a requirement for a prohormone-processing endoprotease. As a negative control, POMC was coexpressed with Kex2p, which also efficiently cleaved/3LPH in BSC-40 cells but, being a yeast enzyme, should not be capable of regulated pathway processing. Finally, these experiments allowed us to determine whether the tissue specificity of prohormone processing can be explained simply by a differential expression of PC3 and PC2. If so, then overexpression of PC2 in BAM cells should result in an insulinoma-processing phenotype. Briefly, replicate plates of BAM cells were coinfected with VV : mPOMC and either VV" WT, VV" KEX2, or VV" hPC2. Sixteen hours postinfection, cells were washed and incubated for 2 hr in medium containing cycloheximide (50/xg/ml). This allowed any processed peptides present in the constitutive pathway to be secreted while those present in the regulated pathway are retained. The cells were again washed and incubated for 45 min in Ca2+-free balanced salt solution containing cycloheximide with or without 3 mM BaC12 (Fig. 4B). This medium was collected and analyzed for POMC peptides by high-performance liquid chromatography (HPLC) (reversed phase and cation exchange) coupled with domain-specific radioimmunoassay. The results show that coexpression of POMC with PC2, but not Kex2p, resulted in

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processing of/3-LPH to y-LPH and /3-endorphin (1-31) in the regulated pathway, demonstrating that PC2 has specialized properties that enable it to be routed to, and function in, this compartment (compare Fig. 4B, bottom, with second and third panels). In addition, PC2 was able to cleave ACTH to a-MSH correctly. The vaccinia reconstitution system has also been used to show that PC3 and PC2 together, and not separately, are capable of cleaving proinsulin correctly when coexpressed in maturation-deficient cells (20). Similarly, PC2 cleaves pro-LHRH to luteinizing hormone-releasing hormone (LHRH) and GAP peptides (49).

Processing o f Vaccinia-Derived Proopiomelanocortin Insulinoma Secretory Granule Type H Activity in Vitro The efficient expression of bona fide prohormone in VV : mPOMC-infected BSC-40 cells (22 pmol of secreted ACTH IR/10 6 cells) affords the ability to prepare significant quantities of POMC to be used as a substrate for processing reactions in vitro. Depending on the requirements, the prohormone can be prepared as a nonradiolabeled [approximately 40/zg of mPOMC can be prepared from 1 x 108 cells (5 • 150-mm plates)] or a radiolabeled [3 x 10 6 cpm are incorporated into mPOMC (4000 cpm/ng)] substrate. Methods describing the production of each have been described (16). To compare cleavage site specificity of the insulin secretory granule type II endopeptidase in vitro with that of the endogenous endoproteases in insulinoma (8) (Fig. 3) and bovine adrenal medullary (BAM) cells (23) (Fig. 3), as well as transfected PC2 and PC3 (9), 10/~g of nonradiolabeled mPOMC was incubated (30~ for 3.5 hr) with an aqueous-phase Triton X-114 extract of insulin secretory granules prepared from an insulinoma propagated in NEDH (New England Deaconess Hospital strain) rats (16). This preparation is highly enriched for the type II (PC2) activity, with lesser amounts of the type I (PC3) activity (---10-30%). The digested sample was resolved by reversed-phase HPLC and the resultant fractions assayed with domain-specific RIAs as previously described (8, 9, 23, 45) [see, e.g., Fig. 4C; Note, compare with the profile of intact POMC shown in Fig. 4A (POMC/WT)]. Analysis showed that the processing in vitro of POMC by the insulinoma PC2 preparation resulted in the excision of a set of peptides highly similar to that produced by Rin m5F cells expressing mPOMC and BSC-40 cells expressing POMC, PC2, and PC3 [compare with Fig. 3 and 4A (POMC/PC3 + PC2)]. Briefly, the major fl-LPH domain peptides identified were 7-LPH and a 7-LPH cleavage product (cleavage at the Leu~~ ~ bond in 7-LPH) as well as native and oxidized fl-endorphin

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(1-31). Thus, like the endogenous Rin m5F cell endoproteases (8) and transfected PC2 (11, 24), the type II endopeptidase efficiently cleaves the Lysl63-Arg 164site at the ACTH//3-LPH and LysZ~ TM y-LPH//3-endorphin junctions but not the LysZ32-Lys233 at the C terminus of/3-endorphin. Prominent ACTH domain peptides included c~-MSH, CLIP, ACTH, and the ACTH/y-LPH processing intermediate. Thus, like PC2 and PC3 expressed in transfected BSC-40 cells (9), the type II activity correctly cleaves at the LyslZZ-Arg 123 site between J peptide/ACTH junction (Fig. 1). Furthermore, like PC2 expressed in transfected cells (9), the type II endopeptidase excised the ACTH/y-LPH processing intermediate from mPOMC.

Acknowledgments The authors thank Dr. Steve Arch for critical reading of the manuscript. This work was supported by NIH Grants DK-44629 and DK-37274. J.V. is supported by NIH Neuroendocrinology Training Grant DK-07680.

References 1. C. Flexner and B. Moss, Annu. Rev. Immunol. 5, 305 (1987). 2. D. E. Hruby, Clin. Microbiol. Rev. 3, 153 (1990). 3 R. J. Leonard, A. Karschin, S. Jayashree-Aiyar, N. Davidson, M. A. Tanouye, L. Thomas, G. Thomas, and H. A. Lester, Proc. Natl. Acad. Sci. U.S.A. 86, 7629 (1989). 4. F. Quan, L. Thomas, and M. A. Forte, Proc. Natl. Acad. Sci. U.S.A. 88, 1898 (1991). 5. A. Karschin, B. Y. Ho, C. Labarca, O. Elroy-Stein, B. Moss, N. Davison, and H. A. Lester, Proc. Natl. Acad. Sci. U.S.A. 88, 5694 (1991). 6. M. Chinkers and E. M. Wilson, J. Biol. Chem. 267, 18589 (1992). 7. R. H. Edwards, M. J. Selby, W. C. Mobley, S. L. Weinrich, D. E. Hruby, and W. J. Rutter, Mol. Cell. Biol. 8, 2456 (1988). 8. B. A. Thorne, L. W. Caton, and G. Thomas, J. Biol. Chem. 264, 3545 (1989). 9. L. Thomas, R. Leduc, B. A. Thorne, S. P. Smeekens, D. F. Steiner, and G. Thomas, Proc. Natl. Acad. Sci. U.S.A. 88, 5297 (1991). 10. P. A. Bresnahan, R. Leduc, L. Thomas, J. Thorner, H. L. Gibson, A. J. Brake, P. J. Barr, and G. Thomas, J. Cell Biol. 111, 2851 (1990). 11. S. Benjannet, N. Rondeau, R. Day, M. Chr6tien, and N. G. Seidah, Proc. Natl. Acad. Sci. U.S.A. 88, 3564 (1991). 12. G. Thomas, B. A. Thorne, and D. E. Hruby, Annu. Rev. Physiol. 50, 323 (1988). 13. K. I. Andreasson, W. W. Tam, T. O. Feurst, B. Moss, and Y. P. Loh, FEBS Lett. 248, 43 (1989). 14. I. Lindberg and G. Thomas, Endocrinology (Baltimore) 126, 480 (1990).

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I MOLECULAR APPROACHES 15. S. S. Molloy, P. A. Bresnahan, S. H. Leppla, K. R. Klimpel, and G. Thomas, J. Biol. Chem. 267, 16396 (1992). 16. C. J. Rhodes, B. A. Thorne, B. Lincoln, E. Nielsen, J. C. Hutton, and G. Thomas, J. Biol. Chem. 268, 4267 (1993). 17. S. Hallenberger, V. Bosch, H. Angliker, E. Shaw, H.-D. Klenk, and W. Garten, Nature (London) 360, 358 (1992). 18. G. Thomas, B. A. Thorne, L. Thomas, R. G. Allen, D. E. Hruby, R. Fuller, and J. Thorner, Science 241, 226 (1988). 19. L. Thomas, A. Cooper, H. Bussey, and G. Thomas, J. Biol. Chem. 265, 10821 (1990). 20. S. P. Smeekens, A. G. Montag, G. Thomas, C. Albiges-Rizo, R. Carroll, M. Benig, L. A. Phillips, S. Martin, S. Ohagi, P. Gardner, H. H. Swift, and D. F. Steiner, Proc. Natl. Acad. Sci. U.S.A. 89, 8822 (1992). 21. E. B. Stephens, R. W. Compans, P. Earl, and B. Moss, EMBO J. 5, 237 (1986). 22. G. Seethaler, M. Chaminade, R. Vlasak, M. Ericsson, G. Griffiths, O. Toffoletto, J. Rossier, H. G. Stunnenberg, and G. Kreil, J. Cell Biol. 114, 1125 (1991). 23. B. A. Thorne, O. H. Viveros, and G. Thomas, J. Biol. Chem. 266, 13607 (1991). 24. G. L. Smith and B. Moss, Gene 25, 21 (1983). 25. G. Thomas, L. Thomas, B. A. Thorne, and E. Herbert, Biochimie 70, 89 (1988). 26. G. Thomas, E. Herbert, and D. E. Hruby, Science 232, 1641 (1986). 27. B. Moss, in "Virology" (B. N. Fields and D. M. Knipe, eds.), Vol. 2, p. 2079. Raven Press, New York, 1990. 28. S. J. Geobel, G. P. Johnson, M. E. Perkus, S. W. Davis, J. P. Winslow, and E. Paoletti, Virology 179, 247 (1990). 29. B. Moss, Annu. Rev. Biochem. 59, 661 (1990). 30. T. Fuerst R. E. G. Niles, F. W. Studier, and B. Moss, Proc. Natl. Acad. Sci. U.S.A. 83, 8122 (1986). 31. O. Elroy-Stein, T. R. Fuerst, and B. Moss, Proc. Natl. Acad. Sci. U.S.A. 86, 6126 (1989). 32. T. R. Fuerst, M. P. Fernandez, and B. Moss, Proc. Natl. Acad. Sci. U.S.A. 86, 2549 (1989). 33. W. A. Alexander, B. Moss, and T. R. Fuerst, J. Virol. 66, 2934 (1992). 34. J. S. Hayflick, W. J. Wolfgang, M. A. Forte, and G. Thomas, J. Neurosci. 12, 705 (1992). 35. P. L. Earl, N. Cooper, and B. Moss, Curr. Prot. Mol. Biol. Supplement 15, p. 16.15.1 (1991). 36. M. Mackett, G. L. Smith, and B. Moss, J. Virol. 49, 857 (1984). 37. H. Shida, T. Tochikura, T. Sato, T. Konno, K. Hirayoshi, M. Seki, Y. Ito, M. Hatanaka, Y. Hinuma, M. Sugimotor, F. Takahashi-Nishimaki, T. Maruyama, K. Miki, K. Suzuki, M. Morita, H. Sashiyama, and M. Hayami, EMBO J. 6, 3379 (1987). 38. P. L. Feigner, T. R. Gadek, M. Holm, R. Roman, H. W. Chan, M. Wenz, J. P. Northrop, G. M. Ringold, and M. Danielsen, Proc. Natl. Acad. Sci. U.S.A. 84, 7413 (1987). 39. C. A. Franke, C. M. Rice, J. H. Strauss, and D. E. Hruby, Mol. Cell. Biol. 5, 1918 (1985). 40. F. G. Falkner and B. Moss, J. Virol. 62, 1849 (1988).

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41. S. Chakrabarti, K. Brechling, and B. Moss, Mol. Cell. Biol. 5, 3403 (1985). 42. J. H. Richardson and W. E. Barkley, "Biosafety in Microbiological and Biomedical Laboratories." Health and Human Services Publication (NIH) 88-8395 (1988). 43. J. Sambrook, E. F. Fritsch, and T. Maniatis, "Molecular Cloning: A Laboratory Manual," Vols. 1-3. "Cold Spring Harbor Lab., Painview, NY, 1989. 44. D. R. O'Reilly, L. K. Miller, and V. A. Luckow, "Baculovirus Expression Vectors: A Laboratory Manual," Vol. 1, p. 161. Freeman, New York, 1992. 45. B. A. Thorne and G. Thomas, J. Biol. Chem. 265, 8436 (1990). 46. D. Julius, A. Brake, L. Blair, R. Kunisawa, and J. Thorner, Cell (Cambridge, Mass.) 1075 (1984). 47. D. F. Steiner, S. P. Smeekens, S. Ohagi, and S. J. Chan, J. Biol. Chem. 267, 23435 (1992). 48. P. J. Barr, Cell (Cambridge, Mass.) 66, 1 (1991). 49. W. C. Wetsel, L. Thomas, J. S. Hayflick, H. R. Rivera, N. Lautermilch, and G. Thomas, Endocr. Soc. Abstr. 74, 453 (1992).

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Overexpression of Neuropeptide Precursors and Processing Enzymes Iris Lindberg and Yi Zhou Although the existence of neuropeptide precursors has been recognized for over 10 years, few neuropeptide precursors have as yet been overexpressed. The overexpression of both neuropeptide precursors and their processing enzymes represent important endeavors for several reasons. These studies will allow the production of enough material for protein crystallization. Obtaining sufficient quantities of different neuropeptide precursors and the common processing enzymes will also permit specificity studies using a variety of precursors. Both of these efforts should ultimately provide the information needed to address the general principles underlying differential processing of neuropeptide precursors by a restricted number of processing enzymes. Overexpression of recombinant proteins in a stable mammalian system has many advantages over other methods of overexpression. Mammalian cells contain enzymes capable of performing appropriate posttranslational modifications such as glycosylation and phosphorylation; disulfide bond formation is also correct. These properties mean that the expressed protein is likely to possess biological activity. We have chosen the Chinese hamster ovary/dihydrofolate reductase (CHO/DHFR) system (1) in order to supply milligram quantities of recombinant secreted proteins in a soluble form. One of the advantages of this system is the ability of these eukaryotic cells to make appropriate posttranslational modifications. The final cell line produced represents a stable system, ensuring that multiple harvests can be obtained from the same roller bottles (which provides cost effectiveness). In addition, supertransfection using other selectable markers (such as G418) is feasible using this method; it is thus possible to add other proteins to the cell line already overexpressing one protein. Another advantage of the CHO/DHFR system is that the final result is a series of cell lines each expressing higher levels of the desired protein; the effect of overexpression dosage can thus be explored. A disadvantage of the DHFR method is the length of time required to obtain the overexpressing cell lines, usually no less than 6 months. The principle of the method revolves around the requirement of all cells for the enzyme dihydrofolate reductase for nucleoside synthesis; when this enzyme is lacking, cells must be growth in medium containing nucleosides. Transfection of a CHO cell line lacking the endogenous DHFR gene with a mixture of cDNAs encoding DHFR and the desired protein results in cointegration in the genome of these two cDNAs. The addition of the drug methotrexate (MTX), a tightly binding inhibitor of DHFR, depletes the cell

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of active enzyme. Amplification of the transfected DHFR, together with the cotransfected cDNA, then ensues in a subpopulation of cells, resulting in the generation of clones expressing higher levels of DHFR and of the desired cDNA. Progressive introduction of these new cell lines to higher levels of MTX forces overexpression to its maximum, reached when the transport of MTX across the cell membrane is limiting. The method described below is adapted from procedures described in Ausubel et al. (2); further information can also be found in Kaufman (3). Overexpression and characterization of recombinant proenkephalin and the prohormone convertase PC1 have been published (4, 5).

T r a n s f e c t i o n of Cells Before beginning the procedure, nucleoside-free fetal bovine serum must be prepared. This serum is obtained by thorough dialysis, which is extremely important for the success of the procedure (L. Chasin, personal communication, 1994). The 500-ml bottle of serum [ICN (Costa Mesa, CA) or other source] is first treated at 60~ for 20 min. We then dialyze it against three changes of 10 liters each of Dulbecco's phosphate-buffered saline [made from GIBCO (Grand Island, NY) stock powder; no calcium added] at 4~ Buffer changes are made at 8- to 16-hr intervals as convenient, and penicillin-streptomycin is included at standard concentrations in the last dialysis buffer change in order to guard against microbial growth. The serum is sterilized by filtration and stored frozen at -20~ in 45-ml aliquots. All glassware is washed by rinsing; no detergents are used. Chinese hamster ovary cells that have the endogenous gene for DHFR deleted (DG44) (6) are obtained from L. Chasin (Department of Biological Sciences, Columbia University, New York, NY). The DHFR vector can be obtained from the Genetics Institute (Cambridge, MA). Both of these sources retain rights to the expressed materials. DG44 cells are grown in Ham's F12 medium with 10% (v/v) fetal calf serum at 37~ in an atmosphere of 5% CO2. Split 2-5 x 10 6 cells into 10-cm dishes the day before transfection; cells should be about 40% confluent with plenty of space between them. For maximum amplification leverage, the ratio of the plasmid bearing the cDNA to be overexpressed to the DHFR-containing plasmid should be high. We find that ratios much greater than 100:1 result in few clones and ratios less than 50"1 result in hundreds of clones. It should be borne in mind, however, that having few clones is not necessarily bad. With the proenkephalin overexpression, two parallel transfections using different amounts of the DHFR vector yielded 1 clone in one transfection and about 100 in the other. After screening by radioimmunoassay, it was found that the single clone had

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a higher initial expression than any one clone obtained from the other transfection. Transfections should be carried out using the calcium phosphate technique, following established methods (3, 4), but using 5-20/zg/10-cm plate of the plasmid bearing the cDNA of interest (which must represent a eukaryotic expression plasmid, purified using a Qiagen kit (Chatsworth, CA), cotransfected with 0.1-0.5/zg of DHFR-coding plasmid. We obtained PC1 and PC2 expression plasmids from N. Seidah (IRCM, Montreal, Canada) (7), and constructed proenkephalin-containing expression vectors from plasmids and inserts provided by others (4). Two days after transfection, cells are selected for the expression of DHFR by splitting into six to ten 10-cm dishes containing Alpha minus minimal essential medium, which lacks nucleosides (GIBCO), containing 10% (v/v) well-dialyzed fetal calf serum (FCS). Two to four T-150 flasks with peelaway tops (Costar, Cambridge, MA) can also be used. DG44 cells should also be split into this medium in order to provide a control for cellular selection; no DG44 cells should survive transfer to nucleoside-free medium. If the cDNA vector encoding the desired protein also contains the gene for G418 resistance, cells can first be selected in G418 (600/zg/ml; 50% active; GIBCO) and selected clones or pools of clones subsequently placed into Alpha minus medium. While this procedure was followed for the PC 1 amplification with good results, it was not particularly successful for our first PC2 expression attempt. The plates are fed twice weekly with flesh medium until the appearance of clones 10 days to 2 weeks later. Between 15 to 30 large, isolated clones are randomly picked using either glass cloning rings (Bellco, Vineland, NJ) or the agarose-overlay method. The latter procedure involves overlaying the cells with 3-5 ml (for a 10-cm plate; 25 ml for a 150-cm flask) of a fleshly prepared 1:1 mixture of 2% (w/v) agarose (autoclaved and maintained at 50~ and 2x Dulbecco's modified Eagle's medium (DMEM) containing 2 • trypsin-EDTA at room temperature; circled clones are picked through the gel layer with a pipettor and sterile 200-/zl truncated pipette tips. Clones are placed into either 48- or 24-well plates (depending on the number of clones selected) containing 1 or 2 ml of medium per well, respectively. When these wells are about 70% confluent, each well is expanded to a 35mm well (in 6-well plates) or to a T-25. When these are again near confluence, a portion is frozen down [in Alpha minus medium with 10% (v/v) serum and 10% (v/v) dimethyl sulfoxide (DMSO)], a portion is maintained in singlicate, and a 10-cm dish or 35-mm well is set up for screening. Almost any plate arrangement should work as long as precautions are taken to guard against loss of clones by contamination, by maintaining duplicates and/or by immediate freezing. The maintenance of clones prior to obtaining screening results can be extremely time consuming; it is sometimes preferable to freeze the

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clones until all of the screening data are in, and then bring up the selected clone.

Screening If antisera are available that are known to react with the protein, screening of secretion medium by Western blotting is rapid and convenient. The wells/ plates are washed with phosphate-buffered saline (PBS) to remove serum proteins and then placed into an appropriate volume (1 ml/well of a 6-well plate; 6 ml for near-confluent 10-cm dish) of Opti-MEM (GIBCO) containing sterile aprotinin (100/~g/ml; Sigma, St. Louis, MO) and allowed to secrete at 37~ overnight. At this stage allowances can be made for clonal growth variations by scoring cells (heavy, medium, light) and adjusting the volume of Opti-MEM appropriately; minimum volumes that allow covering the bottom of the well must be used. The following morning, the conditioned medium is removed, centrifuged to remove any floating cells, and a 1/10 volume of 10 x Laemmli sample buffer is added. The samples are boiled and subjected to electrophoresis on an appropriate percentage sodium dodecyl sulfate (SDS)-polyacrylamide gel [15% (w/v) was used for proenkephalin, 8.8% for PC1] and Western blotting for analysis of expression of an appropriately sized protein. Another screening method involves radioimmunoassay of conditioned medium; for overexpression of proenkephalin, clones were screened using radioimmunoassay of the overnight-conditioned Opti-MEM for Metenkephalin-Arg-Phe immunoreactivity (this peptide represents the carboxylterminal heptapeptide of proenkephalin). For overexpression of prodynorphin, we used a Leu-enkephalin radioimmunoassay (following digestion of the conditioned medium with trypsin and carboxypeptidase B to release this cryptic peptide). We also used a mixture of antisera directed against the amino and carboxyl termini of this protein in Western blots. If no antisera of the desired protein are available, clones can also be screened by preparing RNA and performing Northern blots. Owing to the length of time required for growth of all of the clones, preparation of RNA, and Northern blotting, the clones are stored frozen until all of the data are obtained. We prepare a 10-cm dish of cells from each clone; when 70-80% confluence is reached, the cells are scraped into a small volume of PBS, pelleted, and RNA is prepared from the pellet using the detergent lysis method (2). The pellet can also be stored at -70~ until RNA preparation is convenient. Twenty micrograms of RNA from each clone is subjected to Northern blotting on formaldehyde gels, using standard procedures (2); the blot is photographed under ultraviolet (UV) light in order to provide a record of the relative amounts of rRNA per lane. Following prehybridization at

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42~ in formamide-containing buffer (2), 1-2 • 10 6 cpm/ml of random-primed labeled cDNA probe is included in the 42~ hybridization, and the blot is then washed and exposed to film. With the PC enzymes, for which no antisera were available until well into the amplification, considerable clonal variation in the amount of RNA expression was observed between clones or cell pools derived from early, but generally not late, amplifications. The highest expressing clone is brought up from the freezer into a T-25 flask. When this flask reaches near confluence, it is trypsinized and split 1:6 into two 10-cm plates containing methotrexate at the next highest concentration (see below); the remainder of the cells is frozen again at this time.

Amplification During amplification, the drug methotrexate (MTX) [Sigma, known as (+)amethopterin], which represents a tightly binding inhibitor of DHFR, is applied to the cells in gradually increasing doses. Cellular survival requires the presence of active DHFR and thus amplification of the DHFR gene and coamplification of the gene of interest occur simultaneously following exposure to MTX. For each amplification series, MTX should be purchased in several 100-mg bottles of the same specific activity. This compound is kept frozen in sterile 5-ml aliquots as a stock solution in Alpha minus medium lacking serum (the solution should be warmed and well mixed prior to sterilization by filtration to ensure that all material is soluble). Because MTX is considered hazardous, we do not weigh out this material but instead dissolve the entire contents of the bottle in 42 ml of medium (5 mM stock). The amplification procedure cannot be performed quickly because cells must be slowly adapted to increasing concentrations of MTX. We split cells 1 96 or 1 98 into a four- to fivefold higher concentration of MTX. The addition of increased MTX generally slows cell growth such that the cells do not overgrow the dishes; however, occasionally the cells express sufficient DHFR such that they grow more rapidly and need to be split again within a few days. Under the proper conditions for DHFR starvation, the cells take on a flat, stretched look and begin dying. The plates are fed twice a week with fresh medium containing MTX at the appropriate concentration. Within 10 days to 2 weeks, clones will begin to form, and 12 of these clones are then grown up and again screened for production of the protein of interest. Often, however, clonal growth does not occur; instead, the cells grow slowly over the entire dish. In this case, the cells are expanded and screened as a pool. Simultaneous amplification of six such pools has been recommended by others (3). Provided that expression of the desired protein has been verified at a previous step, this method usually works well at later steps and saves

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considerable hands-on time in screening clones. However, we always compare conditioned medium obtained from clones at various levels of amplification in order to ensure that increasing expression is being obtained over several concentrations of methotrexate. While occasionally no increase is observed from one step to another, over several stages of amplification increasing expression should be observed; otherwise the final cell pool may exhibit low expression (this was the case for PC2). In this case, it may be possible to perform clonal selection at the stage where increasing expression is lost, with the idea of potentially locating a higher expressing subclone among the cells at a particular amplification level. The following amplification steps (concentrations of methotrexate) are employed; 5 nM, 20 nM, 100 nM, 500 nM, 2.5 tzM, 10 tzM, and 50 tzM. Cells must be growing well before the next level is attempted (i.e., attain near confluence from a 1:8 split in 3-4 days) and this usually takes 3-4 weeks. The entire amplification procedure takes about 6 months. It is extremely important to freeze cells at each level of amplification in order to guard against the loss of time by accidental contamination of the current line. For the proenkephalin overexpression, expression of immunoreactive Met-enkephalin-Arg-Phe increased steadily from about 1 pmol/106 cells at 5 nM MTX to 380 pmol/10 6 cells at 50 tzM MTX. However, we have observed a great variability in the expression of various proteins; PC2 was expressed at about 0.5 ~g/ml, PC1 at 2/~g/ml, and proenkephalin at 30 izg/ml. The reason for this variability is unknown but is potentially related to integration-dependent expression as well as possible toxicity of the expressed protein.

Collection of Conditioned Medium The collection schedule of conditioned medium is important. First, the optimal length of incubation for each harvest needs to be determined. Generally, the longer the cells are incubated, the more protein will be secreted into the medium; however, this does not necessarily result in a higher level of the intact protein or greater amounts of enzymatic activity in the medium. Protein degradation is more apparent during longer incubations, especially for smaller peptide precursors; thus the presence of certain proteinase inhibitors or serum proteins in the medium is helpful. For proenkephalin, we found 2% serum to exhibit a protective effect with regard to degradation (4); however, as described below, we now employ Opti-MEM containing aprotinin because it is easier to purify proenkephalin from this solution. Because medium conditions do not resemble intracellular conditions, not only stability but also enzymatic activity may decrease during prolonged incubations. We find that PC1 activity does not always increase in parallel with increasing

100

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MOLECULAR APPROACHES

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75

/

0

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i

I

,

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,

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FIG. 1 Incubation time course: enzymatic activity of PC1 does not always correlate with incubation time. CHO/PC1 cells were cultured in regular medium containing serum in roller bottles, until 80% confluence was reached; bottles were then washed and incubated in serum-free Opti-MEM (120 ml/roller bottle) for 24 hr. One milliliter of medium was removed from each roller bottle at each time point, centrifuged, and the supernatant kept at 4~ until the last time point was collected. PC1 enzymatic activity was then assayed in all aliquots using the RSKR-AMC fluorogenic substrate in the presence of an enzyme inhibitor cocktail to block nonspecific proteinases. incubation time, suggesting that PC1 is inactivated during prolonged incubation (Fig. 1). Western blotting and radioimmunoassay data indicate, however, that immunoreactive PC 1 continuously increases over the same time period. This apparent inactivation of PC 1 is in large measure due to protein aggregation (Y. Zhou, unpublished results, 1994). Second, one needs to determine how many harvests can be carried out using the same flask or roller bottles. Normally, CHO cells can be continuously cultured in serum-free medium from several days to several weeks, depending on the particular cell line and the culture conditions. However, longer culturing in serum-free medium, as compared to regular medium, can sometimes cause cells to detach, possibly due to the rolling movement of the roller bottle. In addition, CHO cells continue to divide slowly in serumfree medium; thus overcrowding may also contribute to cell detachment. Probably because of this cellular detachment and damage, nonspecific proteinase activities (distinguished from PC1 by differential sensitivity to proteinase inhibitors and neutral pH optimum) are increased in PC 1-expressing CHO cell-conditioned medium at later harvests. In contrast, proenkephalinconditioned medium has been successfully harvested from the same roller bottle up to 13 times. Therefore, to achieve the highest yields of the recombinant protein (or activity), a time course experiment such as the one shown in Fig. 1 should be performed to determine the optimal harvest pattern.

[5]

OVEREXPRESSION OF PROTEIN PRECURSORS

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Collection of conditioned medium is performed from roller bottles containing cells at about 80% confluence. CHO cells producing the highest yield of either PC 1 or proenkephalin are cultured in roller bottles with MEM Alpha medium and 10% FBS (MTX is not necessary). To minimize contamination by serum proteins, serum-free medium, Opti-MEM (GIBCO), is used during the collection phase. After two successive washes with 40 ml each of warmed PBS, 100-120 ml of Opti-MEM containing aprotinin (100 txg/ml; Miles Labs, Kanakee, IL) is added. Aprotinin is used to prevent degradation of secreted proenkephalin and prodynorphin; however, this proteinase inhibitor does not appear to be necessary for protection of PC1. To collect PC1, roller bottles are incubated for 12 hr at 37~ to collect proenkephalin, bottles can be incubated up to 24 hr. Increasing degradation of proenkephalin in this medium is seen at 48 hr. These degradation products are extremely difficult to purify away, therefore it is worth preventing their appearance. The freshly collected conditioned medium is centrifuged at 1000 rpm for 15 min to eliminate remaining cells and stored frozen. For proenkephalin, we remove a 1-ml aliquot of each harvest prior to freezing for quality control of harvests by Western blotting; PC1 and nonspecific proteinase activity in each harvest are monitored by performing an enzyme assay in the presence and absence of an inhibitor cocktail (see below). We generally store proenkephalincontaining medium frozen prior to purification. PC 1-containing conditioned medium is also stored frozen; after one freeze-thaw cycle, PC1 activity is decreased 20-40%.

Purification of Secreted Recombinant Protein To obtain secreted recombinant protein from large quantities of conditioned medium, purification should be initiated with a step that affords concomitant concentration. Generally, precipitation methods, ultrafiltration, or certain chromatographical methods are used for this purpose. However, when conditioned medium is concentrated more than 10-fold, protein degradation is dramatically increased, even when procedures are performed at 4~ therefore, adding proteinase inhibitors (such as 1 mM phenylmethylsulfonyl fluoride) is necessary during and after direct concentration of conditioned medium. We have attempted to avoid ultrafiltration because we have observed major degradation of proenkephalin during this procedure. Although they are more time consuming, chromatographic methods appear to be preferable to concentrate both proenkephalin and PC1. The major contaminants in the conditioned medium are residual serum proteins and inactive or degradation fragments of the recombinant proteins. Even when cells are well washed with PBS, trace quantities of serum proteins,

102

I MOLECULARAPPROACHES predominantly albumin and immunoglobulin, still remain bound to the cells. Opti-MEM contains about 50/zg of protein per milliliter (mainly transferrin and insulin), while the concentration of recombinant protein will range between 1 and 30/zg/ml. On the basis of the extent of overexpression achieved, the biochemical properties, and the biological activity of the particular protein overexpressed, specific purification strategies must be devised for each recombinant protein.

Purification of Proenkephalin The highest resolution technique for the purification of recombinant proenkephalin is reversed-phase chromatography. If no serum has been used in the collection of medium, the chromatofocusing step previously employed (4) can be omitted. Using a preparative C4 column (Vydac, 2.2 • 25 cm; Separations Group, Hesperia, CA) it is possible to purify proenkephalin to homogeneity from conditioned medium in one step; substantial, although not complete, separation of glycosylated from unglycosylated proenkephalin is also achieved in this step. This gradient is based on the method of Thomas et al. (8). One hundred and fifty to 200 ml of thawed medium is acidified by the addition of 0.1% (v/v) trifluoroacetic acid and centrifuged for 20 min at 10,000 g. The supernatant is filtered by hand through a 45-/zm Rainin (Ridgefield, NJ) Nytran filter into a 60 ml syringe attached to pump A, and pumped directly onto the HPLC column [equilibrated with buffer A, 0.1% (v/v) trifluoroacetic acid (TFA)] at 6 ml/min. The column is washed with 16% (v/v) buffer B [80% (v/v) acetonitrile in 0.1% (v/v) TFA] until the UV absorbance at 280 nm returns to 0, and a gradient consisting of the following steps is then applied at 4 ml/min: to 35% B in 5 min, to 51% B in 75 min, and to 90% B in 10 min. Fractions of 4 ml are collected and 50-/zl aliquots of each fraction are dried into small Eppendorf tubes, using a Speed-Vac (Savant, Hicksville, NY) and reconstituted in Laemmli sample buffer. Electrophoresis on a 15% polyacrylamide gel followed by Coomassie staining is used to verify the position of proenkephalin (56 min), which depending on the degree of expression may represent the major secreted protein. Phenol red elutes at about 20 min; bovine serum albumin (BSA) elutes at about 69 min; immunoglobulin G (IgG) elutes at 98 min; proenkephalin fragments elute just prior to proenkephalin. The fractions containing proenkephalin are pooled and lyophilized (although variable loss occurs during drying); alternatively, they can be dialyzed against an appropriate buffer. Proenkephalin is very soluble (at least 10 mg/ml in water) but sticky and it is advisable to keep stock solutions at a high protein concentration (> 1 mg/ml). If it is desired to more completely separate glycosylated proenkephalin from

[5]

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unglycosylated proenkephalin, further purification of fractions containing unglycosylated proenkephalin can be performed on a semipreparative C4 column (Vydac, 1 x 25 cm). Pooled fractions are diluted at least threefold with buffer A, 0.2% heptafluoroacetic acid (HFBA; Pierce, Rockford, IL), and applied to the column by repetitive injection through a 5-ml loop. The column is washed with 16% buffer B (100% acetonitrile containing 0.2% HFBA) until the absorbance returns to 0 and is then eluted with a linear gradient to 51% B in 50 min, to 62% in 25 min, 70% in 15 min, and 100% in 10 min. Unglycosylated proenkephalin elutes around 86 min. In both HPLC systems, glycosylated proenkephalin elutes earlier than unglycosylated proenkephalin. Using only the first column, we obtain between 4 and 5 mg of recombinant proenkephalin per preparation. Quality control of the recombinant protein should be performed by amino-terminal sequencing.

Purification of PC1 PC 1, an 87-kDa glycoprotein, has been implicated in the proteolytic maturation of proopiomelanocortin and thus represents a likely candidate for a neuropeptide precursor convertases (9-11). Owing to its size and its enzymatic lability, it is impossible to purify PC1 by reversed-phase HPLC and maintain its biological activity; therefore, FPLC (fast protein liquid chromatography) methods are used. Like HPLC, FPLC provides high resolution, rapid performance, and reproducible conditions; however, because organic solvents are not employed, protein conformation is not altered and enzymatic activity is better maintained during purification. During purification, by optimal arrangement of purification procedures, concentration and desalting of the protein can be accomplished during each chromatographic step, which will minimize time requirements. The entire purification takes less than 12 hr, and enzymatically active PC1 is purified about 19-fold from the conditioned medium to homogeneity (Table I). To prevent degradation and stabilize the protein, all of the procedures are performed at 4~

Enzyme Assay PC 1 activity is monitored by the cleavage of a custom-synthesized fluorogenic substrate; Cbz-Arg-Ser-Lys-Arg-aminomethylcoumarin (RSKR-AMC; Enzyme Systems Products, Dublin, CA). Alternatively, Pyr-Arg-Ser-Lys-ArgAMC (Peptides International, Lexington, KY) can also be used. Similar fluorogenic compounds may also represent appropriate substrates for PC2 (12) and furin (13, 14) respectively. Assays are conveniently carried out in Parafilm-wrapped polypropylene microtiter plates (Costar), but can be

104

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TABLE I Purification of Recombinant PC1 Protein from Conditioned Medium of CHO/PC1 Cells

Step

Enzymatic activity (units) ~

Protein amount (/xg)

Yield (%)

Specific activity (units//zg)

Purification (-fold)

1050 937 743

5180 4860 444

100 89 70

0.19 0.20 1.65

1 1 8.7

56 330

40 90

1.40 3.67

7.4 19

Conditioned medium DEAE-Blue cartridge Phenyl-Superose (I) Mono Q Peak (I) Peak (II) a

5.3 31

One unit is 1 nmol of AMC/hr.

performed in Eppendorf tubes at pH 5.5 with 10 mM calcium chloride and 0.2 mM substrate (final concentrations). Ten-microliter aliquots of each fraction from the purification are included and the final volume is brought up to 50 /zl with distilled water. Conditioned medium samples and harvest-quality control samples are assayed at the same concentrations of substrate and calcium, but are assayed both in the presence and absence of a proteinase inhibitor cocktail of 2.5 I~M trans-epoxysuccinic acid (E-64), 1/xM pepstatin, tosyl lysyl chloromethyl ketone (TLCK) (50/xg/ml), and tosyl phenyl chloromethyl ketone (TPCK) (100 /~g/ml). Activity measured in the absence of the inhibitor cocktail indicates the presence of nonspecific proteinases, and the harvest is not used if high levels (10-20% of total) of such nonspecific proteinases are detected. This inhibitor cocktail is not required in assays following the hydrophobic interaction purification step. The pH of column fractions is adjusted by adding a 1/10 vol of 1 M Bis-Tris buffer, pH 5.5. After 10 to 16 hr of incubation at 37~ cleavage of the peptide bond is detected by measuring the concentration of free AMC, which is highly fluorescent, with a standard spectrofluorometer or a microtiter plate fluorometer (Cambridge Biotechnology, Cambridge, MA) at 380-nm excitation, 460-nm emission. The latter instrument has the advantage that time course experiments can be performed using the same plate incubated for varying lengths of time. Fluorescence is compared to a standard curve of 1-100/zM AMC (Peninsula, Belmont, CA). PC1 immunoreactivity in aliquots of column fractions is analyzed by SDS-PAGE on 8.8% polyacrylamide gels and Western blotting (Mini-PROTEAN system; Bio-Rad, Richmond, CA) using polyclonal PC1 antiserum directed against residues 84-100 (representing the presumed amino terminus of the mature protein) (15).

[5] OVEREXPRESSION OF PROTEIN PRECURSORS

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Concentration by DEAE-Blue Chromatography Three hundred and fifty milliliters of conditioned medium, representing several successive harvests, is recentrifuged at 10,000 rpm for 20 min. The supernatant is then diluted with an equal volume of 20 mM Bis-Tris buffer (pH 6.5) and is loaded onto a 5-ml DEAE-Blue Econo-cartridge column (BioRad) at a flow rate of 2 ml/min. The column is washed with at least 20 ml of the same buffer and is then isocratically eluted with 30 ml of 1 M sodium acetate in 20 mM Bis-Tris, pH 6.5. The effluent is monitored at 280 nm by UV absorbance and a 20-ml peak of UV-absorbing material is collected. During column loading, most of the phenol red and minor contaminant proteins are removed because they do not bind to the column. This relatively inexpensive column concentrates the medium about 18-fold in 7 hr, and prevents the more expensive FPLC columns from becoming contaminated with phenol red. It can be reused several times before discarding (due to flow-through of enzyme).

Purification by Hydrophobic Interaction Chromatography The peak of UV absorbance at 280 nm, which is obtained in 1 M sodium acetate, 20 mM Bis-Tris buffer (pH 6.5), is applied directly to a phenylSuperose column (HR5/5; Pharmacia, Piscataway, NJ) without further manipulation. Elution of proteins is performed using an 18-ml linear gradient from 1 M sodium acetate, 20 mM Bis-Tris, pH 6.5 to 20 mM Bis-Tris, pH 6.5, followed by an 18-min isocratic elution with 20 mM Bis-Tris buffer, pH 6.5. One-milliliter fractions are collected at a flow rate of 0.5 ml/min. A single broad peak of proteinase activity against the fluorogenic substrate is detected, and this peak should overlap with PC1 immunoreactivity. During this chromatographic step, albumin and most serum proteins do not bind to the column under the loading conditions, while inactive PC1, mostly represented by aggregated forms, elutes later than the peak of enzymatically active PC1. Thus by using this hydrophobic interaction column, active PC 1, which consists predominantly of the 87-kDa form, is purified from the major contaminants and from inactive forms (Fig. 2).

Purification by Anion-Exchange Chromatography Enzymatically active fractions obtained from the phenyl-Superose column are pooled, diluted with 2 vol of 20 mM Bis-Tris, pH 6.5 buffer, and applied to a Mono Q column (HR5/5; Pharmacia). Proteins are eluted with a linear gradient from 20 mM Bis-Tris, 0.1% Brij 35, pH 6.5 to 1 M sodium acetate, 20 mM Bis-Tris, 0.1% Brij 35, pH 6.5. The flow rate is 0.5 ml/min and 1-ml fractions are collected. All of the fractions are screened by enzyme assay and Western blotting. On the anion-exchange column, the single active peak

106

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MOLECULAR APPROACHES 0.5 E c

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500

-]4oo~"

0.4

--'

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13~176 3

1/

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/i

c

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10

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Fractions

FIG. 2 Purificationprofile using hydrophobic interaction chromatography. The curve represents UV absorbance and the bars depict enzymatic activity against the fluorogenic substrate.

from the phenyl-Superose column is separated into two peaks (Fig. 3A), both of which overlap with the peak of immunoreactive 87-kDa PC 1. Because the protein in the first active peak is not homogenous (as shown by SDSPAGE analysis and Coomassie staining; Fig. 3B), further analysis and characterization are performed using the second peak of active PC1 (5). We have further purified the second peak by gel-filtration chromatography (Superdex G-200; Pharmacia) and verified that a single UV absorbance peak was found that overlapped with both enzymatic activity and 87-kDa PC1 immunoreactivity (5). Owing to losses of activity, this gel-filtration step is now not routinely employed in the purification. The final enzymatically active fractions from the anion-exchange column are pooled, brought to 10% glycerol, and aliquoted. The aliquots can be stored frozen at -20 or -70~ Spontaneous protein aggregation was observed to occur in unfrozen samples kept at initial protein concentrations, resulting in a diminution of activity.

Conclusions We have found the DHFR-coupled amplification method a reliable and relatively straightforward way to overexpress a variety of different protein precursors. To date, we have used the method to overexpress proenkephalin, prodynorphin, a mutated proenkephalin, PC1, and PC2. Expression levels

[5]

A

OVEREXPRESSION OF PROTEIN PRECURSORS

107

300

0.i0

E C

250 o~" m

0 O0 Cq

200 k D a peak I

peak II

u 0.05 t-

ltso ~ ~

116 kDa--

loo ~ ~i

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~-,<

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0

10

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20

97 kDa--

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FIG. 3 Purification profile using anion-exchange chromatography and SDS-PAGE analysis. (A) The curve represents UV absorbance and the bars depict enzymatic activity against the fluorogenic substrate. (B) SDS-PAGE analysis and Coomassie staining of the applied sample and of peaks I and II.

for these proteins have ranged from 0.5 to 30/xg/ml. Interesting differences have emerged from these various amplifications; for example, we have consistently found prodynorphin more difficult to recover intact than proenkephalin; and PC2, but not PC 1, is secreted as a zymogen rather than as the active enzyme (17). Using the DHFR-coupled amplification method, neuropeptide precursors can often be obtained in yields appropriate for crystallization efforts, while processing enzymes that are constitutively secreted (i.e., do not possess a membrane-spanning domain) can be enzymatically characterized. Further refinements of this overexpression method should lead to better methods of maintaining enzyme activity, because at present the amount of enzymatically active PC1 recovered and the specific activity of PC 1 are low compared to the amount of enzyme protein secreted. With the overexpression of several different neuropeptide precursors, interesting comparisons of processing specificity of the new neuroendocrine convertases will become possible.

Acknowledgments The work reported herein was supported by NIH Grant DA 05084. I. Lindberg was supported by an RCDA from the NIDDK.

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References 1. R. J. Kaufman and P. A. Sharp, J. Mol. Biol. 159, 601 (1982). 2. F.M. Ausubel, R. Brent, R. E. Kingston, D. D. Moore, J. A. Sith, J. G. Seidman, and K. Struhl, eds., "Current Protocols in Molecular Biology." Wiley, New York, 1987. 3. R. J. Kaufman, in "Methods in Enzymology" (D. Goeddel et al., eds.), Vol. 185, p. 537. Academic Press, San Diego, 1990. 4. I. Lindberg, E. Shaw, J. Finley, D. Leone, and P. Deininger, Endocrinology (Baltimore) 128, 1849 (1991). 5. Y. Zhou and I. Lindberg, J. Biol. Chem. 268, 5615 (1993). 6. G. Urlaub, P. J. Mitchell, E. Kas, and L. A. Chasin, Somatic Cell Mol. Genet. 12, 555 (1986). 7. S. Benjannet, T. Reudelhuber, C. Mercure, N. Rondeau, M. Chr6tien, and N. G. Seidah, J. Biol. Chem. 267, 11417 (1992). 8. G. Thomas, E. Herbert, and D. E. Hruby, Science 232, 1641 (1986). 9. S. Benjannet, N. Rondeau, R. Day, M. Chr6tien, and N. G. Seidah, Proc. Natl. Acad. Sci. U.S.A. 88, 3564 (1991). 10. L. Thomas, R. Leduc, B. A. Thorne, S. P. Smeekens, D. F. Steiner, and G. Thomas, Proc. Natl. Acad. Sci. U.S.A. 88, 5297 (1991). 11. B. T. Bloomquist, B. A. Eipper, and R. E. Mains, Mol. Endocrinol. 5, 2014 (1991). 12. I. Lindberg, B. Lincoln, and C. J. Rhodes, Biochem. Biophys. Res. Commun. 183, 1 (1992). 13. S. M. Molloy, P. A. Bresnahan, S. H. Leppla, K. R. Klimpel, and G. Thomas, J. Biol. Chem. 267, 16396 (1992). 14. K. Hatsuzawa, M. Nagahama, S. Takahashi, K. Takada, K. Murakami, and K. Nakayama, J. Biol. Chem. 267, 16094 (1992). 15. O. Vindrola and I. Lindberg, Mol. Endocrinol. 6, 1088 (1992). 16. F. S. Shen, N. G. Seidah, and I. Lindberg, J. Biol. Chem. 268, 24910-24915 (1993).

[6]

Use of Antisense RNA to Block PeptideProcessing Enzyme Expression Richard E. Mains

Introduction Antisense RNA and DNA have become useful tools with which to investigate the function of specific enzymes in peptide processing. Figure 1 depicts a secretory granule and a lysosome in a neuroendocrine cell; even in a cell synthesizing a great deal of a given peptide, such as a pancreatic/3 cell making insulin or an intermediate pituitary cell making melanotropin, most of the protease activity in a crude cell extract is not found in secretory granules. Thus, the goal in antisense technology is to determine which of the endogenous enzymes are actually involved in peptide processing and how those enzymes work. Overexpression of an exogenous candiate endoprotease may result in the appearance or enhancement of a specific cleavage; this, however, would not prove that the candidate processing enzyme normally carries out that cleavage step. In its simplest form, the basic idea of the antisense method is to lower the rate of synthesis of one specific target protein and then to investigate the consequences for peptide processing in a neuroendocrine cell that is synthesizing biologically interesting peptides. The method may involve the introduction into cells of RNA complementary to the normal coding or sense strand of RNA; the RNA can be introduced into the cell directly or made by the cell that is expressing an appropriate DNA vector, and the antisense RNA (which we will call AS-RNA for convenience) will then pair with only the targeted endogenous mRNA (as for PC1 in Fig. 2). Alternatively, a synthetic oligonucleotide (DNA) is introduced into the cell and pairs with the endogenous sense RNA (as for PC2 in Fig. 2). The hope in either approach is that the antisense RNA or DNA will alter expression of only the targeted protein. In the case of peptidylglycine a-amidating monooxygenase (PAM; EC 1.14.17.3), AS-RNA indeed lowered the level of PAM protein in a stable cell line (1). As a comparison, PAM activity was also blocked in cells by a drug treatment that chelated copper ions essential for PAM activity (2) but such a drug treatment could alter other cellular functions as well. Antisense treatment also avoids the problem of inaccessibility of the peptide-processing enzyme; guanidinoethylmercaptosuccinic acid (GEMSA) is a potent and specific inhibitor of carboxypeptidase H (EC 3.4.17.10), but it is ineffective Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

109

110

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secr.____etory granule/trans-Golgi lysosomal proteases

novel

I| C~

CPH

endoproteases N U

C

f-"~PC1 C

N ~J [ ) /

N

l

t

cyt~

FIG. 1 Use of antisense technology to identify processing enzymes. Various cell organelles contain peptide-processing enzymes and other proteases. Prohormone convertases PC1 and PC2 are shown, along with an unidentified convertase called PCX; the Golgi endoprotease furin, which is closely related to PC1 and PC2, is shown; enzymes acting later in the biosynthetic pathway, such as CPH and PAM, are also shown (1, 3, 6-8, 16, 51, 52). from outside the cell (3). Another advantage of the antisense approach is that it can be adapted to block expression of one member of a closely related family of proteins when known drugs cannot distinguish the family members adequately (4, 5). The antisense method provides an answer to the question of what consequences a selective drop in the expression of a particular enzyme has on peptide biosynthesis. Frequently, selective overexpression of the same enzyme can provide additional important data on changes in peptide processing. It is important to consider that other members of a related family of proteins may also be affected by an antisense treatment. For example, because PC1 and PC2 have some stretches of nearly identical nucleotide sequences (6-8), a poorly chosen antisense oligonucleotide might block expression of both PC1 and PC2. Any one chosen treatment might inadvertently block more than one targeted protein, for example, blocking expression of an additional putative prohormone convertase ("PCX" in Figs. 1 and 2) by an antisense method supposedly targeting PC1 or PC2.

[6]

A N T I S E N S E RNA BLOCKING ENZYME EXPRESSION

111

PC1 ~ nuclease Ii11/1111111111111111111111111111 PC1 -AS-RNA

PCX

0 0 ~0 0 ks-~176 ribosome

FIG. 2 Antisense techniques reduces expression of a single mRNA: endogenous mRNAs (arrows indicate 5'-to-3' direction) in the cytosol of a neuroendocrine cell, with the PC1 mRNA paired with antisense RNA (AS-RNA) about to be destroyed by a nuclease, and the PC2 mRNA paired with an antisense oligodeoxynucleotide blocking the ability of ribosomes to translate the PC2 mRNA. The other cellular mRNAs are unaffected by the antisense treatments.

Antisense Method" Mode of Action There are several distinct ways that AS-oligonucleotides and AS-RNA have been shown to lower the level of synthesis of the targeted protein. One mode of action involves blocking synthesis of the targeted protein by binding tightly to the region of the initiation codon and blocking the initiation of protein synthesis (PC2 in Fig. 2). Another mode of action involves the destruction of the endogenous mRNA and a net lowering of the level of the targeted mRNA (PC1 in Fig. 2) (5, 9-13). In theory, antisense methods might also block transcription or interfere with correct splicing of RNA, but the evidence for such actions is still rather limited (5, 9, 10, 12, 14). In the case of ASRNA and simple DNA AS-oligonucleotides, the destruction of the endogenous mRNA is produced by endogenous nucleases that destroy doublestranded RNA and R N A - D N A hybrids (5, 9-11). Even a single cleavage in the mRNA for a targeted protein could be adequate to precipitate the destruction of much of the endogenous mRNA, because the nicked mRNA would

112

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MOLECULARAPPROACHES

have lost its poly(A) tail and would presumably become unstable (13). Certain derivatized oligonucleotides, such as methylphosphonate oligonucleotides, block translation when binding to the initiation codon but do not initiate destruction of the targeted mRNA (10, 12, 15).

Antisense Method: Requirements An antisense approach is more likely to be effective, first, if the targeted protein is rate limiting (5, 9, 10). For targeted enzymes present in excess, dramatic reductions in expression are required before functional consequences can be observed. In the case of peptide-processing enzymes, the prohormone convertases and PAM are clearly rate limiting, because there can be significant amounts of the substrates for these enzymes (high molecular weight precursors; glycine-extended peptides) remaining in tissue extracts (3, 7, 8, 16, 17). By comparison, carboxypeptidase H is probably not rate limiting, because there are usually negligible levels of peptides with COOHterminal basic resides in tissue extracts. Second, a convenient cell or animal system is needed. The system can be primary tissue cultures, cell lines, transgenic animals, or embryos that can be manipulated. In each case, the neuroendocrine target cells must synthesize peptides efficiently enough and in large enough quantity so that rigorous examination of changes in peptide processing is possible. Along with the neuroendocrine tissue, there must be a reliable method of delivering the ASRNA or oligonucleotide to the inside of the cells. These methods usually involve uptake of synthetic oligonucleotides or AS-RNA with a transient disruption of normal peptide biosynthetic processing, or else transient or stable expression of a DNA vector that encodes the AS-RNA. Third, the nucleotide sequence of the targeted protein needs to be known. For the synthetic oligonucleotide approach, a published sequence is all that is needed to choose a short sequence to use as the target. For the AS-RNA method, a cloned fragment of DNA is required, although the cloned fragment need not be the full coding region of the protein, and fragments derived from polymerase chain reaction (PCR) (even containing a few errors) are perfectly suitable. For either approach, the potential for confusion from related members of a family of proteins must be considered carefully (4, 6, 8). Finally, assays are needed to ascertain the success of the antisense treatment, preferably several independent assay methods. Northern analyses of the endogenous mRNA are essential, as is some sort of assay for the protein target. The assays for lowered protein levels can be enzyme assays and Western blot analyses, as in the case of PAM (1). Immunostaining could

[6] ANTISENSE RNA BLOCKING ENZYME EXPRESSION

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also work, but detecting a loss in immunostaining signal for a rate-limiting protein requires high-quality antisera. Clearly, rigorous assays of the changes in peptide processing are needed, with microsequencing and peptide mapping often being crucial to allow full interpretation of the results.

Advantages and Disadvantages of Antisense Oligonucleotide Method The most immediate advantage of this method is that one can test a hypothesis a few days after a nucleotide sequence is established, if the neuroendocrine target tissue and the assay methods are ready. The method can work effectively using primary cultures (5, 18-21), which means that the limited number of stable, differentiated neuroendocrine cell lines is not an impediment. The controls for the method are simple and direct, because the sense oligonucleotide should not show the inhibition seen with the appropriate antisense oligonucleotide; in fact, a single base mismatch in an antisense 15-mer is usually enough to eliminate the effect seen with the correct antisense oligonucleotide (20-22). The fact that a single base mismatch can make an antisense oligonucleotide ineffective can be a serious limitation, however, because it means that one cannot be certain that an antisense oligonucleotide designed against the nucleotide sequence for a peptide-processing enzyme in one species should work in a second species. The most commonly used oligonucleotides are 15-mers, because they are long enough to have the requisite specificity and still enter cells well;larger oligonucleotides have also been used successfully (5, 9, 10, 23). A major problem with the oligonucleotide approach is that even a trial experiment costs hundreds of dollars, because the oligonucleotides must be used at concentrations in the 10-100/~M range (9, 12, 15, 21, 24). Another problem is that the synthetic oligonucleotides are often unstable in culture medium unless serum is excluded or vigorously heat inactivated, which may introduce additional confusing factors (9, 10, 25). The breakdown products from oligonucleotides used at such high levels can also accumulate inside cells and be toxic (12, 26). Various chemically modified oligonucleotides are more stable in the medium and gain entry into cells more easily, but on a molar basis are usually less effective at suppressing synthesis of the targeted protein (12, 15). To add to the potential confusion, there are several examples in which the level of the targeted mRNA increased rather than decreased after treatment with antisense oligonucleotides (27). Owing to limitations in space, this chapter focuses on the vector methods, and readers are referred to excellent reviews on the antisense oligonucleotide methods (5, 12, 15, 21, 28).

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Advantages and Disadvantages of Antisense RNA Method Stable cell lines expressing AS-RNA to PAM and PC1 have been used to examine the role of those enzymes in peptide biosynthesis (1, 6). For other targeted proteins, it is clear that AS-RNA stably expressed from the appropriate vector can effectively lower the level of a protein when the ASoligonucleotide method did not work in the same system (5, 9, 10, 29). One clear advantage of creating stable cell lines or transgenic animals expressing a given AS-RNA is that a wide range of experiments can then be performed on the same cells, allowing much more complete testing of potential artifacts than is possible with transient experiments. Another advantage is that the method tolerates a mismatch of up to 15-20% of the nucleotide sequence (10, 11), which means that using the established nucleotide sequence of a protein in one species to inhibit expression in a related species has a good chance of success (as for PAM and PC1, where the rat clone was used to block expression of the mouse enzymes) (1, 6). The property of tolerating mismatches makes the choice of the nucleotide region to target much more crucial in the case of proteins that are members of a family, such as the PCs. One obvious disadvantage of this method is the need to clone a region of the cDNA for the targeted protein, but given the advent of the polymerase chain reaction, any published nucleotide sequence can be used to create a cDNA fragment that may work. While transient expression of vectors encoding a new protein can work using primary cultures (30, 31), there are not yet any reports of the transient method working to disrupt peptide biosynthesis using AS-RNA. Another problem with the AS-RNA approach is that there may be one or more open reading frames in the AS-RNA and thus the AS-RNA might encode a protein that could have its own side effects (9, 10, 32). In addition, the presence of double-stranded RNA in a cell can set off a number of self-destructive reactions, such as the phosphorylation of factors crucial to protein synthesis, which have nothing to do with the desired ablation of the one targeted protein (5, 9, 10). There are also a number of reports of apparently stable cell lines that have lost the ability to express AS-RNA (5).

Choice of Strategy For antisense oligonucleotides, it is clear that targeting the initiation codon is the best initial guess for what region of the nucleotide sequence to block, because binding an oligonucleotide to the initiation codon can block translation and also precipitate destruction of the mRNA (PC2 in Fig. 2) (9, 10, 23). For AS-RNA, the 5' end of the targeted mRNA is usually unsurpassed

[6]

ANTISENSE RNA BLOCKING ENZYME EXPRESSION Met 1 i 3'-"

s,ro

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I antisense transcript

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5' Northern analyses with RNA Probes

Expression Vector for Antisense RNA

wt

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AS

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fullsize

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FIG. 3 A vector encoding an antisense RNA and a drug resistance gene is diagrammed, along with possible results of Northern analyses with sense and antisense RNA probes. in effectiveness by vectors aimed at other regions of the mRNA (PC1 in Fig. 2) (9, 10, 23, 33). Using a fragment that is shorter than the full-sized endogenous mRNA creates the opportunity that the endogenous mRNA and the added AS-RNA might be distinguishable on the same Northern analysis probed with a cDNA probe (which would detect both the endogenous sense mRNA and the added AS-RNA; the sum of the patterns for sense and antisense RNA probes in Fig. 3) (1, 34-37). Another crucial choice is whether to employ an inducible or a constitutive promoter for expressing the ASRNA. Inducible promoters, such as metallothionein or mouse mammary tumor virus, provide the advantage that the antisense approach can then be used for targeted proteins whose expression is essential for cell viability, but the induction itself will necessitate additional control experiments (1, 6, 20, 38, 39). Constitutive promoters have a corresponding set of advantages and problems (5).

Determining Whether Antisense Method Is Working Clearly, the most direct measures of whether the antisense method is working involve direct measurements of the targeted protein. These can involve Western analyses and enzyme assays and immunostaining. The peptides

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made in the neuroendocrine tissue must also be analyzed rigorously, to determine precisely which step(s) in peptide processing have been disrupted, if any. Northern analyses are essential to be certain that the decrease in the targeted protein is not merely the result of a clonal variant, but rather was indeed caused by expression of the AS-RNA. This is not a trivial worry; before we succeeded in establishing AtT-20 lines stably expressing PAM AS-RNA, we encountered several AtT-20 cell lines with low PAM activity that seemed to be clonal variants (1). Even when the AS-RNA is designed to be different in size from the endogenous mRNA, detecting the AS-RNA in a Northern analysis is frequently difficult. The AS-RNA to PAM was designed to be much smaller than the endogenous PAM mRNA, and the AS-RNA was detected with a cDNA probe in samples from stably transfected AtT-20 cells (1). However, in AtT20 cells expressing AS-RNA to PC1, the PC1 AS-RNA was designed to be the same size as the endogenous PC1 mRNA, and the AS-RNA was not distinguishable using a cDNA probe (6). The literature has several examples of AS-RNAs that were readily detected using cDNA probes (9, 36, 37) and of AS-RNAs that were undetectable using cDNA probes (10, 33, 35, 40). A more sensitive approach is to use a sense strand RNA probe, which will detect only the AS-RNA (Fig. 3), but even that approach does not always visualize the AS-RNA because of its extremely rapid turnover (5, 35). An additional complexity is added by the existence in cells of endogenous "unwindase" activity, which is now recognized to have both unwinding and modifying activity (5, 11). Unwindase activity unwinds double-stranded RNA, which at first would seem to prevent the successful action of ASRNA to bring about the destruction of the endogenous mRNA. It is now clear, however, that the unwindase also modifies the paired RNAs by changing up to half of the A residues into I residues, which are then read as G residues during protein synthesis (5, 11). This means that apparently intact RNA, as judged by Northern analysis, would encode a thoroughly altered protein, because many codons would no longer result in the insertion of the normal amino acid in the protein during synthesis. This is probably an explanation for the finding that the targeted protein level can sometimes be depressed far more than Northern analyses would predict (5, 11). This also suggests that the use of solution hybridization-RNase protection assays instead of Northern analyses would be a more quantitative and reliable means of determining how much unaltered endogenous mRNA remains after treatment with AS-RNA. The unwinding and modifying activity is thought to be important in the block of fibroblast growth factor expression in developing Xenopus oocytes, because the oocytes naturally express an antisense RNA to the fibroblast growth factor RNA at a set time during development

(5, lO).

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The use of antisense RNA to alter targeted protein expression can occasionally be confused by the fact that some tissues express natural RNAs that are made from the strand opposite the one normally expected. Examples include the Xenopus oocyte as cited above, gonadotropin hormone-releasing hormone and certain transcriptional factors (10), and insulin-like growth factor II (41). In addition, the problem mentioned above of open reading frames in the AS-RNA may be important if the protein encoded can interact with the native targeted protein. This concept is still controversial, with data supporting the notion of antisense peptides and proteins interacting with natural proteins (42-44) and data refuting any important interactions (32, 45, 46).

Detailed Methodology for Antisense RNA Blocking Enzyme Expression On the basis of the literature cited above and our experience with PAM and PC1 AS-RNA, the following protocol is recommended as the most likely to yield positive results in initial experiments. First, roughly the 5' half of the mRNA should be used to construct the antisense vector (Fig. 3), with careful selection to avoid long open reading frames. This region can be cut from the full cDNA with the appropriate restriction enzymes or created using published sequences and reverse transcription-polymerase chain reaction, starting with RNA from the tissue richest in the targeted protein. The cDNA is then inserted into the expression vector in the antisense direction. We have had success with both inducible promoters (1, 6) and constitutive promoters (A. Zhou and R. E. Mains, unpublished observations, 1994). For the overexpression of sense RNAs, encoding a functional protein, we have had success with drug selection built into the vector and on a separate plasmid (47-50); for antisense work, the best results have come using vectors that have drug resistance closely linked to the antisense sequence of interest (Fig. 3). The plasmid needs to be purified by banding on a CsC1 gradient before use; other methods of plasmid preparation, such as polyethylene glycol precipitation, have yielded significantly fewer stable transformants. A 60-mm dish containing a monolayer of cells (about 1 mg of cell protein or 3 x 106 cells) is rinsed in protein-free medium for 30-60 min, to remove as much protein as practical, because protein inhibits the transfection process. During this rinse period, 30 txg of the plasmid is added to 350 txl of 4 M ammonium acetate, pH 5, and then precipitated using 1 ml of ethanol. The precipitate is collected using a refrigerated microcentrifuge, and washed with 70% ethanol. The 70% ethanol is removed completely in a tissue culture hood, and the precipitated DNA is allowed to dry for 20-30 min. The DNA

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is then dissolved in 30/zl of protein-free culture medium. We have had good success with the calcium phosphate precipitation method (47), but more recently the lipofection technique with cationized lipids has given better results (31, 48-50). The dissolved DNA is mixed with 30/zl of lipofectin reagent (Bethesda Research Laboratories, Gaithersburg, MD). The proteinfree rinse medium on the cells is discarded and replaced with 3 ml of fresh protein-free medium. The DNA-lipid mixture is then transferred into the dish of cells and the cells allowed to take up the DNA at 37~ for 4-5 hr (longer times can be toxic to the cells). The cells are then removed from the dish with trypsin and plated into several 96-well plates, using 100/xl of regular growth medium per well. The manufacturer has changed the recommended protocol for this procedure several times; however, all the variations seem to work well. After 24 hr, the wells are overlaid with an additional 100/zl of medium containing the selection drug (e.g., G418 at 0.5 mg/ml or hygromycin at 200 U/ml, depending on the vector used). Cultures are then fed daily until nontransfected cells begin to die off, after which the feeding schedule is reduced to twice or even once a week. The amount of medium to remove and replace depends on the rate at which the cells are metabolizing; the goal is to prevent the wells from ever becoming acidic, yet to keep some of the conditioned medium in each well to help the establishment of resistant clones. After 3-4 weeks, colonies become apparent to the naked eye when the plates are viewed from below; the colonies metabolize rapidly enough to change the color of fresh medium after an overnight incubation. At this point, the cells are trypsinized from the original plate and equal aliquots are plated into duplicate 24-well plates; one 24-well plate will be used for the initial biochemical screening (Northern analyses, Western analyses, or enzyme assays) and the other plate serves as the stock of cells for further expansion. Typically three-quarters of the clones survive this initial expansion step. After this initial selection, we tend to keep all lines that grow vigorously and show any promising results at all in the biochemical analyses, because subsequent analyses of well-established lines sometimes differ on which lines are best. We also freeze aliquots of promising clones before going further in the analyses. It takes 1-2 months from the time of transfection to have established several stable cell lines expressing the desired plasmid. About half the cell lines showing good growth in the selection medium can be expected to express the transfected DNA at clearly detectable levels; we have found that the best results invariably come from the first wave of cell lines that show good growth in the drug selection medium. For several cell types (AtT-20, hEK-293, and GH 3) we have found that treating the 96-well and 24-well plates with the lid removed for 30 min and the ultraviolet light in the tissue culture hood on, greatly improves initial

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cell attachment. We have also found that the use of high serum levels in the initial plating (up to 30% serum, heat inactivated at 56~ for 30 min) greatly increases the number of stable cell lines established. The number of 96-well plates to use must be determined empirically for each cell line; the basic goal is to have enough plates so that the probability of a drug-resistant well being clonal is high (perhaps a maximum of 10 drug-resistant wells per 96well plate), yet to have the cell density high enough in each well so that there is an adequate feeder layer for the few individual cells that stably incorporate DNA and eventually grow into stable lines. Another variable to determine for each line is the feeding schedule and the amount of medium to replace at each feeding, which must be adjusted for each transfection. Cross-feeding with conditioned medium from wild-type AtT-20 or GH3 cells can also be helpful. For AtT-20 cells, the six or eight 96-well plates from one transfection are usually fed with a nearly complete change of medium (remove and replace 150-175/zl of the 200/xl of medium) each day for 7-10 days, after which the feeding schedule drops to twice-a-week removal and replacement of 100/zl of medium. After the establishment of stable cell lines, we have often found that subcloning is essential to ensure that a clone is being studied (49), especially for the slowly growing antisense lines. A confluent 60-mm dish of the desired line is trypsinized and suspended in 10 ml of growth medium. Then 50 Izl of the cells is combined with 12.5 ml of growth medium containing the selection drug and one 96-well plate is seeded with 100/zl of the diluted cell jypension per well. The remainder of the diluted suspension is mixed with an additional 10 ml of medium and a second plate is seeded, continuing to eight plates with serially diluted cells. A dense 60-mm dish of wild-type GH 3 or hEK293 cells is then trypsinized and the cells suspended in 80 ml of selection medium; a 100-tzl aliquot of the wild-type cells is added to the serially diluted clone of interest. The wild-type cells provide a feeder layer of morphologically distinguishable cells that die at a rate that usually matches the rate of expansion of the stably transfected clones. Clones of stably transfected cells should be visually identified and marked after 10 days to 2 weeks, and are expanded when dense enough. Sometimes a feeder layer of nontransfected cells has also proved useful for the initial stage of transfection.

Acknowledgments This work was supported by Public Health Service Grants DK-32948, DA-00266, and DA-00097. I thank Drs. Betty Eipper, Brian Bloomquist, Sharon Milgram, and An Zhou for critical reading of the manuscript.

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References

10. ll. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27.

R. E. Mains, B. T. Bloomquist, and B. A. Eipper, Mol. Endocrinol. 5, 187 (1991). R. E. Mains, L. P. Park, and B. A. Eipper, J. Biol. Chem. 261, 11938 (1986). L. D. Fricker, B. Das, R. S. Klein, D. Greene, and Y. K. Jung, NIDA Res. Monogr. 111, 171 (1991). A. J. Baertschi, Y. Audigier, P. M. Lledo, J. M. Israel, J. Bockaert, et al., Mol. Endocrinol. 6, 2257 (1992). R. Baserga and D. T. Denhardt, eds., Ann. N. Y. Acad. Sci. 660, 1 (1992). B. T. Bloomquist, B. A. Eipper, and R. E. Mains, Mol. Endocrinol. 5, 2014 (1991). N. G. Seidah and M. Chr6tien, Trends Endocrinol. Metab. 3, 133 (1992). S. P. Smeekens, Biotechnology 11, 182 (1993). D. A. Melton, ed., "Antisense RNA and DNA." Cold Spring Harbor Lab., Cold Spring Harbor, NY, 1988. J. A. H. Murray and N. Crockett, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 1. Wiley, New York, 1992. B. L. Bass, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 159. Wiley, New York, 1992. J. J. Toulme, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 175. Wiley, New York, 1992. M. Y. Chiang, H. Chan, M. A. Zounes, S. M. Freier, W. F. Lima, et al., J. Biol. Chem. 266, 18162 (1991). C. Robinson-Benion, N. Kamata, andJ. T. Holt, Antisense Res. Dev. 1, 21 (1991). D. M. Tidd, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 227. Wiley, New York, 1992. D. F. Steiner, S. P. Smeekens, S. Ohagi, and S. J. Chan, J. Biol. Chem. 267, 23435 (1992). R. E. Mains, I. M. Dickerson, V. May, D. A. Stoffers, S. N. Perkins, et al., Front. Neuroendocrinol. 11, 52 (1990). W. Gerdes, W. Brysch, K. H. Schlingensiepen, and W. Seifert, NeuroReport 3, 43 (1992). A. Ferreira, J. Niclas, R. D. Vale, G. Banker, and K. S. Kosik, J. Cell Biol. 117, 595 (1992). R. H. Selinfreund, S. W. Barger, M. J. Welsh, and L. J. Van Eldik, J. Cell Biol. 111, 2021 (1990). F.J. Mangiacapra, S. L. Roof, D. Z. Ewton, and J. R. Florini, Mol. Endocrinol. 6, 2038 (1992). L. Neyses, J. Nouskas, and H. Vetter, Biochem. Biophys. Res. Commun. 181, 22 (1991). S. A. Liebhaber, F. Cash, and S. S. Eshleman, J. Mol. Biol. 226, 609 (1992). C. Boiziau and J. J. Toulme, Biochimie 73, 1403 (1991). J. C. Larcher, M. Basseville, J. L. Vayssiere, L. Cordeau-Lossouarn, B. Croizat, et al., Biochem. Biophys. Res. Commun. 185, 915 (1992). A. C. Yu, Y. L. Lee, and L. F. Eng, J. Neurosci. Res. 30, 72 (1991). L. C. Yeoman, Y. J. Danels, and M. J. Lynch, Antisense Res. Dev. 2, 51 (1992).

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28. L. Neckers, L. Whitesell, A. Rosolen, and D. A. Geselowitz, CRC Crit. Rev. Oncogen. 3, 175 (1992). 29. A. Ao, R. P. Erickson, A. Bevilacqua, and J. Karolyi, Antisense Res. Dev. 1, 1 (1991). 30. J. M. Burrin and J. L. Jameson, Mol. Endocrinol. 3, 1643 (1989). 31. R. A. Maurer, BRL Focus 11, 25 (1989). 32. A. N. Eberle, R. Drozdz, J. B. Baumann, and J. Girard, Pept. Res. 2, 213 (1989). 33. M. C. Moroni, M. C. Willingham, and L. Beguinot, J. Biol. Chem. 267, 2714 (1992). 34. M. I. Munir, B. J. F. Rossiter, and C. T. Caskey, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 97. Wiley, New York, 1992. 35. S. R. Rodermel and L. Bogorad, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 121. Wiley, New York, 1992. 36. S. M. Kaiser, P. Laneuville, S. M. Bernier, J. S. Rhim, Kremer, R., et al., J. Biol. Chem. 267, 13623 (1992). 37. M. Kimura, M. Sato, and M. Katsuki, in "Antisense RNA and DNA" (J. A. H. Murray, ed.), p. 109. Wiley, New York, 1992. 38. M. J. Smith and E. V. Prochownik, Blood 79, 2107 (1992). 39. T. Liu, J. G. Williams, and M. Clarke, Mol. Biol. Cell 3, 1403 (1993). 40. H. Yamada, S. Koizumi, M. Kimura, and N. Shimizu, Exp. Cell Res. 184, 90 (1989). 41. E. R. Taylor, E. A. Seleiro, and P. M. Brickell, J. Mol. Endocrinol. 7, 145 (1991). 42. D. W. Pascual and K. L. Bost, Immunol. Invest. 19, 421 (1990). 43. G. Fassina, M. Zamaia, M. Brigham-Burke, and I. M. Chaiken, Biochemistry 28, 8811 (1989). 44. T. S. Elton, L. D. Dion, K. L. Bost, S. Oparil, and J. E. Blalock, Proc. Natl. Acad. Sci. U. S. A. 85, 2518 (1988). 45. A. N. Eberle and M. Huber, J. Recept. Res. 11, 13 (1991). 46. U. B. Rasmussen and R. D. Hesch, Biochem. Biophys. Res. Commun. 149, 930 (1987). 47. I. M. Dickerson, J. E. Dixon, and R. E. Mains, J. Biol. Chem. 262, 13646 (1987). 48. A. Zhou, B. T. Bloomquist, and R. E. Mains, J. Biol. Chem. 268, 1763 (1993). 49. S. L. Milgram, R. C. Johnson, and R. E. Mains, J. Cell Biol. 117, 717 (1992). 50. F. A. Tausk, S. L. Milgram, R. E. Mains, and B. A. Eipper, Mol. Endocrinol. 6, 2185 (1992). 51. M. C. Kiefer, J. E. Tucker, R. Joh, K. E. Landsberg, D. Saltman, et al., D N A Cell Biol. 10, 757 (1991). 52. O. Vindrola and I. Lindberg, Mol. Endocrinol. 6, 1088 (1992).

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Section II

Immunological and Biochemical Approaches to the Study of Peptide-Processing Pathways

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[7]

Combination of High-Performance Liquid Chromatography and Radioimmunoassay for Characterization of Peptide-Processing Pathways A. Ian Smith and Rebecca A. Lew

Introduction The accurate quantitation of often very low levels of biologically important peptides in tissue or body fluid extracts is generally possible only by using immunologically based assay systems. Given that active peptides are generated from higher molecular weight precursors (which can be differentially processed in different tissues) and that peptides with varying biological functions can contain common amino acid sequences, these assays are often complicated by the lack of uniquely specific antisera. Strategies such as twosite immunoassays ([8] in this volume) can often overcome some of these problems; however, these assays are restricted only to well-characterized peptides, take time to develop, and allow the measurement of only one peptide per assay. An alternative approach is to use antisera with broad specificity to assay the fractions generated following the chromatographic separation of cross-reacting components. The separation of polypeptides has traditionally proved difficult as they are complex molecules that vary in size, charge, solubility, and solution conformation. Over the last decade or so the development and application of reversed-phase high-performance liquid chromatography (RP-HPLC) has to a large extent overcome these difficulties. This chapter focuses on the development of strategies that combine RPHPLC and radioimmunoassay (RIA) to identify and characterize peptides in tissue extracts. In addition, examples are given of how these techniques can be applied to characterize peptide-processing pathways and to show that in addition to expression and secretion, the precise pattern of peptide processing can be influenced by physiological regulators.

Reversed-Phase High-Performance Liquid Chromatography In this chapter particular emphasis is placed on the development and application of HPLC strategies. The generation of antibodies and their use in radioimmunoassay is well described in a previous volume in this series (Methods in Neuroscience, Vol. 13, Chapters 20-22) and in this volume ([8 and 19]). Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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Reversed-phase HPLC has become the method of choice for separating biologically active peptides. The separations are achieved following the partitioning of the peptide solute with the hydrophobic stationary phase in an aqueous buffer (solvent A). The peptides are then sequentially eluted, their retention dependent on the overall hydrophobicity of the peptide, by decreasing the aqueous component while increasing the organic (hydrophobic) component (solvent B) of the mobile phase. The widespread use of RP-HPLC for the separation of biologically active peptides reflects not only the stability and, importantly, the predictable retention characteristics of the stationary phases, but also their ability to handle a wide spectrum of polar, nonpolar, large, and small peptide solutes. In addition, the versatility of RP-HPLC arising from its ability to modulate solute retention via selective ion suppression, and via solvophobic and ion pair alterations, allows the separation of even very closely related peptides.

Sample Preparation The preparation of samples prior to injection represents the crucial first step in any HPLC analysis. The goal of the chromatogapher is to achieve high recovery of the solute in an injectable volume, free of organic solvent and particulate material. In the case of biological extracts, samples must also be free from protease activity, lipid, high molecular weight protein, or any other substance(s) that may bind irreversibly to the stationary phase.

Tissue Extraction The efficient extraction of peptides or peptide fragments from flesh or frozen tissues requires the removal of protein and lipid, and the inactivation of proteases. Extractants generally fall into one of two categories: (a) aqueous acid and (b) organic solvent. HC1 at 0.1 M has proved a highly effective extractant in terms of extraction efficiency (typically >90%); however, one disadvantage is that labile modifications can be cleaved during extraction, for example, prolonged exposure can deamidate glutamine and asparagine residues. Acetic acid at 2 M, perhaps not so efficient, is, however, a more gentle extractant but should be used only in conjunction with protease inhibitors. One effective extractant developed by Bennett (1) consists of 5% (v/v) formic acid, 1% (v/v) trifluoroacetic acid (TFA), 1% (w/v) NaC1 in 1 M hydrochloric acid. This cocktail, although harsh, has the advantage of precipitating most cellular proteins and would potently inhibit most proteases. Acidified organic solvent, for example, 80% methanol containing

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0.1 M HC1, gives a clean protease-free supernatant; however, for larger peptides (>2500 Da) recoveries are low (<60%) and frequently inconsistent. Tissues are usually homogenized in 5-10 vol (by weight) of extractant in a glass-Teflon or Polytron (Brinkmann, Westbury, NY) homogenizer (sonication should be avoided as labile peptides or labile modifications to peptides can be damaged). The homogenate is centrifuged (20,000 rpm for 30 min) and the supernatant decanted. Lipid can be removed by repeated (three times) extraction with an equal volume of petroleum ether or dichloromethane.

Prechromatography Cleanup Prior to analytical HPLC, samples generally go through a preliminary cleanup procedure. For synthetic or recombinant peptides this procedure may simply involve the removal of particulate material by microcentrifugation and also perhaps volume reduction and/or removal of organic solvent, which is easily accomplished by lyophilization or centrifugal evaporation. However, a universally applicable and effective method of sample cleanup, particularly for tissue extracts, involves the use of octadecylsilane (ODS, C~s)-based disposable cartridges, the premise being that anything eluting from the cartridge will elute from the analytical column.

Procedure 1. 2. 3. 4. 5.

Wet the column with 2 ml of 80% acetonitrile (ACN)-0.1% (v/v) TFA. Wash the column with 2 ml of 0.1% TFA. Load the sample(aqueous). Wash the column with 2 ml of 0.1% TFA (to remove salts, etc.). Elute peptides with 3 ml of 80% ACN-0.1% TFA.

The eluant is collected and the organic component removed prior to HPLC. This particular method was designed for the Millipore-Waters (Bedford, MA) Sep-Pak cartridges. It is important that the stationary phase not be exposed to air at any stage during the procedure. Methionine-containing peptides often partially oxidize during extraction, and as the relative hydrophobicity differs between methionine and methionine sulfoxide, this can lead to ghosting or split peaks on RP-HPLC. A useful quantitative and qualitative oxidation procedure involves exposure of the peptide or peptide-containing extract to 10-3 M sodium periodate in 50%

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acetic acid for 30 min, thus allowing the quantitative conversion of methionine to its sulfoxide form but not to the sulfone (2).

Analytical Reversed-Phase Columns The widespread use of RP-HPLC has led to a tremendous proliferation of reversed-phase columns. Not all these columns, however, are acceptable for the chromatography of polypeptides. The RP stationary phase, which normally is derivatized microparticulate porous silica, varies not only by the type of derivatization (C2-C22, phenol, CN, etc.) but also by the degree of derivatization (carbon loading), end capping (blocking of underivatized silanol), particle size/shape, and pore size. All these factors dictate the selectivity and retention characteristics of a particular column type. For the chromatography of peptides and small proteins, it is essential that any free silanol groups be end capped or blocked. If this is not the case, then a salt must be included in the mobile phase to prevent any interactions between the peptide solute and the column packing that can lead to peak tailing (3). Generally, C18 columns are most appropriate for the majority of peptide separations, whereas small spherical particles (5-10/xm) of medium pore size (50-100 ,~) will provide an efficient, high-capacity packed bed. This type of column will resolve a wide spectrum of peptides varying in size, charge, and solubility (Fig. 1). There are, however, a number of circumstances under which these types of columns are not necessarily optimal. First, to separate larger polypeptides and small proteins, the wide pore (300 A), short alkyl chain (C4) columns such as the Brownlee (Santa Clara, CA) Aquapore 300 are particularly useful. These columns, especially in a semipreparative mode, are a particularly effective first chromatographic step in purifying a peptide from a complex matrix, for example, where an extract contains a multitude of peptides, small proteins, and protein fragments. Second, another problem can be the successful separation of highly hydrophilic small peptides. These small molecules, which are often only barely retained, if at all, on conventional C~8 columns, can be well resolved on small particle size, high carbon loading (>30%) C18 columns, such as the Ultracarb C~8 (Phenomenex, Torrance, CA). Third, the separation of very hydrophobic peptides can be achieved by substituting CN (~Bondapak CN; Waters Assoc., Milford, MA) for C~8; this lowers the overall hydrophobicity of the stationary phase, thus allowing the separation of small hydrophobic molecules. One criterion of peptide purity is its elution as a single Gaussian peak in two distinct HPLC systems. Changing solvents and ion-pair agents, as discussed below, is useful, as is changing the stationary phase. However,

[7]

129

COMBINATION OF HPLC AND RIA 4

6

AO[

2

.J 5

lb TIME

1'5

2b

2'5

(rain)

FIG. 1 Separation of standard synthetic peptides. Solvent A was 0.08% TFA, solvent B was 0.08% TFA in 70% ACN, and a 20-min linear gradient from 5 to 70% B was used. Peak 1, His-Pro-diketopiperazine; peak 2, TRH; peak 3, TRH-OH; peak 4, GnRH; peak 5, substance P; peak 6, somatostatin; peak 7, CCK-8 (all 2.5/zg, except peaks 1 and 3, which are 10 ~g); 0.2 AUFS at 206 nm.

changing both column type and mobile phase is optimal in confirming the purity of an isolated or synthetic peptide. Finally, the selection of column size is of course largely dependent on the particular application, although in some cases choice may be limited by the output of the high-pressure pumping system. Preparative HPLC columns (500 • 25 mm), used to purify up to 10 g of peptide per injection, normally run at flow rates in excess of 10 ml/min, which is beyond the capacity of most analytical HPLC systems. The modern analytical HPLC system is designed to generate flows of between 100/zl and 10 ml/min, which is sufficient to drive semipreparative columns (25 x 1 cm) in which sample loads may reach 100 mg/injection. The development of microbore column technology has allowed the peptide chemist to analyze very low levels (_<10 ng) of peptides and, perhaps more importantly, recover the solute in a low volume prior to sequence analysis. The columns are very narrow (50 x 1 mm i.d.),

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II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES

TABLE I Commonly Used Reversed-Phase Chromatography Solvent Systems Mobile phase

Advantages

Disadvantage

A: 0.1% HaPO4 B: 0.1% H3PO4-ACN

UV absorbent, good chromatography, simple to prepare

Nonvolatile

A: 0.1 M triethylamine phosphate (TEAP), pH 3.0 B: 0.1 M TEAP in ACN, pH 3.0

Excellent chromatography, good buffering capacity, frequently used in preparative HPLC

Nonvolatile

A: 5% formic acid B: 5% formic acid in methanol

Good chromatography, volatile

Poor UV absorbance

A: 0.08% TFA B: 0.08% TFA in ACN

Good chromatography, very clean, UV transparent, volatile

A: 0.5 M pyridine formate, pH 3.0 B: 0.5 M pyridine formate in 60% propanol

Good chromatography, volatile, good buffering capacity

A: 20 mM ammonium formate, pH 4.0 B: 20 mM ammonium formate in 60% ACN

Particularly useful for seconddimension HPLC, volatile, UV transparent

Ref.

Unpleasant to use, poor UV absorbance

and typical flow rates are between 25 and 100 /zl/min, thus requiring a specialized microbore pumping system.

Mobile Phase Considerations The ideal eluting solvents should be (a) totally volatile, thus after drying collected fractions are compatible with subsequent bioassay and/or sequence analysis, (b) ultraviolet (UV) transparent below 215 nm, to allow the sensitive detection of peptides without aromatic amino acids, and (c) give good resolution with no tailing peaks for a wide range of peptides varying in hydrophobicity and molecular weight. In addition, the mobile phase is generally acidic (pH < 3.0) both to protect the silica column core (soluble above pH 7.0) and to suppress carboxyl group ionization, thus minimizing ionic interactions with the stationary phase. Although over the years a number of different solvent systems have been developed [Table I (4-9)], trifluoroacetic acid (TFA)-acetonitrile (ACN)-based solvent systems are perhaps the most commonly used.

[7] COMBINATION OF HPLC AND RIA

131

We have found the following TFA-ACN gradient system particularly useful for the first-step analysis of peptides over a wide range of hydrophobicity and molecular weights (Fig. 1). Column Flow rate Solvent A Solvent B Gradient Detection Injection volume Chart speed

Nova-Pak ClS (Waters Assoc.) 1 ml/min 0.08% TFA (pH 2.1) 70% ACN-0.08% TFA 3-70% B (2-49% ACN), 20 min 214 nm, 0.5 AUFS 20/~1 (5/zg each peptide) 0.5 cm/min

We routinely add a small amount of acetic acid (0.02%, v/v) to solvent A, to balance the optical density of solvent A with solvent B, thus eliminating to a large extent sloping baselines. In this system small peptides such as His-Pro-diketopiperazine, thyrotropin-releasing hormone (TRH; a tripeptide), and TRH-OH are resolved early in the chromatogram, while larger and more hydrophobic peptides such as gonadotropin-releasing hormone (GnRH), substance P, and somatostatin elute later (Fig. 1). Under circumstances in which the separation of closely related peptides is suboptimal, manipulation of the acetonitrile gradient may prove necessary. One example is a study examining molecular forms of~-endorphin congeners. As the method of detection relied on radioimmunoassay, it was essential to separate and identify all the cross-reacting components in the extract. The similarities in peptide structure warranted the development of a gradient capable of separating all molecular forms (Fig. 2). The conditions are essentially the same as outlined before; however, the gradient, 30-80% B over 30 min, is much shallower than the analytical gradient used in Fig. 1. Acid extracts of ovine pars intermedia were purified by Sep-Pak chromatography and, after drying, the reconstituted (0.08% TFA) extract was subjected to RP-HPLC. Fractions (0.5 ml) were collected, dried, and reconstituted in buffer prior to radioimmunoassay (10). The immunoreactive (IR) a-N-acetylendorphin (NacEP) profile (Fig. 2) contained significant levels of the C-terminally shortened forms NacEP(1-17) (NacyEP) and NacEP(1-16) (NacaEP), as well as the larger molecular forms NacEP(1-27), NacEP(1-26), and NacEP( 1-31).

Applications We have previously shown that the precise pattern of NacEP processing in the ovine pars intermedia may be centrally regulated (10). Following hypothalamopituitary disconnection (HPD), in which the descending neural

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II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES

I a RI

~EP Noc-'~EP

P Nic'~EI

AO0

214Into

.

Ior IBEPI-27

I I Noc-~P,-=7

I Q_

5

n 14.1 (3 o Z n" ,=_

0

I

I0 ELUTION

I

2O

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TIME (rain)

FIG. 2 (a) Separation of synthetic ovine fl-EP-related peptide standards (all 5/zg) on a Nov-Pak C~8column, eluted with a linear gradient of 30-80% solvent B (solvents A and B as in Fig. l) over 30 min at 1 ml/min, monitored at 214 nm (0.5 AUFS). (b) RP-HPLC IR-NacEP profile of extract of sheep pars intermedia.

input to the pars intermedia is surgically ablated, the pattern of processing is altered with NacEP(1-27) becoming the dominant molecular species (Fig. 3B). To determine if these effects reflect the loss of descending dopaminergic neural input to the pars intermedia, two groups of normal sheep were treated with either the dopamine agonist bromocriptine (2.5 mg, bid, sc, 10 days) or the dopamine antagonist haloperidol (1.0 mg, bid, sc, 10 days). Acid extracts of pars intermedia were submitted to RP-HPLC and the IR-NacEP profiles determined on collected fractions (Fig.3). In the control pars intermedia, the RP-HPLC profile showed the major species as NacaEP (30%), NacyEP (30%), and NacEP(1-27) (35%), with NacflEP (5%) a minor component (Fig. 3A).

133

[7] COMBINATION OF HPLC AND RIA 800

800

A

B

AIB~-27 400 E

0

o) n

v

n Ld U 0 Z

800

Ae~ AIB'-27

400

~

I0

20

C

3_0

Ai~,_2~

0

k

0

800

10

20

30

20 (min)

30

D

!

rr'

A~

400

400

0

0

0

I0 20 TIME (m in)

30

Ac'Y

T

0

w~

10

TIME

FIG. 3 Typical RP-HPLC elution profile of a-N-acetylated endorphin (NAcEP) from (A) control, (B) HPD (90 day), (C) haloperidol-treated (1 mg, bd, sc, 10 days), and (D) bromocriptine-treated (2.5 mg, bd, sc, 10 days) sheep pars intermedia. Solvent as in Fig. 2. In the HPD animal (Fig. 3B), the dominant molecular species is NacEP(1-27), representing 65% of the total IR-NacEP as described previously. Chronic treatment with haloperidol produced an IR-NacEP profile similar to that observed in the HPD animals (Fig. 3C), with NacEP(1-27) (80%) becoming the principal molecular component. Although bromocriptine lowers the total IR content, on RP-HPLC the IR-NacEP profile (Fig. 3D) was qualitatively similar to the control group. These data suggest that not only does dopamine play a role in proopiomelanocortin (POMC) gene expression and POMC product secretion in the pars intermedia (11), but dopamine also may be involved in the regulation of POMC processing. Indeed, evidence suggests that, albeit at the level of gene expression, the POMC-processing enzymes PC1 and PC2 in the pars intermedia of the rat are also sensitive to dopamine (12).

Ion Pair Strategies For particularly difficult separations a hydrophobic ion-pair agent can be introduced into the mobile phase (1). At low pH (2-3) any basic amino acids within a peptide sequence will be fully protonated and available to "pair"

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II IMMUNOLOGICALAND BIOCHEMICALAPPROACHES ACTH

OD 206nrn

=

o

s TIME (rain) A(;TH CLIP

OD 206

6

~b TIME (rain)

Flc. 4 Top: RP-HPLC separation of human ACTH and human CLIP (solvents A and B as in Fig. 1) with a linear gradient of 30-50% solvent B over 30 rain at 1 ml/ rain (both 10/~g); 0.2 AUFS at 206 nm. Bottom: RP-HPLC separation of human ACTH and human CLIP (both ]0 ~g). So|vent A was 0.12% HFBA, solvent B was 70% ACN containing 0.12% HFBA, and a 45-60% solvent B linear gradient over 10 rain at 1 ml/min was used; 1.0 AUFS at 206 nm.

with a hydrophobic anion. In this way an element of cation-exchange chromatography is introduced, in which peptides containing basic amino acids are more strongly retained. This can prove particularly useful in enhancing the separation of peptides poorly resolved in more conventional RP-HPLC systems. For example, adrenocorticotropin (ACTH) and ACTH(18-3"~ (CLIP), products of the anterior pituitary and pars intermedia, are particularly difficult to resolve with an ACN-TFA gradient. Even using a shallow gradient, eluting peaks are broad and unsatisfactory for collecting fractions. However, substituting heptafluorobutyric acid (HFBA) for TFA allows the rapid separation of ACTH and CLIP into two well-resolved discrete peaks (Fig. 4). Column Flow rate

/xBondapak (Waters Assoc.) 1 ml/min

[7] COMBINATION OF HPLC AND RIA

Solvent A Solvent B Gradient Detection Injection volume Chart speed

135

0.13% HFBA (pH 2.1) 80% ACN-0.13% HFBA 40-85% B (32-68% ACN) 206 nm, 1.0 AUFS 20/~1 (10/~g each peptide) 0.5 cm/min

We have successfully applied this strategy to characterize the ACTH-like immunoreactivity described in the rat pituitary and brain (Fig. 5). Acid extracts of anterior pituitary, par intermedia, hypothalamus, hippocampus, and cortex were partially purified on Sep-Pak cartridges (described earlier) and subject to RP-HPLC, using HFBA as the acidic ion-pair agent in the mobile phase. Collected fractions were dried and reconstituted in RIA buffer, prior to RIA analysis using a C-terminally directed ACTH antibody. The data show (Fig. 5) that as expected the majority of IR-ACTH in the anterior pituitary eluted as authentic ACTH whereas in the pars intermedia CLIP [ACTH(18-39)] is the dominant molecular form. The small amount of heterogeneity seen likely arises from other posttranslational modifications such as phosphorylation, glycosylation, and C-terminal shortening, which have been previously described in the rat pituitary (13). In the different brain regions, CLIP or modified CLIP seems to be the principal immunoreactive species, and in all the CNS tissues examined ACTH appears only as a minor component.

Size-Exclusion Chromatography As described above, RP-HPLC is now used extensively in the characterization of many polypeptides and peptide fragments. However, the separation and recovery of the longer, more hydrophobic peptides can still pose problems. Although these difficulties to a large extent can be overcome by manipulating the ionic component of the mobile phase and/or changing the column type, as described earlier, the resultant separations are often achieved at the expense of resolution of smaller, subtly modified fragments. Under these circumstances size-exclusion (SE)-HPLC can prove a useful adjunct to RP-HPLC in a peptide purification protocol. Unfortunately a number of the methods described previously for SE-HPLC include the use of strong protein denaturants in the mobile phase to maintain linearity of size-based protein separations. Such denaturants place severe limitations on sensitivity and method of detection as well as subsequent analytical procedures on collected fractions. In this laboratory we have developed an ACN-PO4 buffer-based mobile phase that allows the high recovery of separated products, and, after

136

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I M M U N O L O G I C A L AND B I O C H E M I C A L APPROACHES 2,5

I~s

HYPDTHALAMUS ~-0 hCLIP 1.5

1.0

O.5

2S

2.5

5.O

30

PARS INTERMEDIA ~4.0

35

40

HIPPOCAMPUS 2.0

,.5

1.5

i

l

2.0

1.0

N LO

0.5

.

L

0,n.q tzy '

&5

5.O

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20

25

30

&O

3.0

1.5

2.0

1.0

1.0

~5

m

30

35

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CORTEX

ANTERIOR LOBE 4.0

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3s

L5

40

FRACTION NUMBER

30

[7]

COMBINATION OF HPLC AND RIA

137

drying, collected fractions can be assayed directly (there being no evidence for phosphate interference in our assay systems). Acetonitrile (20%) is a necessary component, preventing both polypeptide aggregation and peak tailing due to nonspecific interactions between the solute and the stationary phase (14). Column Flow rate Solvent Detection Injection volume Chart speed

TSK G2000 SW (Toyo Soda Co., Tokyo, Japan) 0.5 ml/min 20% ACN in 50 mM phosphate (Sorenson, pH 5.8) 214 nm, 0.5 AUFS 20/~1 (10/~g each peptide) 0.5 cm/min

Although a calibration curve constructed with protein molecular weight standards within the column fractionation range (500-60,000) was not completely linear (curving down in the lower molecular mass region), we have been able to achieve good separations of many polypeptides, especially in the 3000-30,000 range. For example, fl-endorphin (fl-EP) and its immediate precursor fl-lipotropin (/~-LPH) (molecular mass 3500 and 11,500 Da, respectively), which are poorly resolved by RP-HPLC (4), are effectively separated by SE-HPLC (Fig. 6). In this example acid extracts of rat anterior pituitary (Fig. 6A) and intermediate lobe (Fig. 6B) were applied to the column and fractions (0.5 min) subsequently assayed for IR-fl-EP. Both tissues exhibited immunoreactive material corresponding in size to fl-EP, this being the only significant immunoreactive form in the rat intermediate lobe extract. The rat anterior pituitary however, contained two peaks of immunoreactivity, the major peak eluting in the position of fl-LPH with a slightly smaller peak migrating with/3-EP. The retention times of synthetic standards (fl-EP) and highly purified standards (/3-LPH) were calculated from the UV recording trace. The fractions were collected into a carrier solution [50/A of 20% (w/v) mannitol containing 100/~g of Polypep, Sigma Chemical Co., Australia] to help prevent peptides of interest from binding to the walls of the fraction collection tubes during the drying procedure.

FIG. 5 C-Terminal ACTH immunoreactivity profiles of extracts of rat pituitary anterior lobe, neurointermediate lobe, hypothalamus, hippocampus, and cortex, chromatographed on a/~Bondapak column. The upper left-hand diagram shows the elution positions of human ACTH and CLIP. Vertical hatching defines the elution time of CLIP, horizontal hatching that of ACTH.

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IMMUNOLOGICAL

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o-eLP.

28 ~9 C

1

o- BEP

|

24-

0 ,4,-

u

0 k,,

20-

D

16.

a. ILl (11

12

C

I

~ m.

8 4 tJ.

,5

2"0 ~5

~

s'5 ~)

45 i

ELUTION TIME (rain) o-.13EP

5.0 C

o

4.0

G

o-B~

3.0

IL

w

2D

I

h0

"~-~,~

~

~2~

~

,~4,~'

ELUTION TIME (rain)

FIG. 6 Elution profiles of IR-fl-EP from extracts of rat anterior pituitary (A) and rat intermediate lobe (B). The retention times of purified ovine/3-LPH and synthetic ovine/3-EP are marked. A TSK G2000 SW column was eluted with 50 mM phosphate, pH 5.8, containing 20% acetonitrile (0.5 ml/min) and monitored at 214 nm (0.5 AUFS).

Summary Individually, both HPLC and RIA are useful techniques for either the qualitative (HPLC) or the quantitative (RIA) analysis of peptides. In combination, however, the two techniques represent an extremely powerful tool for the

[7] COMBINATIONOF HPLC AND RIA

139

precise and sensitive characterization of biologically important peptides in tissue or body fluid extracts. In this chapter our aim has been to provide the reader with some general guidelines on developing appropriate strategies for peptide characterization. We have also included practical examples of what can be achieved using these techniques. Absolute conditions must, of course, be determined empirically, depending on the biochemical nature of the peptide(s) and tissue matrix. However, it is hoped that this chapter may help in establishing a starting point for subsequent method development.

Acknowledgment The authors would like to thank Nicole Casey for preparing this manuscript and Drs. J. R. McDermott and D. J. Autelitano for their scientific contributions. This work was supported by the National Health and Medical Research Council of Australia.

References 1. 2. 3. 4. 5. 6. 7. 8. 9.

lO. 11. 12. 13. 14.

H. P. J. Bennett, J. Chromatogr. 266, 501 (1983). D. G. Smyth, Anal. Biochem. 136, 127 (1984). A. I. Smith and J. R. McDermott, J. Chromatogr. 306, 99 (1984). E. C. Nice and M. J. O'Hare, J. Chromatogr. 162, 401 (1979). N. G. Seidah, R. Routhier, S. Benjannet, N. Lariviere, F. Gossard, and M. Chr6tien, J. Chromatogr. 193, 291 (1980). Y. Takagaki, G. E. Gerber, K. Nihei, and H. G. Khorana, J. Biol. Chem. 225, 1536 (1980). J. R. McDermott, A. I. Smith, J. A. Biggins, M. Chyad A1-Noaemi, and J. A. Edwardson, J. Chromatogr. 222, 371 (1981). G. J. Hughes, C. De Jong, R. W. Rischer, K. W. Winterhalter, and K. J. Wilson, Biochem. J. 199, 61 (1981). A. N. Starratt and M. E. Stevens, in "Handbook of HPLC for the Separation of Amino Acids, Peptides and Proteins" (W. S. Handcock, ed.), Vol. 2, p. 255. CRC Press, Boca Raton, FL, 1985. A. I. Smith, C. A. Wallace, D. J. Autelitano, M. C. Cheng, I. J. Clarke, and J. W. Funder, Neurosc. Lett. 65, 229 (1986). A. I. Smith and J. W. Funder, Endocr. Rev. 9(1), 159 (1988). R. Day, M. K-H. Schafer, S. J. Watson, M. Chr6tien, and N. G. Seidah, Mol. Endocrinol. 6, 485 (1982). H. P. J. Bennett, C. A. Browne, and S. Solomon, Biochemistry 20, 4530 (1981). A. I. Smith, C. A. Wallace, M. C. Cheng, and D. J. Autelitano, J. Chromatogr. 416, 225 (1987).

[8]

Development and Use of Two-Site Immunometric Assays for Examining Peptide-Processing Pathways Steven R. Crosby

The theoretical advantages associated with the use of two-site immunometric assays for the measurement ofpeptide and protein hormones were recognized long ago (1). These advantages include improved sensitivity and precision, wider working range, greater specificity, and shorter incubation times than radioimmunoassays (RIAs), which use antibodies directed against a single epitope. A further advantage of two-site assays is their use for studying peptide processing pathways. By generating specific antibodies to defined amino acid sequences (domains) within peptide precursor molecules, and combining selected pairs of these antibodies, specific two-site assays for individual products of peptide precursor processing from full-length precursor molecules (2) to small peptides such as adrenocorticotropin (ACTH) (3) can be developed. In this way it is possible to establish a panel of assays for the measurement of many different products of precursor processing without the need to separate these peptides by chromatography. Two-site assays therefore provide a practical approach to studies of peptides processing in vivo because they allow the rapid measurement of peptides in small volumes of sample and enable large clinical studies to be undertaken that would otherwise be impractical. The ability to do this may be of diagnostic interest because the appearance of incompletely processed peptides in fluids may indicate the presence of disease. In this chapter, procedures for the development of two-site immunoradiometric assays for the specific measurement of individual products of peptide precursor processing are described. The proopiomelanocortin (POMC) family of peptides released from the anterior lobe of the pituitary gland is used as a model; however, the techniques may be equally applied to other peptide families derived from common precursors.

Principle of Two-Site Immunometric Assay A schematic representation of the two-site immunometric assay, which uses antibodies directed at two distinct epitopes on the molecule to be measured, is given in Fig. 1. Briefly, the sample containing the antigen is mixed with 140

Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

141

[8] TWO-SITE IMMUNOMETRIC ASSAYS

Q O Antigen

4-

FIG. 1

~

Labeled antibody

1

Primary ~ ~ complex ~ ~

~

)_.

4-

Unbound labeled antibody

Secondary complex Principle of the two-site immunometric assay.

excess labeled antibody, which is directed at one of the epitopes, and after a short incubation a labeled antibody-antigen complex (primary complex) is formed. A second antibody (capture antibody), which is directed toward a separate epitope and which is linked to a solid phase (e.g., Sephacryl S300), is then added (again in excess), and after a further short incubation a secondary complex (i.e., labeled antibody-antigen-solid-phase antibody) is formed. The amount of label associated with this complex is directly proportional to the antigen concentration and the measurement of label activity associated with the solid phase after the removal of unreacted labeled antibody enables the accurate estimation of the antigen concentration in the sample. The sequence of incubations with each reagent need not be in the order described, and in some instances it is possible to add labeled and solidphase antibodies to the sample simultaneously. From this brief description, it is clear that most of the antigen in the sample participates in generating a positive signal in a two-site assay because the use of excess antibody favors the formation of antigen-antibody complexes. However, it is also clear that two-site assays use large quantities of antibodies

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

relative to RIAs and the long-term usefulness of two-site assays is therefore dependent on a continuous supply of homogeneous antibody.

D e v e l o p m e n t of T w o - S i t e I m m u n o m e t r i c A s s a y s

Choice of Antibodies: Polyclonal or Monoclonal Although polyclonal antibodies are easier and simpler to produce, monoclonal antibodies (MAbs) are superior reagents when used in two-site assays for reasons associated with the ease of preparation of assay reagents. Using hybridoma techniques, it is possible to generate large quantities of homogeneous, monospecific antibody relatively easily, without the need for immunoadsorption to antigen. The practical problems associated with the preparation of homogeneous antibody for labeling and coupling to solid supports are therefore overcome with MAbs but represent a significant hurdle with polyclonal antisera.

Choice of Antigens A further advantage of using hybridoma techniques is that in order to produce MAbs a pure preparation of immunogen for immunization is not required. A relatively crude form or extract of the peptide may be used as the immunogen and, after fusion, hybridomas can be screened for those producing antibodies to the desired epitope. However, wherever possible, both N- and Cterminal synthetic or purified peptide sequences coupled to larger immunogenic proteins such as bovine serum albumin (BSA) should be used as immunogens because this maximises the chances of generating compatible pairs of MAbs. In general, this approach is successful, but epitopes on native peptides may not necessarily fold into the same conformation as synthetic peptides.

Choice of Assay Format The choice of assay format (i.e., type of label and solid phase) is determined largely by practical considerations including the required degree of assay sensitivity, the proposed usage of the assay (i.e., how frequently the assay will be used), and the instrumentation available. An appropriate label for use in two-site assays has a high specific activity, is stable in solution, is compatible with many sample media, is easily coupled to antibody, and

[8] TWO-SITE IMMUNOMETRIC ASSAYS

143

displays low nonspecific binding to the solid phase. Radioiodine, alkaline phosphatase, and horseradish peroxidase fulfill these criteria and are the most commonly used labels, although many others have been used. For the solid phase, antibodies have been coupled to many different solid supports ranging from microparticles such as Sephacryl S-300 to microtiter plates and membranes. Generally, the stability of antibodies is enhanced when coupled to solid phases and in our experience MAb coupled to Sephacryl S-300 is stable for up to 6 months when stored as a 50% slurry at 4~ Taking these considerations into account for the development of assays for the POMC family of peptides secreted by the adult human anterior pituitary gland, an assay format using 125I-labeled MAbs and Sephacryl S-300linked MAbs as the solid phase has been adopted in our laboratory. For the separation of the secondary complex from unreacted labeled antibody, we have chosen the sucrose layering technique developed by Hunter and Budd (4). This is a noncentrifugation method that regularly gives precision of 1-3% over most of the standard curve.

Generation and Characterization of Antibodies If polyclonal antibodies are to be used, it is advisable to immunize large animals such as sheep or goats to ensure the production of adequate volumes of antiserum. A total dose of 100/~g of immunogen per animal per immunization, using a multisite immunization technique, usually yields good results. Booster injections are given at monthly intervals, and small volumes of blood are withdrawn 7 to 10 days after these injections. When the antibody in serum from these test bleeds has reached a high titer, a large volume of blood is withdrawn and the process is repeated after 3 days. Standard hybridoma techniques (5) are used to generate MAbs. In brief, activated splenic lymphocytes from immunized mice (1-2 x 108 cells) are fused with 107P3-NS1/Ag4 mouse myeloma cells using 45% polyethylene glycol 4000. The resulting suspension is diluted in HAT medium (hypoxanthine-aminopterin-thymidine) containing 20% (v/v) fetal calf serum and distributed into six 96-well plates on feeder layers of nonimmunized spleen cells (1 x 105/well). All wells containing hybrids are screened for antibodies to radiolabeled peptides. Cloning and antibody production in vivo (in ascites fluid) are carried out as described by Goding (6).

Assessment of Antibodies Irrespective of whether polyclonal or monoclonal antibodies are used, they should be characterized for avidity and specificity. The RIA provides a

144

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IMMUNOLOGICAL

AND BIOCHEMICAL

APPROACHES

100 %%

.,

B ~! 0 m

m

1

50

. . . .

C". ',

tr-

~ . . . . . . . . . .

:~ ,

X

!.. '.

%%

.

i

;

Y

Z

.. Oo

Peptide

FIG. 2 Radioimmunoassay displacement curves for the determination of the degree of cross-reactivity that an antibody (recognizing peptide A) has with related peptides (B and C). Cross-reactivity is calculated from the molar concentrations of peptides B and C required to produce 50% displacement of the radioligand [e.g., the percent cross-reactivity of peptide B is (X/Y) x 100].

convenient means of achieving this and the following protocol is used routinely in our laboratory. The assay diluent is 0.1 M sodium phosphate buffer containing 0.25% (w/v) BSA and 0.01 (w/v) sodium azide. Antibody (100/zl) is incubated with 125I-labeled peptide (10,000 cpm in 100/zl) and peptide standards or diluent (100/zl) for 18 hr at 4~ Separation of bound and free radioligand is with anti-IgG bound to cellulose (Sac Cel; Immunodiagnostic Systems, Washington, Tyne and Wear, U.K.) for 30 min at 21~ and antibody titer is expressed as the final dilution of antibody required to bind 50% of the radioligand. Antibodies are also characterized for avidity, and specificity by constructing displacement curves with potentially cross-reacting peptides and the crossreactivity is calculated from the molar concentration of each peptide required to produce 50% displacement of the radioligand (Fig. 2) as follows: % Cross-reactivity concentration of antigen causing 50% displacement of tracer x 100 concentration of cross-reacting peptide causing 50% displacement This approach enables an initial assessment of cross-reactivity to be made and, by constructing similar displacement curves with overlapping synthetic

[8] TWO-SITE IMMUNOMETRIC ASSAYS

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peptide sequences, it is possible to map the antibody-binding site within a peptide accurately to regions of approximately 10 amino acids. Further characterization to determine whether two antibodies can bind simultaneously to the same molecule is achieved after they have been labeled and coupled to the solid phase.

Preparation of Immunoradiometric Assay Reagents In our experience, highly purified antibodies are required for labeling, but for the preparation of solid-phase reagents it is not necessary to have antibodies purified to homogeneity.

Radiolabeled Monoclonal Antibodies Each MAb is purified from ascites fluid. After filtration (0.4/xm, Millex; Millipore, U.K.), 4 ml of ascites fluid is diluted 1" 1 with 1.5 M glycine, 3 M NaC1 buffer (pH 8.9) and passed through a column (5 ml) of Sepharose-protein A (Pharmacia, Milton Keynes, Bucks, U.K.). Pure antibody is eluted with 100 mM citric acid, pH 4, and neutralized with 1 M Tris-HC1, pH 9. Typical yields from 4 ml of ascites fluid should be between 20 and 30 mg of antibody. Purified antibodies are iodinated by the chloramine-T method, which enables the preparation of radiolabeled antibodies with specific activities up to the theoretical optimum of 12/zCi//xg [incorporation of one iodine per immunoglobulin G (IgG) molecule]. To 50/xg (25 ~1) of purified antibody is added an equal volume of 0.5 M sodium phosphate buffer, pH 7.5. The antibody is reacted at room temperature for 50 sec with Na~25I, (0.5 mCi; Amersham International, Bucks, U.K.) and 25/zg of chloramine-T (BDH, Poole, Dorset, U.K.) in 10/~1 of 0.5 M sodium phosphate buffer, pH 7.5. The reaction is stopped with 62.5/xg of L-cysteine hydrochloride (Sigma, Poole, Dorset, U.K.) in 100/xl of 0.25 M Tris buffer pH 8.5, and the volume made up to 1 ml with potassium iodide (10 mg/ml) in Tris buffer. Unreacted iodide is separated on a Sephadex G-25 column (PD10; Pharmacia) eluted with Tris buffer.

Solid Phase-Linked Antibodies Solid-Phase Activation Sephacryl S-300 (Pharmacia) is activated according to the method of Wright and Hunter (7). The gel is oxidized with sodium metaperiodate and stored at 4~ in 0.1 M bicarbonate buffer, pH 9.0, as a 50% slurry.

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Antibody Purification Antibody in ascites fluid is partially purified on a CM Affi-Gel Blue column (5 ml of gel/ml of ascites fluid) (Bio-Rad Laboratories, Richmond, CA) and eluted with 0.01 M KzHPO4, 0.15 M NaC1, pH 7.25. This step effectively removes transferrin and albumin, which are the two other major protein components of ascites fluid. After dialysis against 0.1 M sodium bicarbonate buffer, pH 9.0, the antibody is stored at -70~ until coupled to the solid phase.

Coupling to Solid Phase Purified antibody in 0.1 M bicarbonate buffer, pH 9.0, is mixed overnight at room temperature with activated Sephacryl S-300 (0.5-1.5 mg protein/ml gel), after which unbound, activated sites on the gel are blocked according to the method of Wright and Hunter (7). The gel is stored at 4~ as a 50% slurry in 0.05 M sodium phosphate buffer, pH 7.4, containing 0.1% (w/v) sodium azide, 0.5% (w/v) BSA, and 0.1 (v/v) Triton X-100 (Sigma). Immediately before use, the solid phase is washed three times in assay diluent to remove loosely bound antibody.

Assay Standards The availability of many synthetic peptides has eased many of the standardization problems encountered with purified peptides. Synthetic peptides are usually well-defined reagents, in contrast to many purified preparations that have been shown to be heterogeneous. Nonetheless, in the absence of international reference preparations, some problems persist with the standardization of many peptide immunoassays, particularly for larger precursor peptides that cannot be chemically synthesized (e.g., POMC) and that are being measured for the first time. Under these circumstances the first description may serve as the reference point for subsequent assays and, therefore, when a new assay system is developed an independent standardization technique should be used for comparison. Having identified a suitable assay standard, it is preferable to dilute stock solutions of this standard into a matrix that is similar to the biological fluid (usually plasma or serum) in which the peptide is to be measured. Peptidefree plasma can be easily prepared for use as an appropriate matrix by using solid phase-linked antibody as an immunoadsorbent. The relative amounts of solid phase antibody and the incubation times required to remove all of the peptide completely will depend on the concentration of peptide in the matrix and must therefore be determined empirically.

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Testing Combinations of Antibodies in Two-Site Immunometric Assay For antibodies to be compatible in a two-site assay the epitopes at which they bind must be sufficiently far apart for two antibodies to bind simultaneously. This can be assessed as follows.

Assay Protocol Peptide standards (100/zl) are incubated with radiolabeled antibody (100/zl containing 100,000 cpm) overnight at 4~ after which antibody linked to Sephacryl S-300 (100/zl, 10% slurry) is added and incubated with constant shaking for a further 2 hr at 21~ Assay diluent is 0.05 M sodium phosphate buffer, pH 7.4, containing 0.25% BSA and 0.1% Triton X-100. Peptide bound to both antibodies is separated from unreacted labeled antibody by sucrose layering (4). Briefly, this involves adding 1.0 ml of assay diluent to the reactants and allowing the Sephacryl particles to settle for 2 min. A 10% sucrose solution (3 ml) is then added underneath the incubate through a stainless steel tube (a manifold enables the addition to be performed on 20 tubes simultaneously), after which the solid-phase particles are allowed to settle. After 15 min, the top layer containing unreacted antibody is aspirated and the procedure is repeated before the tubes are counted for 3 min in a y counter. During settling, the solid-phase particles are effectively washed and nonspecific binding of labeled antibody is routinely below 0.05% of total counts added. Compatible pairs of antibodies can be easily identified by an increase in bound radioactivity in response to increasing concentrations of peptides standard and high signal" background ratios at higher concentrations of peptide standard.

Choice of Labeled and Solid-Phase-Linked Antibodies Assay performance can vary markedly depending on which antibody is used as the label and which antibody is used as the solid phase. Having identified a compatible pair of antibodies, therefore, it is advisable to assess both antibodies as the label and solid phase.

Optimization of Immunoradiometric Assay Using Selected Antibodies Using the combinations of antibodies determined as described above, the immunoradiometric assay (IRMA) is optimized with respect to the following variables: (a) solid-phase antibody concentration, (b) labeled antibody concentration, and (c) incubation times.

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25pg

100,000

1O,000

1000

Peptide FIG. 3 The effect of varying the solid-phase antibody concentration on the standard curve of a two-site immunometric assay. Antibody linked to solid phase was used at 5 (...), 10 (--), and 25/zg (---) per tube.

Solid-Phase Concentration Peptide standards and 125I-labeled antibody (100,000 cpm in 100/zl) are incubated for 16 hr at 4~ Antibody linked to solid phase is then added in 5, 10, and 25% slurries (100/zl, corresponding to 5, 10, and 25/zg of antibody per tube). Figure 3 shows the effect of varying the solid-phase antibody concentration on a standard curve of an IRMA. In this example the effect of increasing the concentration of solid-phase antibody above 10% results in increased nonspecific binding of labeled antibody without significantly increasing specific binding at high concentrations of peptide standard. In this case a 10% slurry of solid-phase antibody would be considered to be optimal and would be used in subsequent assays.

Labeled Antibody Concentration Using the selected concentration of solid-phase antibody, the effect of a range of ~25I-labeled antibody concentrations (50,000-250,000 cpm/100/zl) should be assessed. Figure 4 shows the effect of increasing the labelled antibody concentration on an IRMA standard curve. Bound radioactivity increases throughout the standard curve. However, in this case 100,000 cpm/ 100/xl appears to give low detection limits and acceptable count rates at a given concentration of peptide standard, and for this assay 100,000 cpm/100 /zl would be optimal.

[8]

149

TWO-SITE IMMUNOMETRIC ASSAYS ,.._ _ C

100,000

ssS SSSSSS~

B

sS E 10,000 0.. 0

1000

~

Peptide FIG. 4 The effect of varying labeled antibody concentration on the standard curve of an immunometric assay. Radiolabeled antibody was added at 50,000 (. 9.), 100,000 (m), and 250,000 cpm/100/xl (---).

Incubation Time The effect of different incubation times with labeled antibody should be assessed in one-step and two-step protocols. In the one-step assay, standards and optimal concentrations of labeled antibody and solid-phase antibody are incubated (e.g., for 2-4 hr) at 21~ with constant agitation. In the twostep assays various combinations of labeled and solid phase-linked antibody incubation times are investigated. The incubation protocol that results in the lowest detection limit with the greatest signal:background ratio over the widest range of peptide concentrations is regarded as optimal.

Assay Evaluation

Specificity Because peptide precursor processing can lead to a heterogeneous mixture of peptides in which a common epitope may be repeated in a number of different molecular forms, an assessment of assay specificity is essential. Gel-permeation chromatography, followed by IRMA measurements on the chromatography fractions, will provide information on the size of peptide(s) recognized by the assay. In addition, the ability of related peptides to generate a positive signal in the IRMA is assessed over a range of concentrations of

150

II

IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES 100,000

E

1 O,000

1000

, • OMC ~ " Pro-ACTH

C). O

/.."

i

10

. B-LPH

i

1 O0

|

1000

Peptide (pmol/liter)

FIG. 5 Standard curves of a two-site IRMA for the measurement of ACTH precursors (POMC and pro-ACTH), together with curves for ACTH, N-POC, and fl-LPH.

each peptide. Because two-site assays require binding of antibodies to two separate epitopes in order to generate a positive signal, smaller peptides derived from larger peptides will not cross-react in assays designed to measure the larger peptide. This is illustrated in Fig. 5, which shows a standard curve for an IRMA developed in our laboratory to measure ACTH precursors (POMC and pro-ACTH) together with curves for ACTH, N-POC, and/3LPH (the major end products of POMC processing in the human anterior pituitary gland), covering the range of 1-1000 pmol/liter. ACTH, N-POC, and fl-LPH fail to generate a significant positive signal in the assay, even at very high concentrations (i.e., 1000 pmol/liter), while POMC and pro-ACTH cross-react equally (100%).

Interference by Peptides That Will Bind to One or Other of the Antibodies Antigenic sequences within precursor peptides are also present in smaller end products of precursor processing that may be found in plasma and tissue fluids. These peptides will bind to one or other of the antibodies used in the assay, and although this does not give rise to a positive signal, it is possible that high levels of these peptides may interfere with the binding of the precursor. The extent to which such interference occurs should be assessed by coincubating precursor peptide standards with smaller peptides in the

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assay over a range of concentrations. At high concentrations of the smaller peptide, the signal generated by the precursor standard may be reduced, and it is important to establish that this effect does not occur at concentrations of the smaller peptides that may be found under physiological or pathological conditions.

Parallelism It is important to establish that dilutions of samples containing the peptide to be measured give a parallel response to dilutions of the peptide standard used to calibrate the assay. This is achieved by diluting plasma containing high levels of peptide in assay diluent and peptide-free plasma or serum.

Nonspecific Interference Depending on the biological fluid in which the IRMA measurements are to be made, it is possible that components of the fluids may interfere in a nonspecific manner, and it is therefore important that potential nonspecific interference be assessed. This is done by comparing standard curves diluted in peptide-free fluid and diluent buffer. In contrast to many RIAs, MAb-based IRMAs do not generally suffer from nonspecific serum matrix effects. In our experience, standards diluted in peptide-free serum/plasma show only a slight inhibition of response and it is therefore not usually necessary to employ extraction procedures to remove interfering substances. However, the presence of anti-mouse immunoglobulins (or cross-reacting immunoglobulins) in plasma samples may result in a striking false-positive signal with MAb-based IRMAs, owing to bifunctional antibodies binding to both labelled and solid-phase MAbs. The addition of normal mouse serum (NMS) to the labeled MAb (0.5%) overcomes this problem and NMS should therefore be routinely included.

Stability The stability of endogenous peptide should be assessed by allowing aliquots of whole blood or plasma to stand at room temperature (21~ for various lengths of time. Plasma aliquots from each sample are frozen immediately and stored at -70~ prior to assay. In addition, a known amount of peptide should be added to whole blood and plasma from normal human subjects and treated as just described to assess the stability of exogenous peptide.

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V

N1Cll

E6B2

Pro-A C TH r/ I / I / / I / / I / i / / 1 1 / / 1 / / / / / I / / / / / / / / / / / / / / / / / / / / / / / / / II ,y

N1Cll N-POC J.R W////////////////////////A ~

Y N1E4

Y N1Cll

t//.d

A1A12 ACTH B-LPH V///////I//////~ W//I//////////////////////A

Y A1A12

Y

Y

Y

A2A3

N5Cll

E6B2 B-END II//////////A

V _

E2E10

V E6B2

FIG. 6 Binding sites and combinations of MAbs used to develop a panel of twosite immunometric assays for POMC-derived peptides.

Applications Many peptide hormones/growth factors (e.g., ACTH, insulin, insulin-like growth factors, inhibin, and calcitonin) are synthesized as larger precursors that undergo differential enzymatic processing. Heterogeneity of such peptides in biological fluids is therefore a common occurrence and the appearance of incompletely or abnormally processed peptides may provide useful markers of disease. For example, it has become clear that the POMC gene is expressed in many normal tissues other than the anterior pituitary gland and that posttranslational processing is different in these tissues. In pathological conditions such as pituitary tumors and tumors associated with the ectopic ACTH syndrome, many peptides derived from the ACTH precursor (POMC) are secreted in excess, in addition to ACTH. Plasma profiles of ACTH and related peptides in these conditions therefore show considerable heterogeneity. To study this, a panel of two-site assays for POMC-derived peptides has been developed in our laboratory. Figure 6 shows the binding sites and antibody combinations used to develop these assays and Table I lists the detection limits and cross-reactivity with related peptides for each assay. Using these assays, we have measured POMC peptides in plasma from normal subjects and from many patients with ACTH-secreting tumors of

[8]

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TWO-SITE IMMUNOMETRIC ASSAYS

TABLE I Immunometric Assays for Proopiomelanocortin Peptides a Cross-reactivity (%) Peptide POMC ACTH precursors ACTH N-POC /3-LPH B-END

Detection limit (pmol/liter)

POMC

Pro-ACTH

16.0 2.6

100 100

0 100

0 0

0 0

0 0

0 0

1.0 1.0 0.9 1.7

1 9 10 0

<10 6 NT a 0

100 0 0 0

0 100 0 0

0 0 100 0

0 0 0 100

ACTH

N-POC

/3-LPH

/3-END

a Abbreviations: POMC, Proopiomelanocortin; ACTH, adrenocorticotropic hormone; N-POC, N-proopiomelanocortin; B-LPH, B-Liprotrophic hormone; B-END, B-endorphin. b NT, Not tested.

both pituitary and nonpituitary (ectopic) origin. In all of these cases that we have studied to date, ACTH precursors have been found to be elevated in the plasma of patients with ectopic ACTH syndrome due to small cell carcinoma of the lung. Processing of POMC beyond pro-ACTH therefore appears to be less extensive in such tumors in comparison with pituitary tumors, and ACTH precursors provide a useful marker of the ectopic ACTH syndrome. The extent to which altered processing of other peptide precursors occurs in other peptide-secreting tumors is not known, but it is interesting to note that precursor forms of insulin-like growth factor II are secreted by nonislet cell tumors associated with hypoglycemia (8). Perhaps the major limitation of two-site assays for studying processed peptides is that the technique cannot be applied to peptides below approximately 10-15 amino acids. However, in view of the practical advantages offered by two-site immunometric assays for the analysis of peptide processing in vivo, it is expected that these assays will become increasingly applied for this purpose in future.

References 1. G. M. Addison and C. N. Hales, in "Radioimmunoassay Methods" (K. E. Kirkham and W. M. Hunter, eds.), p. 481. Churchill-Livingstone, Edinburgh, 1971. 2. S. R. Crosby, M. K. Stewart, J. G. Ratcliffe, and A. White, J. Clin. Endocrinol. Metab. 67, 1272 (1988). 3. A. White, H. Smith, M. Hoadley, S. Dobson, and J. G. Ratcliffe, Clin. Endocrinol. (Oxford) 26, 41 (1987).

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II IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES 4. 5. 6. 7. 8.

W. M. Hunter and P. S. Budd, J. lmmunol. Methods 45, 255 (1981). G. Kohler and C. Milstein, Nature (London) 256, 495 (1975). J. W. Goding, J. Immunol. 39, 285 (1980). J. F. Wright and W. M. Hunter, J. Immunol. Methods 48, 311 (1982). A. M. Cotterill, J. M. P. Holly, S. C. Davies, V. J. Coulson, P. A. Price, and J. A. H. Wass, J. Endocrinol. 131, 303 (1991).

[9]

Methods for Identification of Neuropeptide-Proces sing Pathways Paul Cohen, Mohamed Rholam, and Hamadi Boussetta

General Considerations It has become increasingly clear that a number of biologically important peptides and proteins involved in a variety of physiological processes are not directly produced in their active forms after translation of the corresponding mRNAs. Indeed, the relevant genes generally encode higher molecular weight forms called precursors, which are then converted into the bioactive elements by a number of posttranslational modifications termed processing. It is obvious that a control may then be exerted on the generation of these peptide and protein messengers via the processing machinery, including those enzymes that are involved in the relevant reactions. Proteolytic activation of the precursors is performed by putative selective endo- and exoproteases (and peptidases). At the present time those enzymes that are participating in vivo in the limited and selective hydrolysis of given peptide bonds are not unambiguously identified. Only a family of subtilisinlike endoproteases related to the KEX2 gene product of Saccharomyces cerevisiae and the "furin" gene have been identified and implicated in the processing of various proforms. In parallel, other workers in the field were able to identify a large set of endopeptidases, with comparable selectivity, which indeed belong to different enzyme families. They were proposed, on the basis of their in vitro properties toward various substrates, as putative propeptide and proprotein converting enzymes but their exact relevance to specific physiological processes has not been, as yet, clearly established (1). Careful analysis of the primary structures of a large set of peptide and protein precursors indicates that a strong selective pressure was exerted during evolution on the nature of the amino acids serving as potential cleavage sites, that is, mainly basic amino acid residues, arginine and lysine arranged as singlets, or preferentially as doublets (Lys-Arg > Arg-Arg > Lys-Lys Arg-Lys). In contrast, a number of features suggested that the structural basis for the recognition of processing sites by the relevant proteolytic enzyme machinery was more complex. Indeed, the following observations were made (2, 3). Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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1. All potential cleavage sites are not processed in vivo. Less than 60% of the dibasic moieties present in precursors are indeed cleaved. Unprocessed basic residues are recovered within the bioactive (or connecting) fragments resulting from proteolytic processing. 2. Examination of the amino acid sequences situated around the dibasic cleavage sites did not reveal the existence of a unique consensus sequence. Only a few preferences for some residues in given positions appeared and might play a role in enzyme/subsite recognition. This indicated that the conservation of basic residues as signals for endoprotease recognition was not correlated with the existence of a single consensus primary sequence. 3. Although both monobasic and dibasic sites are encountered in proforms there appears to be a hierarchy within the amino acid residues that are cleaved. Arginine is more frequently used as a processing site than lysine, and the Lys-Arg doublet represents about 68% of the moieties that are recognized as processing sites in vivo, versus the Arg-Arg (18%), Arg-Lys (5%), and Lys-Lys (9%) arrangements. 4. Predictive methods to analyze the possible secondary structure around the dibasic cleavage sites in a database of prohormones and proproteins (70 precursor sequences) indicated that these processing loci are preferentially situated in, or in the immediate vicinity of, privileged structures constituted by 13 turns (2) or alternatively larger loops (4). Several questions then arose on the mechanisms underlying these important biological events. These include the following: (a) What are the enzymes involved in these processes? (b) What are the enzyme mechanisms that govern recognition of the substrates? and (c) What is the chronology and topology of these processing reactions? Answering these questions implies the identification of processing intermediates and of the cellular and the subcellular localization of processing events. Therefore, on the basis of these considerations, two main experimental strategies were adopted by workers in the field. One classic strategy, mainly based on biochemical grounds, was to detect, purify, isolate, and then characterize endoproteases exhibiting this type of selectivity toward basic residues included in peptide, or protein, substrates. The next step was to attempt the obtention of partial structural information to achieve cDNA cloning and complete determination of enzyme primary structures. This approach has turned out to be tedious and difficult for a number of reasons: (a) because of the inherent properties of these proteases and/or peptidases, which are poorly represented in the producing tissues, and (b) because proper substrates could not be defined unequivocally in all cases. The other approach was based on the knowledge of a processing endoprotease, the product of the S. cerevisiae KEX2 gene, which is involved in the

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maturation of pro-a mating factor and of the prokiller toxin (5). This subtilisinlike endoprotease, whose sequence has been determined (6), was used as a probe to clone by homology-related cDNAs present in endocrine and exocrine tissues of higher organisms. The deduced sequences of the endoproteases related to the "furin" gene, were classified under the generic term of PCEs (prohormone-converting enzymes) or "kexins." In some cases the corresponding gene expression products, that is, the active enzymes, were obtained by using adequate systems (baculovirus or/ and Cos cell expression). Unfortunately, at the present time, these compounds exhibit little enzyme activity in vitro, a major drawback in obtaining quantitative data on their kinetic properties toward well-defined substrates. In any case, the design and use of suitable substrates to monitor the enzymes and to analyze their mode of action has proved useful. Various substrates have been made use of, including fluorogenic derivatives of amino acids or of small peptides; synthetic peptides of various lengths reproducing, or mimicking, precursor sequences around the cleavage sites; and "fulllength" precursors obtained by hemisynthesis or by use of recombinant DNA techniques. To answer the questions relative to the chronology and topology of biosynthetic events the most successful methods were based on the following strategies" (a) identification of biosynthetic intermediates by the combined use of high-performance liquid chromatography (HPLC) and of selective antibodies against well-defined domains of the precursor; and (b) identification of processing events in situ by use of antibodies directed against precursor epitopes that were unmasked after the proper proteolytic reaction had occurred. In the present chapter, we discuss essentially those methods that have proved useful and efficient (a) in the detection and identification of processing intermediates of the precursor biosynthetic pathway; (b) in the detection and characterization of processing enzymes; and (c) in the identification and localization of posttranslational events occurring in the secretory machinery of producing cells.

Identification of P r e c u r s o r - P r o c e s s i n g Intermediates To establish the biosynthetic pathway of a given peptide it is necessary to identify with precision the precursor as well as the fragments that are generated after the action of one, or many, processing enzyme(s). The task can be considerably facilitated by the knowledge of the complete amino acid sequence of the precursor obtained from cDNA cloning. This allows the

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design of adequate immunochemical tools to monitor the resulting fragments after the careful separation by chromatographic and/or electrophoretic methods. In some cases, partial or extensive amino acid sequencing by conventional or micromethods may be needed. This can be illustrated in the case of the biosynthetic pathways of prooxytocin-neurophysin in the corpus luteum and in the hypothalamoneurohypophyseal tract.

Prooxytocin-Neurophysin Maturation The brain nonapeptide oxytocin (OT) and its associated neurophysin (Np) are present in the ovarian corpus luteum of the cow as well as in the sheep and human. They both derive from a common precursor in which the Nterminal hormone sequence is separated from the C-terminal neurophysin domain by a "processing sequence" Glyl~ 12, which is indeed eliminated on complete processing of the precursor (Fig. 1). To detect and identify the possible processing intermediates and putative corresponding processing enzymes, a strategy was developed on the basis of the following principles (7, 8): (a) a highly enriched preparation of granules was made from bovine corpora lutea and used as a source of enzymes; and (b) the fresh tissues were used as a separate source of peptides that were identified by radioimmunoassay (RIA) following HPLC separation with refer-

[ OT L-Gly-Lys-Arg~

Np

Endoprotease

! OT l_GlyTLys.~Arg + [ , : I

I

I I

I

.P',

I

I Carboxypeptidase B-like I OT LGly + Lys + Arg

I Amidating enzyme i OT I.NH 2 FIG. 1 A schematic representation of prooxytocin-neurophysin processing in the corpus luteum. [From Clamagirand (46).]

[9] IDENTIFYINGNEUROPEPTIDE-PROCESSINGPATHWAYS

159

ence to synthetic standards. The following peptides were produced by solidphase synthesis: pro-OT/Np(1-20) (peptide I) and the corresponding N- and C-terminal fragments pro-OT/Np(1-12)(OT-Gly-Lys-Arg ~2)(peptide II), proOT/Np(1-11)(OT-GlyLys 1~) (peptide V), pro-OT/Np(1-10)(OT-Gly) (peptide IV), and pro-OT/Np(13-20)(Ala ~3~ Arg 2~ (peptide III).

Experimental Procedure Granule Preparation Cows, superovulated during the luteal phase by follicle-stimulating hormone (FSH) and prostaglandin F2~ (PGF2~) injections, and bred by two successive artificial inseminations at 12 and 24 hr after estrus, (INRA, Jouy en Josas, France), are used as a source of corpus luteum. Animals are sacrificed 7-8 days after the heat period and ovaries are immediately collected and rapidly transported on ice to the laboratory. Fifteen corpora lutea are dissected and the 32 g of fresh tissues is homogenized in 20 mM Tris-HC1 (pH 7.0), 250 mM sucrose buffer (1 g of tissue per 10 ml of solution). The homogenate is then subjected to differential centrifugation to yield a granule pellet. Secretory granules are then purified by a gradient centrifugation run in 33% (w/v) Percoll-250 mM sucrose at 64,000 g for 10 min. Each 1-ml fraction of the gradient is measured for its refractive index, oxytocin immunoreactivity, and acid phosphatase activity (9). The oxytocin-containing fractions with a refractive index ranging from 1.3506 to 1.3464 (fractions 6 through 16) are pooled and used subsequently.

Enzyme Fractionation A lysate of purified granules (a total of 13.7 mg of proteins) is obtained by osmotic shock followed by three successive freeze/thaw cycles, then submitted to molecular sieve filtration on Sephadex G-150 (65 x 1 cm) in 50 mM ammonium acetate, pH 7.0, 4~ Fractions of 1 ml are analyzed for enzyme activity (see below). Isoelectric focusing is conducted (on a total of 0.487 mg of proteins) according to the classic method of an LKB (Bromma, Sweden) apparatus (110 ml) using a pH 3.5 through 10 gradient (10, 11).

Peptide Synthesis All peptides used in this work are synthesized by the solid-phase method. Purification and physicochemical analysis are as in Nicolas et al. (12). The following compounds are used in this work, either as substrates or as standards.

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Peptide I: pro-OT/Np(1-20), i.e., CysITyrlleGlnAsnCysProLeuGlyGlyl~

Peptide Peptide Peptide Peptide

2~

II: pro-OT/Np(1-12), i.e., C y s l ~ Arg 12 (OT-GlyLysArg) III: pro-OT/Np(13-20), i.e., Ala 13--~ Arg 2~ IV: pro-OT/Np(1-10), i.e., C y s I ~ Gly 1~(OT-Gly) V: pro-OT/Np(1-11), i.e., C y s 1 --~ Lys l~ (OT-GlyLys)

Peptide Isolation and Identification Bovine corpora lutea are first homogenized (0~ then extracted overnight (4~ in 0.1 N HC1 (10 ml/g fresh tissue). After centrifugation (30 min, 12,000 g, 4~ the supernatant is filtered through a Millipore (Bedford, MA) filter (0.45-/~m pore size), then applied to a Sep-Pak C18 cartridge (Waters, Milford, MA) previously washed with methanol, water, then 0.1% trifluoroacetic acid (TFA) successively. After peptide and protein adsorption (extracted from 3 g of fresh tissues) the cartridge is washed with 32 ml of 0.1% (v/v) TFA and peptides are eluted with 4 ml of 50% (v/v) acetonitrile in 0.1% (v/v) TFA. After evaporation, peptides are further analyzed by HPLC, thin-layer chromatography (TLC), and radioimmunoassay. Thin-layer chromatography of the peptides recovered from HPLC is performed on HP-KF silica gel plates (Whatman, Clifton, NJ), using as eluent the upper phase of the mixture [butanol-pyridine-H20-0.1% AcOH, 50:30:110, v/v).

Enzyme Assay Endoprotease and carboxypeptidase B-like activities are monitored by using peptide I substrate and measuring the production of both peptides II and III (endoprotease) and of both peptides IV and V (carboxypeptidase B-like). Routinely, 20/zg of either substrate is incubated with an aliquot of the fraction to be tested (containing, on average, 6/zg of protein) in ammonium acetate (50 mM, pH 7.0) for 24 hr at 37~ After acidification the remaining substrate and the products of reaction are analyzed by HPLC [~Bondapak C18 column eluted isocratically with 20% (v/v) acetonitrile in H20, TFA 0.1%, (v/v), then by a 20-40% (v/v) acetonitrile gradient in the same aqueous TFA solution]. Production of peptides II and III from peptide I is quantified by ultraviolet (UV) absorbance using a D-2000 (Merck, Rahway, NJ) integrator coupled to the HPLC. Endoprotease activity was expressed as quantities of either peptide II or III produced (in micrograms) (Fig. 2).

Neurophysin Isolation and Characterization Cows, superovulated and bred as previously described (7), are used as a source of corpus luteum. Animals are sacrificed at different stages after estrus and ovaries are immediately collected and transported on ice to the

[9]

161

I D E N T I F Y I N G NEUROPEPTIDE-PROCESSING PATHWAYS

Q.

O o O

L._

I

Z~ o.,

IIIL

...:]~o-'

~_

o~

o.1

|n

l

T.,

0

.

.

.

.

.

.

1,,1

"

~'

Jl~ .

9

~g

II •

I 0

T.

~ ~g

~ .

.

.

.

.

30 Retention

..

,.'111

Ill

/

60

0

time, min

FIG. 2 Identification ofprooxytocin-neurophysin(1-20) (peptide I) and its fragments by HPLC: separation and detection at 220 nm: Peptide II, Cys 1 --* ArgO2; peptide III, Ala ]3 ~ Arg2~ peptide V, Cys 1~ Lys 11. Upper trace: Elution positions of the peptides used as references (peptides II, V, III, and I). L o w e r trace: Elution of the fragments generated after exposure of peptide I to pro-OT/Np convertase, the putative processing endoprotease isolated from bovine corpus luteum and neurohypophysis. [From Clamagirand et al. (10).] Copyright 1987 American Chemical Society.

laboratory. Neurophysins, small proteins of Mr -" 10,000, are extracted from corpora lutea as usual (11), in the presence of protease inhibitors [aprotinin (4/zg/ml), pepstatin (1 /zg/ml), and 5 mM phenylmethylsulfonyl fluoride], and obtained hormone free with others proteins of similar molecular weight by two successive molecular sieve filtrations. Proteins (Mr "~10,000) are analyzed by isoelectric focusing (IEF) using the Phast System apparatus (Pharmacia, Uppsala, Sweden) with Phast Gel IEF 4-6.5, as recommended by the manufacturer. The proteins are transferred by diffusion to nitrocellulose membrane (Hybond C; Amersham, UK) at 4~ for 90 min in the presence of 25 mM Tris, 150 mM glycine, pH 8.3. After transfer the membrane is treated with 3% (w/v) bovine globulin-flee albumin (Sigma, St. Louis, MO)

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

in 10 mM phosphate buffer, pH 7.5, containing 130 mM NaC1. It is then incubated with rabbit anti-bovine neurophysin I serum (618-11) prepared in the laboratory. Goat anti-rabbit IgG-alkaline phosphatase conjugate and color development reagents 5-bromo-4-chloro-3-indolylphosphate p-toluidine salt and nitroblue tetrazolium chloride are used to visualize neurophysin-anti-neurophysin complexes (Bio-Rad, Richmond, CA) To characterize prooxytocin-neurophysin, proteins of Mr "10,000 prepared from corpora lutea 2 or 3 days after estrus are subjected to enzyme cleavage by trypsin and by prooxytocin convertase. One microgram of Ntosyl-L-phenylalanine chloromethyl ketone (TPCK)-treated trypsin (Sigma) is incubated with proteins, free of protease inhibitors by previous filtration on Sephadex G-25 and containing 2/xg of immunoreactive neurophysin species, in 100 mM ammonium bicarbonate, pH 8, at 37~ for 60 min. The reaction is stopped by addition of 0.1 N HC1. Immunoreactive oxytocin is measured by RIA and the released peptides are identified by HPLC. Pr0oxytocin convertase is purified by isoelectric focusing as previously described (10). Proteins of Mr "~10,000 containing 9/zg of immunoreactive neurophysin are incubated with the convertase in 100 mM ammonium acetate, pH 7, at 37~ for 16 hr. After acidification, the reaction mixture is analyzed by HPLC. Granulosa Cell Culture Cows, superovulated and bred as previously described (7), are used as a source of granulosa cells. Animals are sacrificed 40 hr after the beginning of estrus. Ovaries are immediately collected, washed with sterile 0.9% (w/v) NaC1, and immersed in cold culture medium before transportation to the laboratory. Nonovulated follicles are used. At first, follicular fluid is removed by aspiration with a 27.5-gauge needle and 5-ml syringe and replaced by culture medium. The granulosa cells are harvested through a slit by gentle scraping of the inner wall, using a microinoculation loop. Medium and cells are recovered with a Pasteur pipette and centrifuged (150 g, 5 min, 4~ The cell pellet is washed with culture medium, and the cells are resuspended in culture medium, counted, and distributed into Nunclon Delta 24-well plates (Nunc, Copenhagen, Denmark): 0.3 to 1 • 10 6 cells in 1 ml of culture medium per well. The culture medium is Dulbecco's modified Eagle's medium and Ham's mixture F12 (DMEM/F12) containing penicillin (100 U/ml), streptomycin (0.1 mg/ml), (Eurobio, Paris, France), insulin (2/zg/ml), and without fetal calf serum. The cells are incubated at 37~ in a 5% CO2 humidified atmosphere (Sanyo incubator, Tokyo, Japan). Two to 4 hr after incubation, attached cells are washed with culture medium to eliminate dead cells and erythrocytes. Cultures are carried out in the presence or absence of the following hormones"

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FSH (NIH-FSH-S 1), 1.5 mU per milliliter of culture medium, 0.5 IxM testosterone, or 0.5 txM dehydrotestosterone. Culture media are changed daily. Cell number is deduced from DNA concentration. DMEM/F12 and hormones are supplied by Sigma.

Neurophysin and Oxytocin Isolation from Culture Media Before RIA, immunoreactive neurophysin and oxytocin secreted into the culture media are isolated using Sep-Pak C~8 cartridges (Waters Assoc.). One milliliter of culture medium is filtered through the cartridge previously washed with methanol, water, and 0.1% (v/v) trifluoroacetic acid, and elution is obtained with 3 ml of 40% (v/v) acetonitrile in 0.1% (v/v) trifluoroacetic acid. Eluate is evaporated in a Speed-Vac concentrator (Savant Instrument Co., Hicksville, NY), then dissolved into RIA buffer. HPLC identification of oxytocin peptides is also performed on culture media purified using Sep-Pak C18 cartridges. The procedure is the same, but elution is obtained with 20% (v/v) acetonitrile in 0.1% (v/v) trifluoroacetic acid. The recovery yield of oxytocin-Gly-Lys-Arg, oxytocin-Gly-Lys, oxytocin-Gly, and oxytocin-NH2 is determined as 83, 77, 67, and 69%, respectively.

Neurophysin, Oxytocin, and Progesterone Radioimmunoassay Neurophysin RIA is carried out with purified bovine neurophysin I (11) as standard and rabbit anti-neurophysin I serum (618-12) prepared in the laboratory and used at a final dilution of 1 : 75,000. Separation of free from bound neurophysin is obtained by zirconyl phosphate. Sensitivity is 40 pg and 50% displacement of the 125I-labeled neurophysin I is obtained by either 0.6 ng of neurophysin I or 200 ng of neurophysin II. Oxytocin RIA is carried out as previously described (7). Oxytocin and Cterminally extended oxytocin peptides used as reference in RIA or HPLC are synthesized (see above). A minimum of 2 pg of oxytosin is detected and 50% displacement of the tracer (~25I-labeled oxytocin) is obtained with 75 pg of oxytocin, 300 pg of oxytocin-Gly-Lys-Arg, 250 pg of oxytocin-Gly-Lys, and 210 pg of oxytocin-Gly, thus allowing a sensitive detection of the mature oxytocin and the corresponding extended forms. Concentrations of progesterone in culture media are directly determined using an RIA kit (PROGCT) (CIS; Bio Industries, Paris, France). Sensitivity is 15 pg.

Results This system is particularly interesting because in this organ the appearance of a peak of pro-OT/Np mRNA at days 1-4 after the heat period is followed by the delayed appearance of free and active oxytocin at days 8 to 13 of the

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cow estrous cycle. The measurements made indicated clearly the presence of a set of C-terminally extended forms of OT, that is, OT-Gly-Lys-Arg, OTGly-Lys, and OT-Gly, the substrate for the c~-amidating enzyme producing OT(NH2) (Fig. 2). A similar pattern was obtained in human granulosa cells in culture (Fig. 2). interestingly, the neurophysin-like material produced in corpus luteum 3-4 days after heat contained a higher molecular weight species with a pHi of 4.7. This material was indistinguishable from hemisynthetic pro-OT/Np (13) and was converted by a putative pro-OT/Np convertase from either neurohypophysis or corpus luteum into stoichiometric amounts of mature neurophysin and OT-Gly-Lys-Arg, which was identified by HPLC (as described above). Analysis of the corpora lutea 4 to 7 days after estrus revealed the presence of "Np-like" material, including the pHi 4.3 "OT-associated" Np, indicating that precursor processing had taken place. These data (8) unequivocally demonstrated that pro-OT/Np mRNA, which appears as early as day 1 after estrus, is translated into the precursor but that the latter is not processed further before day 4 after estrus. Moreover, the production of C-terminally extended forms of OT may indicate that processing is incomplete because mature OT(NH 2) appeared later. This might suggest a physiological role for these intermediary processing forms that remains, as yet, unidentified in ovary.

Detection and Characterization of Processing Enzyme Activities The major difficulty in detecting proteolytic enzymes in tissue, or cell, extracts resides in the adequate definition of a specific substrate. Homogenization of biological materials results in removal of cellular barriers, which add to the complexity of the resulting proteolytic activities in crude extracts. Therefore, the definition of a specific activity at the early stages of purification of a given, specific, proteolytic activity remains awkward. In attempting to quantitate the presence of a processing activity in biological materials the researcher should in general perform a first fractionation step before meaningful numbers may be obtained (14). Because cleavage of propeptides generally occurs at basic residues, the various substrates used by several workers are either basic amino acids or peptides bearing in their sequence the monobasic or dibasic moieties.

Use of Fluorogenic Substrates Fluorogenic substrates are the most convenient tools for the detection of proteolytic activities and for the rapid determination of kinetic constants. Either derivatives of arginine and lysine or of dibasic moieties Lys-Arg, Arg-

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Arg, Arg-Lys, and Lys-Lys can be used. Because of the lack of solution structure of these substrates, they may be recognized by several proteolytic activities with a "basic residue" selectivity. Therefore they are not recommended for use at the early stages of protease purification, but can be efficient tools to determine enzyme parameters on purified preparations of processing endoproteases (or exoproteases). This can be illustrated in the cases of enzymes that cleave on the carboxyl side of paired basic residues (15, 16). In the first case the substrate used is Boc-Gln-Arg-Arg-MCA, a tert-butyloxycarbonylmethylcoumarin derivative of Gln-Arg-Arg. In a typical assay the synthetic substrate (obtained by solidphase synthesis) is incubated (20 nmol) in the presence of the S. cereoisiae enzyme in a final volume of 250/zl of solution containing 0.2 M Tris-HC1 buffer (pH 7.0) and 0.1% (v/v) Lubrol and 1 mM CaC12 (because the enzyme is a membrane-bound species, activated by Ca 2§ The reaction is kept for 1-24 hr at 37~ and at the end of the incubation 3 ml of H20 is added and the amounts of 7-amino-6 methylcoumarin (AMC) released from the substrate are measured with a fluorometer (excitation at 380 nm and emission at 460 nm). An arbitrary unit system can be defined as the enzyme quantity that can release, in 1 hr of incubation under the standard conditions, a certain quantity (in nanomoles) of AMC. To attempt the definition of an enzyme specificity, various derivatives are designed (see Table I) and percent activity is measured. It would be preferable in either case to measure the Km and Vmax from Line-weaver-Burk or Hofstee-Eadie plots. A comparable approach was used by Brenner and Fuller (16) to analyze the behavior of the Ca 2+-dependent, subtilisin-like endoprotease product of the KEX2 gene of S. cereoisiae. Boc or Ac derivatives of tetra- or pentapeptides were designed in which the C terminus is derivatized by a methylcoumarinamide group (see Table II). This study analyzed the influence of the doublet on Kex2 specificity. Lys-Arg, Arg-Arg, and Lys-Lys derivatives were compared, but not the Arg-Lys arrangement (Table II). Because the P'I and other P' positions are not filled by amino acid residues, only the P1, P2, P3, P4, and, in a few cases, P5 positions were analyzed with respect to the kinetics of Arg-MCA (or Lys-MCA) bond hydrolysis. A typical standard assay is as follows: 100/xM Boc-peptide-AMC in 50 /xl of 200 mM Bis-Tris-HC1 (pH 7.0), 1 mM CaC12, and 0.5% (v/v) dimethyl sulfoxide (DMSO) in 0.01% (v/v) Triton X-100. Initiation of the reaction is achieved by adding ---75 units of the enzyme and incubation is performed for 4 min at 37~ then stopped by placing tubes in ice water and adding 950/~1 of 0.125 M Z n S O 4. The AMC released from digested substrates is evaluated by fluorometry [excitation (hex) 385 nm; emission (hem) 465 nm]; 1 unit of Kex2 is defined as the release of 1 pmol of AMC/min.

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I M M U N O L O G I C A L AND B I O C H E M I C A L APPROACHES TABLE I

E n d o p r o t e a s e Specificity t o w a r d Various Boc Peptidyl-MCA or Aminoacyl-MCA a

Substrate b

Activity (%)

Boc-Gln-Arg-Arg-MCA Boc-Leu-Arg-Arg-MCA Boc-Gly-Arg-Arg-MCA Boc-Leu-Lys-Arg-MCA Boc-Gly-Lys-Arg-MCA Boc-Val-Pro-Arg-MCA Boc-Ala-Pro-Arg-MCA Boc-Glu-Lys-Lys-MCA Pro-Phe-Arg-MCA Z-Phe-Arg-MCA Bz-Arg-MCA Arg-MCA Leu-MCA

100.0 97.8 14.3 118.1 74.9 51.4 25.3 0.8 0.5 0.3 1.1 1.1 0.5

a Using a Ca: §-dependent membrane-bound enzyme from yeast (15). Protease activity for Boc-Gln-Arg-Arg-MCA was taken as 100%. Each substrate (20 nmol) was incubated for 10 hr with the enzyme at 37~ (as described in Use of Fluorogenic Substrates). MCA released was measured by fluorometry. b Boc, tert-butyl oxycarbonyl; Z, benzyloxycarbonyl; Bz, benzyl.

TABLE II

S t e a d y S t a t e K i n e t i c P a r a m e t e r s for C l e a v a g e o f Boc-peptidyl-MCA Substrates a Substrate

P5

P4

Ac-Pro- MetBocBocBocB o c - Arg Boc BocBocBocBoc a

P3 Tyr LeuLeu Gin Val Val GlnLeuGin Glu -

P2

P1

P'I

LysLysArgArgArgPro Ala ThrGly Lys -

Arg-MCA Arg-MCA Arg-MCA Arg-MCA Arg-MCA Arg - M C A Arg-MCA Arg-MCA Arg-MCA Lys - M C A

By Kex 2 endoprotease from S. cerevisiae (16).

kcat

gm

kcat/grn

(sec_l)

(/zM)

(sec -1 M -1)

25 23 45 21 21 31 26 18 6.1 0.19

2.2 3.9 17 13 19 150 210 800 320 55

11,000,000 5,900,000 2,600,000 1,600,000

1,100,000 210,000 120,000 22,000 19,000 3,500

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Results This technique allowed convenient, rapid determination of steady state kinetics, optimal pH, and of kcat, gm, and k c a t / g m ratios (Table II). However, with respect to analyzing the influence of substrate structure on these parameters, one major drawback arose from the following features: (a) the relatively short length of the substrates and the absence of ordered peptide structure in solution, and (b) the absence of amino acid sequence or the C terminus of the dibasic and principally the lack of P'I residue, which turned out to be critical in processing at the dibasic-Xaa (P' 1) site (3).

Use of Synthetic Peptides Some laboratories have used a different approach, taking into account the basic considerations about the expected substrate requirements of the endoproteolytic processing enzymes (General Considerations, above). Therefore, synthesized peptides included in their sequence the mono- or dibasic moieties, which constitute a part of the putative signals for enzyme recognition. This strategy can be illustrated in the case of prooxytocin-neurophysin (proOT/Np) (7) and the prosomatostatin family (17). In a typical experiment, the substrate reproducing or mimicking the precursor sequence around the monoor dibasic cleavage site is exposed to the putative processing endoproteolytic activity. At the end of the incubation period, the remaining substrate and the generated fragments are separated by HPLC and identified by either (or a combination) of the following methods: (a) by reference to synthetic peptide standards, (b) by RIA with the appropriate antibodies, and (c) by amino acid composition and/or sequencing. Because the unequivocal identification of fragments is critical, some variance in the experimental procedure was designed; that is, either the substrate or the generated fragments are derivatized using an amino-terminal reagent absorbing in the visible region (e.g., DABITC) (18) (see Fig. 3).

Unlabeled Peptides A typical illustration of this technique is provided by pro-OT/Np(1-20), the N-terminal domain of the common precursor to oxytocin and neurophysin. To detect a putative processing endoprotease capable of cleaving at the Lys~-Arg 12 pair, a 20-amino acid residue synthetic peptide reproducing the N-terminal sequence of pro-OT/Np was synthesized by the Merrifield technique (Fig. 4). It was used to monitor the purification and characterization of a metalloendoprotease detected in bovine neurohypophysis and in the corpus luteum (14).

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II IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

A LysArgI

I

Endoprotease

LysArg +

I

LysArg

I I

DABITC

LysArg

Endoprotease

LysArg +

C

I

LysArg

I I

Endoprotease

.LysArg +

DABITC

LysArg +

FIG. 3 Schematic representation of the various methods used for analysis of a peptide substrate and its fragments generated after endoproteolytic cleavage at the Lys-Arg $ Xaa dibasic site. (A) The peptide fragments were analyzed after HPLC by UV (Azz0nm). (B) The peptide substrate was prederivatized with DABITC and both the intact precursor and its N-terminal fragment were identified by HPLC and detection at 436 nm. (C) Both the N- and C-terminal peptide fragments were derivatized after endoproteolytic cleavage by DABITC and detected, after HPLC separation, by their A436n m values.

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IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

Cs~Tyr-I l e -lev-Glanl-Asn-Ct - L eSus-Pr~ -Asp-Leu-Asp-val-ArgS 1

10

11

12

10

11

12

I I

I I

13

169 20

Endopeptidase I

13

20

Cys-Tyr-lle-Gln-Asn-Cys-Pro-Leu-Gly-Gly~Lys~Arg + Ala-Val-keu-Asp-keu-Asp-Val-Arg

l--s

s

I

Carboxypeptidase B-like I

1o

Cys-Tyr-Ile-Gln-Asn-Cys-Pro-Leu-Gly-Gly IS s~l . I

I Amidating enzyme 1

9

Cys-Tyr-lle-Gln-Asn-Cys-Pro-Leu-Gly-NH 2 Oxy tocin

FIG. 4 A schematic view of prooxytocin-neurophysin(1-20) processing by enzyme activities from secretory granules lysates. [From Clamagirand (7).]

In a standard assay, 20 txg of the reference substrate is incubated with an aliquot of enzyme preparation (representing 1 txg of protein) in 100 mM ammonium acetate buffer, pH 7.0, in a final volume of 200 txl for 24 hr at 37~ The resulting fragments are analyzed by HPLC after acidification with 15 lxl of 1 N HC1, using a Cl8 column (Whatman), and eluted by a gradient of 0.05% (v/v) TFA in acetonitrile on an LKB apparatus. Peptide elution is monitored at 220 nm. Both the (1-12) and (13-20) fragments are identified by reference to synthetic standards and by amino acid composition (Fig. 4). The identification of both N- and C-terminal peptides resulting from the ArglZ-Ala ~3 cleavage is essential because contaminating exoproteases such as amino- and/or carboxypeptidase(s) can further degrade the generated products. Quantitation of the enzyme reaction can be achieved by several methods, including (a) evaluation of the disappearance of substrate, and (b) measurement of the generated N- and/or C-terminal fragments.

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

R5

/

t

R8

aB

R8

OR

~ I,o.Tol

Rs

A

E

abc

111

I"

0

TIME (rain)

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171

A comparison of the numbers obtained via the two techniques may be useful to reveal contaminating activities that may affect the stability of either, or both, generated products during the incubation conditions. This technique may also be applied to a number of derivatives of the reference sequence to study the role of the dibasic moiety, of peptide length, secondary structure, and of given amino acid residues in the kinetics of peptide cleavage and in substrate recognition (13, 19). However, careful interpretation of the kinetic data suggests that a certain number of conditions be respected. These include the following: (a) only peptides of similar length should be compared, and (b) structural information on the solution conformation of the peptide substrates should be obtained by circular dichro~sm, Fourier transform-infrared, and possibly one- and two-dimensional highresolution ~H nuclear magnetic resonance (NMR) spectroscopies (19, 20). Prederivatization of Peptide Substrate In this case the N terminus of the synthetic substrate is first reacted with a chromophore reagent [such as dimethylaminoazobenzene isothiocyanate (DABITC)] to generate the dimethylaminoazobenzene thiocarbamoyl peptide (DABTC-peptide). This is well illustrated in the case of the detection and purification of the "RXVRG" endoprotease from skin exudate of Xenopus laevis (Fig. 5). There the substrate was obtained by reaction of DABITC (21) with a tetradecapeptide (peptide I) mimicking the conserved region of X. laevis skin hormonal precursors, that is,

DABTC-peptide I = DABTC-AspValAspGluArgAspValArgGlyPheAlaSerPheLeu-NH2 Both amino- and C-terminal blockage prevent the substrate from amino- and carboxypeptidase(s) attack. The preparation of substrate and steps of the enzyme assay are as follows. 1. The substrate (peptide I) and the fragments generated by its cleavage at the R-G (i.e., a = AspValAspGluArgAspValArg) or R-D (i.e., b = Asp-

FIG. 5 An enzyme test to detect "RXVRG" endoprotease from X. laevis skin secretions. The peptide used as substrate, called "Kermit," mimicked the consensus sequence of hormone precursors around a processing locus (R $ G)-DABTC-Kermit = DABTC-D1VDERSDVR8GFASFL-NH2). Detection of the remaining substrate as well as of its generated fragments by cleavage at R8 (a) and R5 (b) was made after HPLC separation by A436n m measurements. (c) Elution position of the unmodified DABTC-Kermit. [From Kuks et al. (21).]

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

ValAspGluArg) or S-F (i.e., c = AspValAspGluArgAspValArgGlyPheAlaSer) bonds are reacted with DAB ITC. 2. Fifty micrograms of peptide is derivatized with dimethylaminoazobenzene isothiocyanate as described (18) and purified by HPLC (Milton Roy) on a Nucleosil 5-tzm C~8 column (146 x 4.6 mm) eluted with 35% (v/v) acetonitrile in aqueous 1% (v/v) N-methylmorpholine acetate buffer at pH 5.3 (flow rate, 1 ml/mn). DABTC-peptides (retention time, 25-40 min) are collected, dried, and then redissolved in acetonitrile-water (2:1, v/v) and stored at -20~ 3. Fifty picomoles of DABTC-peptide I is incubated with 12.5 Izl of enzyme preparation for 2 hr. The reaction is stopped by the addition of 12.5 Izl of pyridine and the mixture is analyzed by HPLC [30% (v/v) acetonitrile; flow rate, 1.5 ml/min]. All cleavage fragments elute separately within 5 min. 4. The fractions that generate the fragment DABTC-peptide I-(1-8) with little or no DABTC-peptide I-(5) (Fig. 5) are defined as active. 5. Effects of amino acid substitution on the endoprotease action are monitored by a competition test with the reference standard (21).

Postderivatization of Generated Fragments This technique can be illustrated in the case of the detection and characterization of an arginine-specific monobasic endoprotease from rat intestinal mucosa (22, 23). In this case a synthetic peptide (LeuGlnArgSerAlaAsnSerNH2) is synthesized corresponding to the prosomatostatin(62-68) region as a carboxamidated form: that is, pro-S(62-68)-NH2 is used as substrate and exposed to the enzyme. Then the generated fragments and the recovered intact substrate are subjected to N-terminal derivatization with DAB ITC, and the derivatized fragments are analyzed by HPLC separation (Fig. 6). The standard assay is as follows. Aliquots of enzyme are incubated with 2 nmol of pro-S(62-68)-NH2 in 10 mM sodium phosphate, pH 7.4, for 1 hr at 37~ in a final volume of 15 tzl. Controls establish that the enzyme activity is perfectly stable for at least 5 hr at 37~ The reaction is stopped by addition of dimethylaminoazobenzene isothiocyanate in pyridine (2 mg/ml) and incubated at 70~ for 50 min. The excess reagents are extracted with heptane-ethyl acetate (2 : 1). The aqueous phase is dried and redissolved in 50 ~1 of pyridine-H20 (1 : 1), and one-tenth is analyzed by HPLC using a $5ODS2 column (4.6 x 250 mm) (Prolabo, Paris, France) eluted with triethylamine acetate [1% (v/v) buffer], pH 5.3, containing 36% (v/v) acetonitrile, at a flow rate of 1 ml/mn. The dimethylaminoazobenzene thiocarbamoyl peptides (DABTC-peptides) are monitored at 436 nm. Under these conditions, DABTC-pro-S(62-64), DABTC-proS(65-68)-NH2, and the DABTC-substrate are visualized on a single run in 12 min (see Fig. 6).

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C 0.01

B

A

r

E rm-

v

0.005

t <

relenlion time (min)

FIG. 6 HPLC separation and A436detection of the peptide fragments generated after cleavage of the peptide substrate pro-S(62-68)L-NH2 = LQRSANSL-NH2 and its N-terminally derivatized DABTC fragments, that is, DABTC-pro-S(65-68)L-NH2, DABTC-pro-S(62-68)L-NH2, and DABTC-pro-S(62-64) (A-C, respectively). [From Bourdais et al. (23).]

Evaluation of the amounts of digested substrate is made either by measurement of the remaining peptide substrate and/or by measurement of the generated fragments. K m values are therefore determined using various peptide concentrations (80-400 txM) under conditions of initial velocity in which cleavage does not exceed 5%. The technique is similarly applied to a number of related peptide derivatives (23) to evaluate the influence of structural parameters and amino acid substitutions on the selectivity of the enzyme.

Use of "Full-Length" Hormonal Precursors Evidently, when attempting to correlate an endoproteolytic activity with its putative endogenous substrate, the researcher would like to test the enzyme with a genuine precursor. One of the main problems centers around the availability of these proforms and their particular properties (essentially their marked hydrophobicity, in some cases). Whereas proinsulin is readily available, prosomatostatin is somewhat difficult to purify from producing

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

cells (24). In some cases using cell-free translation systems or translational expression in transfected heterologous cells (i.e., Cos or baculovirus systems) may be recommended to produce the adequate prohormone substrate from mRNAs or cDNAs. Whatever the source of proform used, the detection of fragments should be based on the use of an adequate separation technique (electrophoretic and/or HPLC) followed by immunochemical detection and identification (including Western blotting on gels). The choice of the technique will depend essentially on the respective size of the precursor and the expected processing fragments. Also, the specificity of the relevant antibodies will be critical in assessing the correct generated segments. In general those experiments are time consuming and the extent of information that is obtained is limited. In particular it should be noted that unless selective antibodies are used, which may recognize well-defined N- or C-terminal epitopes, the precise identification of the peptide bond(s) that are cleaved during processing may require some amino acid sequencing. Because C-terminal sequencing remains a difficult task, there may be some doubt about the actual C-terminal sequence of the relevant fragment. Finally, although this kind of experiment may give a satisfactory answer about the actual capacity of a given endoprotease to cleave the adequate processing site(s), this approach is not appropriate for structure-function relationship studies. An illustration of this technique can be found in the example of prooxytocin-neurophysin, either obtained by hemisynthesis (13) (Fig. 7) or purified from producing organs (8). It should be noted that, in principle, the tools used for the detection of processing intermediates in extracts (see Identification of Precursor-Processing Intermediates, above) or in situ (Use of Selective Antibodies, below) may apply for the detection and identification of precursor fragments generated by cleavage using a putative prohormone convertase. A hemisynthetic form of prooxytocin/neurophysin I was obtained as described in Brakch et al. (13) and tested for its ability to be cleaved by the convertase isolated from bovine neurohypohyseal or corpus luteum granules. The resulting products were separated by chromatography and the fragments were analyzed by UV absorbance at 220 nm by reference to the elution times of standards. The data demonstrated that solely NpI and the OT-GlyLysArg ~2 peptide were generated, indicating that the only bond to be cleaved under these conditions was the ArglZ-Ala ~3 one. Experimental Procedure

Details of the pro-OT/NpI synthesis are in Brakch et al. (13). The reference peptides are numbered as in Identification of Precursor-Processing Intermediates (above).

175

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

t

A

B

9

0.1

0

10

20

30

40

0

Retention

10 time

20

30

40

(min)

FIG. 7 Conversion of hemisynthetic prooxytocin-neurophysin by the endoprotease from bovine neurohypophysis and corpus luteum. (A) HPLC elution pattern of the two components of the hemisynthesis, that is, OT-GlyLysArgAla 13 (left arrow) and [des-Ala']di(DABTC)-neurophysin I (right arrow), and of the resulting covalent adduct, that is, hemisynthetic pro-OT/NpI (middle peak). (B) Elution pattern of the fragments obtained after exposure of hemisynthetic prooxytocin and neurophysin I to the endoprotease. (a) Hemisynthetic precursor; (b) OT-GlyLysArgl2; (c) di(DABTC)-neurophysin I. Peptides were identified by their retention times (a and c) or amino acid composition (b). For details, see Brakch et al. (13).

Twelve micrograms of hemisynthetic precursor is incubated with an aliquot of the enzyme preparation (~2/zg of protein) in 100 mM ammonium acetate, pH 7.0, in a 200-/zl final volume, for 24 hr at 37~ Then the reaction is stopped by 15/zl of 1 N HC1 and the resulting fragments are separated by HPLC, monitored by UV absorbance at 220 nm combined with RIA using anti-oxytocin and anti-Np antibodies (see Identification of Precursor-Pro-

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cessing Intermediates). The column is a Nucleosil 5 CN column (SFCCShandon, France) eluted first with 10% acetonitrile, 90% of 0.05% aqueous TFA for 20 min. Then a gradient is established from the original mixture to 60% acetonitrile, 40% of 0.05% TFA for 40 min. The precursor, Np, and N-terminal fragment (peptide II) were identified by the appropriate RIA using anti-Np and anti-OT antibodies and by amino acid composition. A similar approach was used by Camier et al. (8) to identify a pI 4.7 neurophysin-like material isolated from cow corpus lutea 2-3 days after estrus. This was exposed to either trypsin or to the above-described convertase and the resulting products were separated by HPLC and identified by their retention times coupled with their reactivity toward anti-Np and anti-OT antibodies. The treatment released OT-GlyLysArg (and OTGlyLys) from the pI---4.7 precursor material (8).

Critical Evaluation of Different Techniques In the absence of full-length precursor, the use of unlabeled peptide is by far the most reliable because the peptide sequence (and, hence, conformation) is not affected by derivatization, a factor that should not be neglected when the influence of substrate structure is to be evaluated (19). The technique is tedious and the worker must properly evaluate the extent of cleavage. Additionally, large amounts of peptide and of enzyme may be consumed. Prederivatization offers the advantage that the generated N-terminal fragment is rapidly and easily identifiable, but its possible effect on substrate conformation and on the selectivity of cleavage should be at first carefully evaluated. Postderivatization has the advantages of both techniques except that the kinetics of DABITC reaction with the three N termini (of the intact substrate, and of the N and C terminal-generated fragments) is not necessarily comparable and that discrepancies can be observed. In this case quantitation of the generated fragments may be awkward. The difficulty can be overcome by monitoring a single fragment. In any case, the researcher should choose a technique depending on the purpose of the study (detection of activity, rapid determination of kinetics, or more detailed analysis of structural parameters on the enzyme reaction). Large amounts of enzyme and substrate(s) may often be needed to obtain a complete set of kinetic determinations.

Use of Selective Antibodies to Identify Processing Events in Situ One of the questions regarding posttranslational proteolytic processing concerns the subcellular localization of the enzyme(s) implicated in the relevant peptide bond cleavage(s). On the basis of experiments run on the prooxytocin

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

177

and provasopressin systems in the hypothalamoneurohypophyseal tract, it was thought that the maturation of the precursors occurred within the membrane-limited secretory granules during their translocation from the cell bodies to the nerve terminals (25). This was established on the basis of time course experiments (25). Serious attempts to analyze these proteolytic events more precisely were based on a general strategy adopted by a few laboratories. Indeed, antibodies were raised against structural motifs of a given precursor that were either (a) unmasked after dibasic or monobasic cleavage of the precursor, as in the case of proinsulin and prosomatostatin(26-28), and used in immunochemistry to monitor these processes, or (b) generated after proteolytic cleavage at the processing locus, as in the case of proopiomelanocortin (29), and used similarly in immunocytochemistry. Interestingly, these methods suggested that these proteolytic events occur in the Golgi apparatus, or at least initiate in a late compartment of the Golgi (transGolgi network).

Proinsulin System Proinsulin is an 8-kDa single polypeptide chain that contains three disulfide bridges and undergoes dibasic processing at the Lys62-Arg63 and Arg3~-Arg 32 sites, respectively (see Fig. 8). To analyze these processing events in pancreatic islets, Orci and co-workers raised a selective monoclonal antibody against the C-terminal sequence of insulin B chain, a proinsulin motif situated next to the dibasic cleavage site (Fig. 8) (35). In parallel, a selective antiproinsulin antibody was used to monitor unprocessed precursor. They developed a strategy based on the combined use of these antibodies to measure concomitantly the appearance of this epitope by proteolytic processing with the labeling of a pH probe (DAMP) used as an in situ monitor of pH.

Experimental Procedure Normal adult rats are fixed by vascular perfusion with 1% (v/v) glutaraldehyde in 0.1 M phosphate buffer (pH 7.4); pancreas fragments are dehydrated in ethanol and embedded at low temperature in Lowicryl K4M. Thin sections are collected on nickel grids and immunolabeled by the protein A-gold technique (31) with monoclonal anti-insulin (32) (MAb3, diluted 1 : 100,000) or monoclonal anti-proinsulin (GS-9A8 MAb, diluted 1:3000) antibodies, followed by rabbit anti-mouse IgG (diluted 1 : 200), and the protein A-gold (size of gold particles, 10 nm) solution (1 : 70, v/v). Sections are stained with uranyl acetate (10 min) and lead citrate (1 min) before examination with the electron microscope. Clathrin immunolabeling is performed as described

178

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FIG. 8 Schematic representation of proinsulin. Insulin A and B chain amino acids are in black and the connecting peptide C residues are in white. [From Ellis et al. (30).] 9 1968 AAAS. in Orci et al. (33). The quantitative evaluation of insulin and proinsulin immunoreactive sites is carried out as previously described (33). Data on the proinsulin immunolabeling are from Orci et al. (33). Exposure of B cells to DAMP is carried out on islets of Langerhans isolated by collagenase digestion (34) and incubated for 1 hr at 37~ with 30/~M DAMP (35). At the end of the incubation, islets are processed to localize insulin and DAMP on pairs of consecutive thin sections of the same cells. As judged by immunofluorescence (33), DAMP does not appear to interfere with the clearance of proinsulin out of the Golgi. Results

The observations made indicated that the immunoreactivity was absent in the Golgi but became detectable in the clathrin-coated vesicles emerging from the trans-Golgi pole. This process was accompanied by a marked modification of the so-called

[9] IDENTIFYINGNEUROPEPTIDE-PROCESSINGPATHWAYS

179

pH from neutral to more acidic values (apparent pH 7.0-6.8 to 5.5-6.0), reflecting a progressive acidification of the compartment. The use of a metabolic poison, antimycin (33), was not accompanied by the appearance of antiinsulin immunoreactivity in the Golgi apparatus, indicating that a prolonged sojourn of the precursor in the Golgi stack did not result in the generation of mature, immunoreactive insulin. The interpretation of these elegant studies was challenged by the more recent discovery that the two dibasic cleavages releasing insulin from its precursor may occur in different subcellular compartments and by the intervention of distinct enzyme species (36). Because insulin generation was monitored by a single antibody directed toward the C terminus of the hormone, the experiments did not report on proteolytic events occurring at the N terminus, that is, after cleavage of the Lys62-Arg63 locus at the peptide C-insulin A chain junction (see Fig. 8). These experiences underline the strong need for adequate immunochemical reporters when monitoring the processing of a multifunctional precursor (see below).

Prosomatostatin System Prosomatostatin is the common precursor in a number of bioactive fragments in mammalian producing cells (Fig. 9). Indeed, mono- and dibasic cleavages at the arginine site generate somatostatin-28 (S-28) or its amino-terminal domain S-28(1-12), whereas dibasic maturation at the level of the Arg-Lys moiety liberates somatostatin-14 (S-14) plus the N-terminal domain of the precursor, called "8 kDa" (Fig. 9). In teleostean fishes such as the anglerfish (Lophius piscatorius) these hormonal peptides are produced in the pancreatic islets of the Brockmann organs via two distinct expression products, prosomatostatin I (PSI) and prosomatostatin II (PSII), from two genes. Whereas PSI maturation gives rise to S-14 I, processing of PSII produces exclusively S-28 II (Fig. 10).

Rat Brain Cortical Cells Lepage-Lezin et al. (27) have made use of selective antibodies against S-14 and S-28(1-12), respectively, to evaluate the situation in rat brain cortical cells. The following combined techniques were performed to analyze prosomatostatin processing: (a) a high-performance equilibrium sedimentation separation of granules/synaptosomes, Golgi, and ER fractions from cell homogenate [adapted from Trifaro and Duerr (37)], coupled with both enzyme markers and immunochemical identification of the subcellular fractions, (b)

180

II

IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES R64

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Ab S20(1-12)

FIG. 9 A schematic representation of the prosomatostatin-processing products in rat brain cortical cells. [From Lepage-Lezin et al. (27).]

HPLC separation of the intact precursor and of the main prosomatostatin derivatives that were analyzed by RIA, with the relevant antibodies, and (c) use of anti-S-14 and anti-S-28(1-12) antibodies on the Golgi fractions for immunocytochemical detection of the precursor, S-14, and S-28 compounds and of the S-28(1-12) motif either in the precursor deleted from S-14 or/and in the dodecapeptide itself (Fig. 9).

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS AF-prosomatostatin

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72

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Ab Belle

Ab S 14 $28

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FIG. 10 Prosomatostatin I and II proteolytic processing in L. piscatorius Brockmann bodies. Top: AF-prosomatostatin I. Bottom: AF-prosomatostatin II. Antibody (Ab) S-14, Ab S-28(1-12), and Ab Bella are the various polyclonal antibodies used for the cytoimmunochemical detection of the precursor (S) and their fragments.

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Experimental Procedure Subcellular Fractionation Male Wistar rats (100-150 g) from Centre d'Elevage Robert Janvier (St. Bertheon, France) are maintained under normal housing conditions (12-hr light-dark period) and fed ad libitum (Purina Chow). After decapitation, the optic chiasma is cut, before the brain is removed from the skull, and placed over ice. Hypothalamus containing the median eminence is dissected, and the rest of the brain is cut perpendicularly into three pieces. The cortex is separated from the midbrain with a sharp blade, avoiding the white matter. Either cortical tissue is minced using a pair of scissors, or hypothalamic fragments are homogenized [15% (w/v)] with a Potter-Elvehjem homogenizer in ST buffer [0.3 M sucrose, 20 mM Tris-HC1, pepstatin and aprotinin (5 mg/ml), pH 7.2] containing 0.005 M MgC12, 10/zM NaC1. Subcellular fractionation is performed by a modification of the technique described in Schnabel et al. (29). The homogenate is centrifuged at 300 g for 5 min at 4~ followed by 800 g for 10 min at 4~ the supernatant is centrifuged at 20,000 g for 8 min at 4~ (P2). The recovered supernatant ($2) is centrifuged again at 100,000 g (P3). Each of these pellets, P2 and P3, is resuspended in ST buffer, placed over a discontinuous gradient (gradient I) made of 0.8, 1.0, 1.2, 1.4, and 1.6 M sucrose in 20 mM Tris, pH 7.2, containing pepstatin and aprotinin (5 /zg/ml), and centrifuged at 113,000 g for 70 min at 4~ The band sedimenting at the interphase between 0.8 and 1.0 M sucrose from the P3 gradient, corresponding to the "crude Golgi" (i.e., mixture of Golgi and ER), is adjusted to 1.1 M and layered over a second discontinuous sucrose gradient (gradient II) made of 1.25, 1.3, and 1.4 M sucrose in ST buffer; the tube is then covered with 0.5 M ST and centrifuged at 100,000 g for 90 min. Fractions from the different gradients are treated according to the assay performed (see below). Characterization of Subcellular Fractions The different subcellular fractions are characterized (a) by measuring enzyme activities recognized as markers of lysosomes, Golgi complex, and endoplasmic reticulum (ER) (38), and (b) by Western blot identification of ER-specific proteins (39). Serotonin is chosen as the synaptosomal marker, a neurotransmitter known to be present in the cortex only in nerve endings. Electron microscopy analysis is performed on ER- and Golgi-enriched fractions. Enzyme activities are quantified in 1-ml fractions of gradient II (obtained from the bottom of the tube). Glucose-6-phosphatase (EC 3.1.3.9), recognized as an ER marker (38), is measured using glucose 6-phosphate (Sigma) as substrate according to Morr6 (40) and UDPgalactose: N-acetyl-o-glucosamine 4-/3-o-galactosyltransferase activity (EC 2.4.1.90, N-acetyllactosamine synthase [UDP[~4C]galactose, from Du Pont-New England Nuclear (Boston, MA), and N-acetylglucosamine, from IBF (Villeneuve-la-Garenne,

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

183

France)] as Golgi marker (37). Acid phosphatase (EC 3.1.3.2) is measured in 1-ml fractions obtained from the first gradients as an indication of lysosomal activity. An aliquot of 20 Ixl obtained from the bands recovered after gradient I or II is diluted with 500 ml of 5 ~M Tris-HC1 (pH 7.2) buffer, sonicated, precipitated with trichloroacetic acid [10% (v/v) final], and centrifuged. Electrophoresis and transfer are performed according to Laemmli (41) and Towbin et al. (42). An antibody specific to ER membrane proteins (32) is used at 1 : 5000 dilution, incubated with the transfer membrane for 10 hr, and revealed after thorough washing with the alkaline phosphatase system (Bio-Rad). Serotonin is assayed in all fractions obtained from the two gradients, by aspiration, further adjustment to 0.45 M sucrose, and centrifugation at 100,000 g for 60 min. The pellet is homogenized and assayed by radioenzymology. Microscopy examination is performed on aliquots obtained from the bands sedimenting at 0.9 and 1.18 M sucrose of gradient II. The tube is pinched with a hypodermic needle and the band suctioned with a syringe. For conventional thin sectioning the subcellular fractions are fixed in suspension by glutaraldehyde at 2.5% (v/v) final concentration. After repeated washing, the pellets are postfixed in 1% (v/v) osmium tetroxide solution in cacodylate buffer, dehydrated, and embedded in an Epon-Araldite mixture (43). For immunogold labeling, the Golgi-enriched fraction is spread onto cover slides coated with polylysine, fixed with a few drops of 3% (v/v) paraformaldehyde in phosphate-buffered saline, and then processed according to Dunia et al. (43). Somatostatin-14- and Somatostatin-28(1-12)-Like Peptide Identification All fractions under study, obtained from the different gradients, are aspirated and carefully adjusted to 0.45 M with Tris-HC1, pH 7.3, centrifuged at 100,000 g, and both these pellets and the P2 and P3 pellets (see above) are resuspended in 30% (v/v) acetic acid, homogenized, sonicated, and then centrifuged at 100,000 g for 45 min at 4~ The acid supernatant is subjected to molecular sieve fractionation on a Sephadex G-50 column both equilibrated and eluted with 10% (v/v) acetic acid. One-milliliter fractions are collected, and the S-14 and S-28(1-12)-like peptides are quantitatively evaluated using specific radioimmunoassays as described (44, 45). The antibody 20-44 directed against the COOH-terminal portion of S-14 (provided by C. Rougeot, Pasteur Institute, Paris) recognizes the somatostatin precursor, S-28, and S-14, and is used at a 1 980,000 dilution as described. The antiserum S-320 (donated by R. Beno~t, Montreal, Canada) is used at a final dilution of 1:9000 to detect pro-S(1-76) and S-28(1-12).

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Blockage of lntracellular Transport Two treatments (27) are performed: incubation of cortical fragments with (a) monensin or (b) at reduced temperatures (19~ In (a), cortex fragments are cut until they pass through a Pasteur pipette and are placed in Krebs-Ringer bicarbonate, pH 7.2 (Sigma). They are preincubated for 15 min at 37~ under a 5% CO2/95% 02 atmosphere and for 2 hr with 100 /zM monensin (Calbiochem, La Jolla, CA). After incubation, fragments are rinsed with Krebs buffer and then with 0.32 M sucrose and subjected to subcellular fractionation as described above. In (b), cortex fragments as in (a) are incubated at 19~ in Krebs-Ringer bicarbonate with 50 mM cycloheximide under 5% CO2/95% 02 and agitation for different periods of time and subsequently homogenized and subfractionated as described above. Results Together these approaches unequivocally established that both monobasic and dibasic cleavages occur in the Golgi apparatus of these cells, possibly in the trans pole of these stacks (Fig. 11). These conclusions were reinforced by two independent approaches. First, the process of trans-Golgi budding was impaired by a drug (monensin) and the recovered Golgi apparatus from the treated cells was analyzed for prosomatostatin and its maturation products (27). Second, the primary culture of these cells was kept at 19~ a temperature shown to slow down dramatically, but reversibly, the formation of the trans-Golgi network (TGN) vesicles. Prosomatostatin and corresponding processing products were analyzed similarly by HPLC/RIA in extracts of the purified Golgi fractions. The data from both approaches allowed similar conclusions: although TGN budding was blocked, prosomatostatin conversion in both S-14 and S-28 could be shown to occur (see Fig. 12). The appearance of the latter peptide was noticeably delayed in time (27).

Anglerfish Pancreatic Islets Bourdais et al. (28) have studied anglerfish PSII production and maturation in Brockmann organs of L. piscatorius. In this case, a peptide called Bella, corresponding to the C-terminal motif of the proregion of PSII is synthesized and polyclonal antibodies against this region are raised (Fig. 13) This antibody, called anti-Bella, does not cross-react with intact prosomatostatin but only with the C-terminal sequence of PSII(1-72) (Fig. 10) after this epitope is unmasked. Therefore this immunochemical tool could be used to reveal cleavage occurring at the level of A r g 73 in the 101-amino acid residue precursor (PSII) and leading to the (1-72) fragment plus S-28 II. This antibody is useful to reveal, first, that only a minor proportion of the somatostatin

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

185

producing cells is labeled (--~10% of the total) (Fig. 11); second, that immunogold labeling, using the gold-protein A technique, indicates the unmasking of the corresponding Bella epitope in the TGN and in the dense core granules (Fig. 14). Together these data demonstrate that monobasic cleavage of prosomatostatin II in a small percentage of pancreatic islet cells of the Brockmann organs from fish, generates S-28 II and that this cleavage initiates in the TGN of these cells. Experimental Procedure Four antibodies are used in this study (see Fig. 10); two are directed against somatostatin-14. Another is raised against synthetic AF-S(1-12) I peptide, that is, AlalAlaSerGlyGlyProLeuLeuAlaProArgGlu 12. The fourth is directed against a synthetic peptide named Bella, corresponding to AF-pro-S(64-72) II sequence, that is, Ala64ThrGluGlyArgMetAsnLeuGlu 72. Both peptides and Tyr o Bella are synthesized following a modification of the solid-phase method (12). Their purity is checked by routine procedures [HPLC, amino acid composition, and fast atom bombardment (FAB) spectroscopy]. AF-S(1-12) I peptide is coupled to thyroglobulin and Bella to bovine serum albumin, respectively (28), using a ratio of 10"1 (peptide'protein). The conjugates (100 txg) are mixed with an equal volume of Freund' s complete adjuvant and injected into the foot pads of female outbred Bouscat white rabbits, weighing 2-3 kg and obtained from the Institut Pasteur (Paris, France). Animals are given a booster injection after 15 days and subsequently at monthly intervals until an acceptable titer is attained. Immunofluorescence and immunogold labeling for electron microscopy are both performed on cryosections (1-1xm average thickness) (28). Immunogold labeling is also applied to thin sections of Epon-embedded material. In this case, the samples are fixed in 2% (v/v) glutaraldehyde in cacodylate buffer (0.2 M), pH 7.4, for 30 min and then in 1% (v/v) osmium tetroxide, prior to dehydration and inclusion steps. The thin sections are bleached by sodium metaperiodate for 10 min and immunogold labeling is performed (28).

Pro-ACTH/Endorphin System The intracellular sites where the common precursor for ACTH and endorphin (POMC) is processed have been identified by using a cytoimmunochemical approach based on purified antibodies that recognize POMC products exclusively (29). Several immunochemical tools were developed, including specific antibodies to detect an epitope unmasked at the C terminus of the ACTH

186

I1 IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

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FIG. 12 Prosomatostatin maturation in rat brain cortical cells after "T ~ block" at 19~ The effects of intracellular transport blockage by reduced temperature were evaluated on rat cortical fragments. Golgi-enriched fractions were collected and the proportions of various fragments of the precursor were measured by RIA. [From Lepage-Lezin et al. (27).]

molecule and an a-amidated carboxy group generated at the C-terminal end of the pro-POMC joining peptide(77-94) (Fig. 15). These were able to recognize mature ACTH, but also its glycosylated form or its biosynthetic intermedaite, that is, the POMC molecule deleted at its C-terminal end of the entire fl-lipotropin domain (see Fig. 15). Because the intact precursor was not detectable by these antibodies the technique allowed discrimination between POMC and its processing products, resulting from dibasic cleavage at the Lys137-Arg 138 site (see Fig. 15). The antibodies were used in cytoimmunochemistry and performed on AtT-20 cells taken as models for POMC processing.

FIG. 11 Immunogold labeling of the Golgi-enriched fraction with anti-S28(1-12) serum. In the absence of permeabilization gold particles were visible at the external surface of the Golgi vacuoles and vesicles [arrows in (a)-(c)]. When Triton X-100 permeabilization was applied [(d)-(g)] gold particles were more abundant but still mainly associated with the outer surface of the membrane profiles. Bar: 100 nm. [From Lepage-Lezin et al. (27).]

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FIG. 13 Immunofluorescence staining of Brockmann organs. Cryosections with (A) anti-S-14, (B) anti-Bella, and (C) anti-S-28(1-12) antisera. Bar: 30 ~m [for (A) and (C)] and 20 ~m for (B). [From Bourdais e t a l . (28).]

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FIG. 14 Immunogold labeling of Brockmann bodies. Ultrathin sections with antiserum directed toward Bella. Large granules containing cells are abundantly stained with gold particles (a) and in these cells the trans-Golgi network profiles are also labeled [arrows in (b)]. [From Bourdais et al. (28).]

190

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES Pro-optomelanocorttn

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139

176

6-endorphln

179

20g

Y

Ab Jamle

FIG. 15 pro-ACTH/endorphin processing. A schematic representation ofthe various POMC fragments generated by proteolytic processing in AtT-20 cells; Ab Jamie and Ab Kathy represent the various selective antibodies used for detection of the fragments as shown. [Adapted from Schnabel et a/.(29).] 9 1989 The Endocrine Society.

Experimental Procedure

Details can be found in Schnabel et al. (29). Preparation and Culture of AtT-20 Cells AtT-20/D-16v cells are grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% (v/v) horse serum in an atmosphere of 95% air/5% CO2 and are replated at 5-day intervals when they reach a density of approximately 5 x 105/flask. For individual experiments they are plated on 35-mm plastic tissue culture dishes and cultured for 4-5 days. For morphological studies, AtT-20 cells are fixed for 1 hr at room temperature with 2% (v/v) glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4. They are then postfixed at 4~ with 1% (v/v) osmium tetroxide in ace-

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

191

tate-veronal buffer (1 hr), stained in block with 0.5% (v/v) uranyl acetate in the same buffer (1-2 hr), dehydrated in ethanol, and embedded in Epox. All fixation and embedding steps are carried out in the 35-mm culture dish. After polymerization overnight (60~ pieces of the embedded cell layers are broken out of the culture dish, mounted on Epox blocks, and ultrathin sections (60-80 nm) cut on a Reichert Utracut E ultramicrotome. Sections are stained with 2% (w/v) uranyl acetate and lead citrate, and micrographs are taken on a Philips 301 electron microscope operated at 80 kV. Cells to be processed for immunofluorescence labeling of semithin (0.5-1 ~m) frozen sections and immunogold labeling of ultrathin frozen sections are harvested by scraping into aldehyde fixative [3% (v/v) paraformaldehyde, 0.05% (v/v) glutaraldehyde in 0.1 M phosphate buffer]. They are then pelleted in a Beckman (Fullerton, CA) microfuge for 10 min, and fixation of the pellet is continued for a total of 1 hr. The pellets are cryoprotected in sucrose and frozen in liquid nitrogen. Immunogold Labeling of Ultrathin Frozen Sections Ultrathin cryosectioning of AtT-20 cell pellets and rat pituitary tissue is carried out on a Reichert Ultracut E equipped with a cryoattachment at -110~ following the techniques of Tokuyasu [referenced in (29)]. Sections are transferred to hexagonal nickel grids (2090 mesh) that have been coated with Formvar and carbon. Subsequent incubations and washing steps are carried out by floating the grids on droplets of the filtered solutions. After quenching with 10% (v/v) fetal bovine serum (FBS) containing 0.01M glycine (to block free aldehyde groups), the sections are incubated for 1 hr with one of the above-described primary antibodies [diluted in phosphate-buffered saline (PBS) with 10% (v/v) FBS] followed by 5-nm colloidal gold conjugated to goat anti-rabbit IgG (diluted 1:50). For double immunolabeling (see Table III) the sections are first incubated for 1 hr with Kathy (specific for processed ACTH) followed by incubation in 10-nm protein A-gold conjugate. Sections are then quenched with free protein A (0.1 mg/ml) and incubated with Danielle (recognizes the precursors as well as the processed products) or with Jamie (specific for the c~-amidated C-terminal of JP) followed by 5-nm protein A conjugate. After labeling, the sections are postfixed in 2% (v/v) glutaraldehyde in PBS (10 min), stained with 2% (v/v) osmium tetroxide (15 min), followed by 2% (v/v) acidic uranyl acetate (15 min), and finally absorption stained (5 min) with 0.002% (v/v) lead citrate in 2.2% (v/v) polyvinyl alcohol as described by Tokuyasu [see Schnabel et al. (29)]. Results

The results (29) of these studies demonstrated the following: POMC-related peptides were found in the Golgi, fully processed (and amidated) ACTH

192

II

IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

TABLE III

Specificities of Pro-ACTH/Endorphin Antibodies a

Antibody

POMC

fl-LPH

ABI

Danielle Rebecca

+ +

+ +

. .

Georgie Kathy Jamie

+ .

-

+ +

.

.

.

ACTH . .

. .

+

16K

nJP

. .

fl-Endorphin + +

+ +b

+ +

_

Details in Schnabel et al. (29). o Positive binding ( + ) has been defined by immunoprecipitation by cross-reaction in RIA at antibody concentrations comparable to those used for immunocytochemistry, by graded abolition of immunostaining in the presence of 10-200 mM peptide and failure of irrelevant peptides to block staining. Lack of binding ( - ) indicates lack of immunoprecipitation at high antibody concentration, and failure to show any signal in RIA. Jamie detects POMC(1-94)NH2 but not POMC(1-95), both of which are in the 16K pool originally defined by use of antibody Georgie [details in Schnabel et al. (29)].

a

and the A C T H biosynthetic intermediate were detected in the trans-Golgi cisterna; the use of an antibody selectively directed against the a-amidated C terminus of the joining peptide showed that this epitope could be detected in the dilated rims of the trans-most Golgi cisterna and was not restricted to the secretory granules. It should be noted that the formation of this latter processing product requires a cascade of reactions, including endoproteolysis at two dibasic sites (Arg75-Arg 76 and Lys96-Arg97), then exoproteolysis, and finally a-amidation of the C terminus. These data conclusively demonstrate the occurrence of endoproteolytic maturation in a late compartment of the Golgi apparatus, a finding corroborated by similar observations made on rat pituitary corticotropes.

Acknowledgments This work was supported in part by funds from the Universit6 Pierre et Marie Curie and the Centre National de la Recherche Scientifique to URA 1682. We wish to thank Sandrine Cadel for reading and Brigitte Sockeel for typing the manuscript.

References 1. N. Darby and D. Smyth, Biosci. Rep. 10, 1 (1990). 2. M. Rholam, P. Nicolas, and P. Cohen, FEBS Lett. 207, 1 (1986). 3. M. Rholam, N. Brakch, H. Boussetta, and P. Cohen, submitted for publication (1994).

[9] IDENTIFYING NEUROPEPTIDE-PROCESSING PATHWAYS

10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30.

193

E. Bek and R. Berry, Biochemistry 29, 178 (1990). D. Julius, A. Brake, L. Blair, R. Kunisawa, and J. Thorner, Cell (Cambridge, Mass.) 37, 1075 (1984). K. Mizuno, T. Nakamura, T. Ohshima, S. Tanaka, and H. Matsuo, Biochem. Biophys. Res. Commun. 156, 246 (1988). C. Clamagirand, Biochem. Biophys. Res. Commun. 156, 246 (1987). M. Camier, D. Benveniste, N. Barre, N. Brakch, and P. Cohen, Mol. Cell. Endocrinol. 282, 363 (1990). C. Clamagirand, M. Camier, H. Boussetta, C. Fahy, A. Morel, P. Nicolas, and P. Cohen, Biochem. Biophys. Res. Commun. 134, 1190 (1986). C. Clamagirand, C. Cr6minon, C. Fahy, H. Boussetta, and P. Cohen, Biochemistry 26, 6018 (1987). C. Camier, R. Alazard, P. Cohen, P. Pradelles, J. Morgat, and P. Fromageot, Eur. J. Biochem. 32, 207 (1973). P. Nicolas, A. Delfour, H. Boussetta, A. Morel, M. Rholam, and P. Cohen, Biochem. Biophys. Res. Commun. 175, 565 (1986). N. Brakch, H. Boussetta, M. Rholam, and P. Cohen, J. Biol. Chem. 264, 15912 (1989). I. Plevrakis, C. Clamagirand, C. Cr6minon, H. Boussetta, N. Brakch, M. Rholam, and P. Cohen, Biochemistry 28, 2705 (1989). K. Mizuno, T. Nakamura, K. Tanada, S. Sakakibara, and H. Matsuo, Biochem. Biophys. Res. Cornmun. 144, 807 (1987). C. Brenner and R. Fuller, Proc. Natl. Acad. Sci. U.S.A. 89, 922 (1992). P. Gluschankof, S. Gomez, A. Morel, and P. Cohen, J. Biol. Chem. 262, 9615 (1987). J. A. Norman and J. Y. Chang, J. Biol. Chem. 260, 2653 (1985). N. Brakch, M. Rholam, H. Boussetta, and P. Cohen, Biochemistry 32, 4925 (1993). L. Paolillo, M. Simonetti, N. Brakch, G. D'Auria, M. Saviano, M. Dettin, M. Rholam, A. Scatturin, C. Di Bello, and P. Cohen, EMBO J. 7, 2399 (1990). P. Kuks, C. Cr6minon, A. M. Leseney, J. Bourdais, A. Morel, and P. Cohen, J. Biol. Chem. 264, 14609 (1989). M. C. Beinfeld, J. Bourdais, P. Kuks, A. Morel, and P. Cohen, J. Biol. Chem. 264, 4460 (1989). J. Bourdais, A. Pierotti, H. Boussetta, N. Barre, G. Devilliers, and P. Cohen, J. Biol. Chem. 266, 23386 (1992). A. Morel, P. Nicolas, and P. Cohen, J. Biol. Chem. 258, 8273 (1983). Y. Peng Lob, M. Brownstein, and H. Gainer, Annu. Rev. Neurosci. 7, 189 (1984). L. Orci, M. Ravazzola, M. Storch, R. Anderson, J. Vasali, and A. Perrelet, Cell (Cambridge, Mass.) 49, 865 (1987). A. Lepage-Lezin, P. Joseph-Bravo, G. Devilliers, L. Bendetti, J.-M. Launay, S. Gomez, and P. Cohen, J. Biol. Chem. 266, 1679 (1991). J. Bourdais, G. Devilliers, R. Girard, A. Morel, L. Benedetti, and P. Cohen, Biochem. Biophys. Res. Commun. 170, 1263 (1990). E. Schnabel, R. Mains, and M. Farquhar, Mol. Endocrinol. 3, 1223 (1989). R. Chance, R. Ellis, and W. Bremer, Science 161, 165 (1968).

194

II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES 31. J. Roth, M. Bendayan, and L. Orci, J. Histochem. Cytochem. 26, 1074 (1987). 32. M. Storch, K. Petersen, T. Licht, and L. Kerp, Diabetes 34, 808 (1985). 33. L. Orci, M. Ravazzola, M. Amherdt, O. Madsen, J. Vasali, and A. Perrelet, Cell (Cambridge, Mass.) 42, 671 (1985). 34. P. Lacy and M. Kostianovsky, Diabetes 16, 35 (1967). 35. L. Orci, M. Ravazzola, M. Amherdt, O. Madsen, A. Perrelet, J. Vasali, and R. Anderson, J. Cell Biol. 103, 2273 (1986). 36. H. Davidson, C. Rhodes, and J. Hutton, Nature (London) 333, 93 (1988). 37. J. Trifaro and A. Duerr, Biochim. Biophys. Acta 421, 153 (1976). 38. C. de Duve, R. Wattiaux, and P. Baudhin, Adv. Enzymol. 24, 1291 (1962). 39. D. Louvard, H. Reggio, and G. Warren, J. Cell Biol. 92, 92 (1982). 40. D. Morr6, in "Methods in Enzymology" (W. Jakoby, ed.), Vol. 22, p. 130. Academic Press, New York, 1971. 41. U. Laemmli, Nature (London) 227, 680 (1970). 42. H. Towbin, J. Staehelin, and J. Gordon, Proc. Natl. Acad. Sci. U.S.A. 76, 4350 (1979). 43. I. Dunia, S. Manenti, A. Rousselet, and E. Benedetti, J. Cell Biol. 105, 1679 (1987). 44. P. Gluschankof, A. Morel, R. Benoit, and P. Cohen, Biochem. Biophys. Res. Commun. 128, 1051 (1985). 45. M. Lauber, M. Camier, and P. Cohen, Proc. Natl. Acad. Sci. U.S.A. 76, 6004 (1979). 46. C. Clamagirand, M. Camier, C. Fahy, C. Clavreul, C. Creminon, and P. Cohen, Biochem. Biophys. Res. Commun. 143, 796 (1987).

[10]

Immunological and Related Techniques for Studying Neurohypophyseal Peptide-Processing Pathways Harold Gainer, Mark O. Lively, and Mariana Morris

Introduction The neurohypophyseal peptides, oxytocin and vasopressin, are synthesized as part of larger precursor proteins that are subsequently processed into the final peptide products (1-3). Figure 1 illustrates the precursor (prohormone) structures and the anticipated peptide intermediates for oxytocin and vasopressin. Proteolytic and other processing of these prohormones appears to occur in the neurosecretory granules (4), and a preliminary evaluation of the processing enzyme genes being expressed in the hypothalamoneurohypophyseal system (HNS) has been reported (5). Intact precursors can be isolated from hypothalamic regions containing the neuronal perikarya whereas, as a rule, only fully processed products are found in the nerve terminals in the neural lobe of the pituitary (6-8). In this chapter we describe the development and availability of antibody reagents that, in combination with various separation techniques, can detect and quantify the intermediate peptide forms shown in Fig. 1 for oxytocin (OT) and vasopressin (AVP).

Antibody Reagents" Production and Specificities Most anti-OT and anti-AVP antibodies are directed against the amidated form of these peptides, because these are the peptide antigens that historically were used in the immunizations. In addition, most of the conjugations of these peptides to carrier proteins were previously done using glutaraldehyde as the coupling agent, further ensuring that the antibody would be C-terminally directed. Examples of such antibodies are OT-MM (9) (developed by M. Morris for amidated OT; NIAAA, Bethesda, MD), and AVP-JR and AVP-ES [developed by J. Russell (NICHD, Bethesda, MD) and R. Eskay, respectively, for amidated AVP]. It is possible to bias the epitope of the antibody to the N terminal peptide by coupling the free carboxyl group to the carrier, as was done for the VA series of OT and AVP antibodies Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All fights of reproduction in any form reserved.

195

196

II

I M M U N O L O G I C A L AND BIOCHEMICAL APPROACHES

OXYTOCIN Prloul'lolr Peptide Intermediates

! OT I'Gly-Lys-Arg-i 1 9 13 ! OT ]-Gly-Lys-Aig 1 9 1

Final Peptidos

Tryptlc

Cloavago Products

Np-OT

i'Arg 105

Np-OT

I'A~ 105

o

! OT l~v

1 9 ! OT I-NH2 1 9

loT I-eu

1

I 13

9

l

20

Np.OT

]-~

VASOPRESSIN Precursor Peptide Intermediates

I

] Final Peptides Tryptio Cleavage Products

Np-AVP

AVP EGly-Lys-Arg'I 9 13 ! AVP I-Gly-Lys-Arg 1 9

1

I-A~{

105 107

~

]

145

9

! AVE,I~

1 9 I AVP ]-NH'2 1 9 ~-] 1 8

13

! 20

Np-AvP

105 107

I

145

98

FIG. 1 Structures of oxytocin and vasopressin precursors (prohormones), the fully processed (final) peptides found in vivo, and theoretical peptide hormone intermediates resulting from precursor processing. Also shown are the products of in vitro incubation of the precursors with trypsin. Abbreviations: OT, oxytocin; AVP, vasopressin; Np-OT, oxytocin-associated neurophysin; Np-AVP, vasopressin-associated neurophysin; GP, glycopeptide; AVP(1-8), deglycinamide-AVP. (Table I) (6, 7). The latter procedure has been derived from Moore et al. (10), and is briefly described below. Peptides are conjugated to high molecular weight carriers as follows: Peptide (1-3 mg) and carrier [3 mg of either keyhole limpet hemocyanin (KLH), bovine serum albumin (BSA), ovalbumin, or thyroglobulin] are dissolved in 450/zl of distilled water and 300/zl of an aqueous solution (3.5 mg/ml) of 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) is added. The mixture is incubated at room temperature for 18 hr, and is terminated by 750/zl of 0.05 M ethanolamine followed by an incubation of 5 hr. The peptide is emulsified in 0.5 ml of Freund's adjuvant and injected (0.5 ml/rabbit)

197

[10] N E U R O H Y P O P H Y S E A L PEPTIDE PROCESSING

TABLE I Cross-Reactivities of Anti-Oxytocin and Vasopressin Antibodies with Oxytocin and Vasopressin Intermediate Forms Relative displacement factor (%)a OT-specific antibodies

Peptide

OT (amidated) OT-Gly 1~ OT-Glyl~ ~1 OT Gly1~ AVP (amidated) AVP-GIy ~~ AVP-Glyl~ I~ AVP-Glyl~ AVT (vasotocin) MT (mesotocin)

~2

~2

VP-specific antibodies

OT-MM (1 : 100,000)

OT-VA17 (1:60,000)

AVP-JR1 (1:45,000)

AVP-VA4 (1 : 16,000)

100 0.01 0.01 <0.01 0.01

100 87 35 78 <2

0.02

<2

nt nt nt

nt nt nt

nt nt

nt nt

nt 0.04

<2 <2

100 <0.4 <0.4 <0.4 241

nt

43

nt

100 46 76 51 <2 <2

" Relative displacement factor represents the ratio (in percent) of the peptide concentration required to displace 50% OF [~25I]AVP OR [125I]OT (amidated forms) from a given antibody versus the unlabeled amidated peptide ligand (set at 100%). Cross-reactivity was determined by conventional radioimmunoassay procedures using 6-10 dilutions of hormone or analog, in duplicate. The numbers in parentheses indicate antibody dilutions used in the assays, nt, Not tested. [Data from Altstein e t al. (6) and Altstein and Gainer (7).]

subdermally at multiple sites along the back. Rabbits are injected every 2 weeks for 6 weeks and then every 6 weeks for an additional period of 3 months. The first immunization is carried out in Freund's complete adjuvant, using KLH as a carrier protein. All boosts are carried out using Freund's incomplete adjuvant and in the sequence of BSA, ovalbumin, thyroglobulin, KLH, and thyroglobulin as carrier proteins. Any further boosts utilize KLH again as the carrier protein. The first bleed is collected 52 days after the initial immunization and successive bleeds are drawn at 32- and 52-day intervals. Antisera are aliquoted and stored at -20~ containing 3 mM sodium azide as a preservative. We have found that these variations in carrier proteins are effective in the generation of high-titer peptide antisera. The antibodies are characterized by standard radioimmunoassay (RIA) techniques using peptides iodinated by the Iodogen method (11) and purified by high-performance liquid chromatography (HPLC) using a C~8 reversedphase column. Briefly, 12 serial dilutions (starting from 1 9500) of each antisera are made in 0.1 M sodium phosphate buffer containing BSA (1.5 mg/ ml), pH 7.4 (RIA buffer) and 500/~1 of each dilution is incubated with 100 ~zl of 125I-labeled homologous tracer diluted in Krebs-Ringer solution [125

198

II IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

mM NaC1, 4.5 mM KC1, 1.5 mM MgSO4, 2.0 mM CaC12, 2 mM HEPES, and BSA (1.5 mg/ml), pH 7.4] containing approximately 10,000 cpm. Reactions are incubated for 48 hr at 4~ and are terminated by the addition of 1.26 ml of cold ethanol (68% final concentration). The bound complex is separated from the free tracer by centrifugation at 1500 g for 30 min at 4~ The supernatant is discarded and the pellet (bound fraction) is counted for radioactivity. Nonspecific binding (3-5%) is determined in the absence of antibody and is subtracted from every tube. Data are plotted as a percentage of total counts bound. The dilution that provides 30-60% binding is typically used. Cross-reactivity is determined from RIA curves. Reaction mixtures containing 100 /xl of 125I tracer (approximately 10,000 cpm) diluted in Krebs-Ringer solution, 100/zl of hormone or analog, and 400/xl of antisera (at a dilution that provides 30-60% tracer binding), all in a total volume of 600/zl of RIA buffer, are incubated under equilibrium conditions for 48 hr at 4~ Incubations are terminated as described above, and data are plotted as percent binding versus peptide concentration. Cross-reactivity is determined by comparing the peptide concentration that displaces 50% of a labeled tracer. Table I illustrates the characterization of OT- and AVP-specific antibodies that are C-terminally directed and also amidation specific (i.e., OT-MM and AVP-JR1, respectively) and N-terminally directed antibodies that are highly specific for these two peptides and can detect all the intermediate forms (i.e., V A -17 and V A -4 for OT and AVP, respectively). The latter antibodies can be used in conjunction with various separation techniques (see below) to measure all the intermediate forms. Alternatively, the same immunization protocol described above, but using glutaraldehyde coupling of the specific (C-terminally extended) peptide forms to the carrier, could be used to generate specific antibodies to the specific extended forms. There are obvious advantages and disadvantages to both approaches. Antibodies to a specific extended form could be used directly in an RIA procedure in one step. In contrast, the pan-specific antibodies measure all forms, and hence require more effort to evaluate the contributions of a given form.

Extraction of Peptides from Tissues Tissues are rapidly dissected, quickly frozen on dry ice or liquid nitrogen, and then transferred to precooled glass-glass homogenizers containing 0.1 N hydrochloric acid (HC1) and homogenized manually. Samples are centrifuged at 12,100 g for 15 min at 4~ and the supernatant (S1) is used for the analysis of fully processed forms of AVP and OT by means of RIA. The remaining sample is treated with trichloroacetic acid (TCA, 10T final

[10] NEUROHYPOPHYSEAL PEPTIDE PROCESSING

199

concentration) for 2 hr on ice and spun at 10,000 g for 10 min at 4~ The resulting supernatant ($2) is ether washed, lyophilized, resuspended in half the original volume of 0.1 M sodium phosphate buffer containing BSA (1.5 mg/ml), pH 7.4 (RIA buffer), and assayed for fully processed and partially processed (C-terminally extended) forms of AVP and OT by means of RIA and high-voltage electrophoresis (HVE) chromatography (see the next section). The pellet (P2) containing putative unprocessed prohormone is ether washed, air dried, dissolved in either 2.0 ml (brain) or 0.14 ml (pituitary) of 10 mM Tris-HC1, pH 8.0 (trypsinization buffer), and incubated in the presence or absence of trypsin (10/zg/ml) for 2 hr at 37~ Incubation is terminated by tasyl lysyl chloromethyl ketone (TLCK; 0.025 mg/ml), and samples are assayed for tryptic cleavage products by means of RIA.

Separation Methods" High-Voltage Electrophoresis Because the peptide intermediates differ from one another by various charged groups, that is, free carboxyl versus amidated, additional basic amino acids (lysine or arginine) (see Fig. 1), high-voltage electrophoresis (HVE) of the OT and AVP intermediates offers a rational choice for their separation. In fact, such separations are easily done with the neurohypophyseal peptides and their intermediates, and detection using the antibodies described in Table I is effective. Figure 2 illustrates such a standard run for the OT- and AVP-related peptides. The procedure is as follows. The TCA-treated supernatants of the tissues (described above) are partially purified on a Sep-Pak C reversed-phase column. Samples are applied onto the column and washed with 5 ml of 0.1% trifluoroacetic acid (TFA) to remove nonpeptide contaminants. The peptides are eluted from the Sep-Pak by two 1-ml washes of 80% methanol containing 0.1% TFA. The eluate is dried, reconstituted in 20 txl of 0.1 N HC1, and analyzed by HVE on cellulose plates in a pH 2 system (formic acid-acetic acid-water, 20:80:900). Electrophoresis is carried out at 900 V and the migration front (15.5 cm from origin) is detected using a methyl green marker. The plate is then cut into 1-cm fractions beginning 1 cm below the origin, peptides are extracted from the cellulose with 500 Ixl of 0.1 N HC1, and all fractions are assayed for immunoreactive material by means of RIA as described above. Standard peptides (AVP, OT, and their C-terminally extended forms) are run as markers on adjacent lanes and visualized by fluorescamine and by RIA. Sample recovery is evaluated by using synthetic AVP and OT, and is >98% for the Sep-Pak column and 20% for HVE. Applications of these techniques to rat brain tissues are illustrated in Figs. 3 and 4, and are discussed elsewhere (6, 7).

200

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

~)CO < 250

T II II II II II

"~ 2oo .Q

; c E E

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High-voltage electrophoresis (HVE) of AVP (left), OT (right), and their C-terminally extended analogs. A mixture of amidated and C-terminally extended peptides (10/.r each, dissolved in 20/~1 of 0.01 N HC1) was applied on the HVE plate and electrophoresed as described in text. At the end of the run 1-cm fractions were cut, eluted with 500/zl of 0.1 N HCI, and 0.10 ~1 of each cellulose-HC1 extract was analyzed for amidated or C-terminally extended immunoreactivity by means of RIA, using [125I]AVP or [125I]OT as a tracer. Migration of AVP or OT (open circles) was determined using AVP-JR1 (1 : 45,000) and OT-MM (1 : 100,000) antisera, respectively, and migration of the extended forms, AVP-X and OT-X (filled circles) was determined using AVP-VA4 (1 : 16,000) and OT-VA17 (1 : 60,000) antisera, respectively. Single-letter amino acid abbreviations: G, glycine; K, lysine; R, arginine. [From Peptides 9, Alstein et al. (6), copyright 1988, with kind permission from Elsevier Science, Ltd., The Boulevard, Langford Lane, Kidlington OX5 1GB, U.K.] FIG. 2

Separation Methods" High-Performance Liquid Chromatography and Radioimmunoassay The use of H P L C separation coupled with specific RIAs provides another method for the identification and quantitation of the products of peptide processing. This methodology has proved useful in the study of the development of peptide systems in the fetal sheep (12).

[10]

NEUROHYPOPHYSEAL

250

500-

?o

/

~.

~400-

.~,

300-

rt

~-'

~-'

0

0

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PEPTIDE PROCESSING

;?

[.oo ~

A

0

0

!-,so

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100

200-

100-

0

w 5

I I0

I 15

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;

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I 15

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so

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FIG. 3 Extracts of adult brain (hypothalamus; left) and neural lobe (of the pituitary; right) were analyzed by HVE for fully (OT, open circles) and partially processed (OT-X, closed circles) forms of OT. Hypothalamic (0.1 ml) and posterior pituitary (10 /zl) supernatants ($2) were passed through a Sep-Pak column, eluted, dried, resuspended in 20/xl of 0.01 N HC1, and applied to an HVE plate for electrophoresis as described in text. Peptides were eluted with 500/xl of 0.1 N HC1 and 50/xl of each cellulose-HC1 extract was analyzed for OT or OT-X immunoreactivity as described in the caption to Fig. 2. Oxytocin content was determined from OT-MM immunoreactivity and OT-X content was calculated from the difference between the immunoreactivity of OT-VA17 and OT-MM. Markers (depicted by arrows) were visualized by fluorescamine and by means of RIA. OT-X, C-Terminally extended OT. [From Peptides 9, Alstein et al. (6), copyright 1988, with kind permission from Elsevier Science, Ltd., The Boulevard, Langford Lane, Kidlington OX5 1GB, U.K.]

P l a s m a and T i s s u e E x t r a c t i o n The plasma or tissue samples are subjected to an initial Sep-Pak extraction. C~8 Sep-Pak cartridges (Waters Chromatography, Milford, MA) are washed with ! 0 ml of 100% methanol followed by 10 ml of distilled water. The plasma (5 ml) or the acid extract of the tissues (15 ml) is applied slowly to the column, using a 10-ml syringe. The column is washed immediately with 20

202

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~

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10

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FIG. 4 Brain (left) and neural lobe extracts (right) (postnatal days 7-21) analyzed by HVE for fully and partially processed forms of OT. Brain and pituitary supernatants were applied on an HVE plate, after a Sep-Pak step, and electrophoresed as described in the caption to Fig. 2, except that in this experiment a 100-/xl aliquot (brain) and a 2-/xl aliquot (neural lobe) of the 500-/xl cellulose-HCl extract were analyzed for OT and OT-X immunoreactivity. Oxytocin immunoreactivity (dashed line) was determined using OT-MM antiserum (1 : 100,000), and OT-X immunoreactivity (solid line) was calculated from the difference between the immunoreactivities of OT-VA17 (1 : 60,000) and OT-MM. Electrophoretic positions of standards (arrows) were visualized by fluorescamine and measured by means of RIA as described in text. The value of OT-GK was corrected for the cross-reactivity of anti-OT-VA17 with OT-GK. Abbreviations: OT-G, OT-glycine; OT-GK, OT-glycine-lysine; OT-GKR, OT-glycine-lysine-arginine. [From Altstein and Gainer (7).]

ml of cold 4% acetic acid, followed by 5 ml of distilled water. The peptides are eluted with 4 ml of 75% (v/v) acetonitrile-25% (v/v) of 4% acetic acid. The eluant is lyophilized and stored at -70~ The plasma is applied directly to the column while the tissue samples are sonicated in cold 0.1 N HC1, with a ratio of 10" 1 solvent to tissue. The extract is centrifuged at 3500 rpm for

[10] NEUROHYPOPHYSEAL PEPTIDE PROCESSING

203

20 min at 4~ with the supernatant further purified by the ion-exchange method described above.

High-Performance Liquid Chromatography Peptide separations are performed using an automated gradient chromatography system from Waters Chromatography (Division of Millipore, Milford, MA). This system consists of a model 845 chromatography workstation with a VaxStation 4000 computer for control and data collection. The chromatograph is equipped with a WISP model 712 automated sample injector, two model 510 pumps, a model 484 variable-wavelength detector, and a temperature control oven for the columns. Peptides and tissue extracts are separated using reversed-phase chromatography with a trifluoroacetic acid (TFA)-CH3CN system on a Spheri-5 RP-18 column (4.6 • 230 mm, 5-mm C18 silica particles; Applied Biosystems, Inc., Foster City, CA). Standard samples of authentic OT-amide (Bachen California, Torrance, CA), OT-GlyLys (H. Gainer, NIH, Bethesda, MD), and OT-Gly-Lys-Arg (synthesized in our laboratory) are eluted using the following gradient. The flow rate is constant at 1 ml/min with ultraviolet (UV) detection at 215 nm. The column is initially equilibrated in 100% solvent A (0.1% aqueous TFA). Following injection, the mobile phase is maintained at 100% solvent A for 5 min. From 5 to 12 min, the concentration of solvent B (70% CH3CN in 0.1% TFA) is increased linearly to 7%, then to 35% over the following 40 min. Fifty-two minutes after injection, the concentration of solvent B in the mobile phase is increased to 100% over the next 3 min. Prior to injection, lyophilized extracts of tissue or plasma prepared using the Sep-Pak method are dissolved in 800 ml of 0.1% TFA, filtered through a Nylon 66 membrane (0.22-mm pore size; CentriFree, Millipore, Baltimore, MA). Fractions (1 ml) are collected and reduced to dryness by vacuum centrifugation. The fractions are redissolved in RIA buffer containing BSA (1 mg/ml), then analyzed by RIA. Table II shows the separation of peptide standards as monitored by UV detection (215 nm) and RIA of the HPLC fractions. Using a gradient HPLC system, there was a clear separation of the peptides; OT eluted first, followed by OT-GK and OT-GKR. OT-G was shown to migrate closely to OT in previous work. A similar HPLC and RIA combination can be used to separate the AVP family of amidated and C-terminally extended peptides. These methods have been used for the study of the oxytocin peptide forms present in fetal sheep plasma and hypothalamus (Figs. 5 and 6). In the late gestation fetal sheep (134 days, term of a approximately 142 days) there is evidence for the presence of three forms of oxytocin in the circulation. A comparison of the results with the two OT assays [OT-MM, which is specific

204

II IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES TABLE II

High-Performance Liquid Chromatography Separation of Oxytocin Peptide Standards a Peptide peak HPLC fraction number Peptide

OD

RIA

OT-NH2 OT-GK OT-GKR

49 53 56

50 54 58

Peptide standards were separated by a gradient HPLC method and evaluated by UV absorbance (215 nm) and RIA of the HPLC fractions. The HPLC fraction number for the peptide peak is provided. The separation system is the same as that described in text, using a C~8 silica column and a TFA-CH3CN gradient.

200 '

Z o

150 -

OT

OT-GK OT-GKR

I--o nLL 1 0 0 -

O

[]

50-

0

1

. . . . ~=J"' ' ~ ' ~ ' - ' - ~ ' - - ' ~ - i ~ - ~

8 16 24 3 2 4 D 4 5 4 7 4 9 5 1

53 55 57 59 61 63 65

HPLC FRACTION

FIG. 5 Measurement of OT forms in fetal sheep plasma. Plasma (5 ml) from a 134day sheep fetus was purified using a Sep-Pak C18 cartridge with acetonitrile-acetic acid elution. The lyophilized extract was resuspended in 0.1% TFA and separated by HPLC on a C18 column with a TFA-CH3CN gradient (described in text). The fractions were lyophilized, resuspended in RIA buffer, and measured by two RIAs with different specificities. OT-MM (11) is specific for OT, whereas OT-X ([2) recognizes OT and the C-terminal extended forms. [125I]OT was used as the tracer with the antisera used at final dilutions of 280K for OT-MM and 48K for VA-17 (OT-X). The arrows indicate the elution pattern of the peptide standards OT, OT-NH2, OT-GK (OT-glycine-lysine), and OT-GKR (OT-glycine-lysine-arginine).

[10]

NEUROHYPOPHYSEAL

PEPTIDE

OT ~ Z 0

205

PROCESSING

OT-GKR

i

6

o rr U..

0

4

2

1

8 16 24 32 4{) 45 47 49 51 53 55 57 59 61 63 65 HPLC FRACTION

FIG. 6 Measurement of OT forms in fetal sheep hypothalamus and posterior pituitary. The medial basal hypothalamus (0.6 g) and the posterior pituitary from a 134day-old sheep fetus were sonicated in 0.1 NHC1 and purified by C18Sep-Pak extraction and HPLC separation as described in the caption to Fig. 5. The HPLC fractions were lyophilized and measured by RIA using OT-X, which cross-reacts with OT and the C terminal-extended OT peptides. (n) Hypothalamus; ([]) posterior pituitary. The arrows indicate the elution positions of the peptide standards OT-NH2 and OT-GKR.

for amidated OT, and VA-17, which cross-reacts with the amidated and the C-terminal extended forms (OT-X)] shows that there was a single peak with OT-MM and three peaks with OT-X. The peak eluting at fraction 48 is amidated OT, with similar amounts measured with the two assays. The two later peaks are thought to be OT-GK and OT-GKR on the basis of the chromatographic migration of peptide standards. These HPLC-RIA data confirm and extend our previous results that demonstrated by assay subtraction methods that fetal sheep plasma contained high levels of the extended OT peptides (9, 12). The existence of these alternative OT peptides was first suggested by the work of Amico and colleagues (13-15). Using different OT antisera, they found evidence for the secretion of C-terminal extended OT during pregnancy and after estrogen stimulation. They reported that the primary plasma form in humans and primates was OT-G (15). However, the identity of the circulating OT form(s) may be questioned because the antisera used in this study could not detect OT-GK or OT-GKR. HPLC separation of fetal sheep hypothalamus and posterior pituitary revealed different patterns of peptide expression (Fig. 6). There were two peaks in the hypothalamic extract, comigrating with OT and OT-GKR. The levels of the amidated and extended peptide were essentially equal. The

206

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I M M U N O L O G I C A L AND BIOCHEMICAL APPROACHES

posterior pituitary showed one major peak that was coincident with OT. These results are consistent with studies in the rat (Fig. 4), which showed that the alternative OT forms, primarily OT-GKR, were present in the hypothalamus, but not the posterior pituitary. Although Amico did not detect the extended OT forms in a variety of primate tissues, this was likely the result of the use of an antiserum that was specific for OT-G (15). Indeed, there is strong evidence for the presence of the C-terminal extended OT peptides in peripheral tissues, including the ovary (16), corpus luteum (17), and thymus (18). We have also used HPLC separation and RIA quantitation to study the OT forms in peripheral tissues from fetal and maternal sheep. The results indicate that the adrenal, thymus, chorion, and amnion all contain the alternative OT forms, OT-GKR and OT-GK (unpublished data).

Conclusions In this chapter we have illustrated uses of antibodies that were specific for the neurohypophyseal peptides (OT or AVP) but did not distinguish between the intermediate and amidated forms of the peptides. Combined with an appropriate separation technique and RIA procedures, these antibodies could be used to distinguish between the intermediate forms. We have also found these antibodies useful for immunoprecipitation and immunocytochemical procedures (not illustrated). Another alternative, not described here, is to make peptide antibodies that are specific for each intermediate form of the peptide. For the OT and AVP peptides this should be relatively easy by linkage of the amino termini of these peptides to carriers (by glutaraldehyde, etc.) before immunization. Given such antibodies, assays of the intermediate forms could be performed in a single step, that is, by RIA or immunoprecipitation. The biological significance of the presence of stable "intermediate" forms of OT but not AVP peptides in three species (rodents, sheep, and primates) remains unclear at present. Is this due to the intrinsic nature (structure) of the OT-prohormone (versus the VP prohormone), to different convertases in the cells containing these peptides, or to different microconditions in subcellular organelles where the processing occurs? Future experiments using these and other techniques will be necessary to answer these questions.

Acknowledgments We would like to acknowledge the assistance of Drs. James Rose and K. Tsai. This work was supported by Grants HL43178 (M.M.) and HDll210 (J.R.).

[10] NEUROHYPOPHYSEAL PEPTIDE PROCESSING

207

References

.

,

8.

9. 10. 11. 12. 13. 14. 15. 16. 17. 18.

H. Gainer, Prog. Brain Res. 60, 205 (1983). R. Ivell, H. Schmale, and D. Richter, Neuroendocrinology 37, 235 (1983). E. Breslow and S. Burman, Adv. Enzymol. 63, 1 (1990). H. Gainer, J. T. Russell, and Y. P. Loh, Neuroendocrinology 40, 171 (1985). M. K. H. Sch~ifer, R. Day, W. E. Cullinan, M. Chr6tien, N. G. Seidah, and S. J. Watson, J. Neurosci. 13, 1258 (1993). M. Altstein, M. H. Whitnall, S. House, S. Key, and H. Gainer, Peptides (N. u 9, 87 (1988). M. Altstein and H. Gainer, J. Neurosci. 8, 3967 (1988). W. G. North, E. F. O'Conner, and C. B. Gonz~dez, Peptides (N. Y. ) 13, 395 (1992). M. Morris, S. W. Stevens, and M. R. Adams, Biol. Reprod. 23, 782 (1980). G. Moore, A. Lutterodt, G. Burford, and K. Lederis, Endocrinology (Baltimore) 101, 1421 (1977). P. R. P. Salacinski, C. McLean, J. E. Sykes, V. V. Clement-Jones, and P. J. Lawrey, Anal. Biochem. 117, 136 (1981). M. Morris, M. Castro, and J. C. Rose, Am. J. Physiol. (Regulatory Integrative Comp. Physiol.) 32, R738 (1992). J. A. Amico, M. G. Ervin, F. M. Finn, R. D. Leake, D. A. Fisher, and A. G. Robinson, Metab. Clin. Exp. 35, 596 (1986). J. A. Amico, M. G. Ervin, R. D. Leake, D. A. Fisher, F. M. Finn, and A. G. Robinson, J. Clin. Endocrinol. Metab. 60, 5 (1985). J. A. Amico, in "Recent Progress in Posterior Pituitary Hormones" (S. Yoshida and L. Share, eds.), p. 207. Elsevier, New York, 1988. M. D. Guillou, N. Barre, I. Bussenot, I. Plevrakis, and C. Clamagirand, Mol. Cell. Endocrinol. 83, 233 (1992). C. Clamagirand, M. Camier, C. Fahy, C. Clavreul, C. Creminon, and P. Cohen, Biochem. Biophys. Res. Commun. 143, 789 (1987). V. Greenen, F. Robert, H. Martens, A. Benhida, G. De Giovanni, M. P. Defresne, J. Boniver, J. J. Legros, J. Martial, and P. Franchimont, Mol. Cell. Endocrinol. 76, C27 (1991).

[11]

Approaches to Assessing Ontogeny of Processing Enzymes Richard G. Allen and Julianne Stack

Introduction

Posttranslational Processing of Preproopiomelanocortin Preproopiomelanocortin (POMC) is one of the most thoroughly characterized prohormones (12, 25, 30). The posttranslational processing of POMC is complex and varies in different cell types (13, 18). Each cryptic peptide encoded in POMC is flanked by two or more basic amino acid residues, for instance, -Lys-Arg-(KR), -RR-, -RK-,-KK-, a motif found in essentially all prohormones (21). Liberation of the bioactive peptides is a two-step process (29): the precursor is cleaved at the carboxyl side of the basic residues by a prohormone convertase (PC) (11, 17) and the remainder of basic residues exposed on the carboxyl-terminal end is removed by an enzyme with carboxypeptidase B activity (15). The biochemical basis for the tissue specificity of the proteolytic processing reactions is not completely understood. Several factors may be involved, including selective expression of the distinct PCs, differential compartmentation of either one or more proteases or the precursor, and modulation of cleavage-site accessibility by differential modification of the precursor (27).

Preproopiomelanocortin Processing: Cell and Tissue Specific Cell populations residing in the anterior and intermediate lobes of the pituitary gland process the common precursor prohormone POMC to different peptide end products (13, 19). In the rodent and monkey (1, 4, 16), anterior lobe corticotropes process POMC to predominantly/3-1ipotropin (/3-LPH), /3-endorphin(1-31), and adrenocorticotropin [ACTH(1-39)]; thus posttranslational processing stops at a certain proteolytic cleavage in the anterior lobe and does not proceed to the additional cleavages and biochemical modifications that define POMC-derived peptides in the melanotrope. Simply stated, the intermediate lobe (IL) POMC end products [a-melanocyte-stimulating hormone (a-MSH), N-acetylated and carboxy-shortened /3-endorphins, and ACTH(18-39) (CLIP)] are smaller (and further biochemi208

Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

[11] ONTOGENY OF PROCESSING ENZYMES

209

cally modified by a-N-acetylation, a-N,O-diacetylation, carboxy-terminal shortening, c~-amidation, and phosphorylation) when compared to the major POMC-derived end products ACTH(1-39) and/3-LPH produced in the anterior lobe (AL) corticotrope (13, 19). Thus,/3-endorphin(1-31) and ACTH(1-39) serve as biosynthetic intermediates requiring further endoproteolytic cleavages by PCs to reach their final forms. Many pulse-labeling schemes have been used to define the order of POMC-processing steps in the corticotrope and melanotrope (2, 14, 31). A strict order of cell-specific cleavages of precursor and intermediates has been assigned (31).

Preproopiomelanocortin Processing during Development Because the cells destined to secrete/3-endorphins, ACTH, and a-MSHs emanate from a common embryonic structure (Rathke's pouch), the POMC system continues to be an interesting model of cell differentiation and development that can be studied at the molecular level (1, 4, 16, 26). Over the last several years this laboratory has been studying POMC processing during fetal pituitary development, in both the monkey and rat (1, 4, 16). Here, we would like to present new data using reversed-phase high-performance liquid chromatography (RP-HPLC) fractionation methods combined with immunoassay of specific POMC-derived peptides, addressing the ontogeny of POMC processing, and discuss these findings in light of what is now known about the prohormone convertases and their endoproteolytic specificities. Methods

Tissue Procurement and Preparation Pituitary tissues obtained at different stages of prenatal [embryonic day 15 (el5)-birth] and postnatal (P1, P2 etc.) development are dissected with the aid of a dissecting microscope and homogenized in ice-cold 30% (v/v) acetic acid containing bovine serum albumin (BSA; 0.5 mg/ml) and phenylmethylsulfonyl fluoride (PMSF; 0.3 mg/ml). After freeze/thawing three times, the insoluble material is removed by centrifugation, an aliquot is taken for total POMC peptide immunoactivity, and the supernatants are diluted, lyophilized, and frozen at -80~ until fractionation by RP-HPLC.

Fractionation by RP-HPLC After lyophilization, samples are redissolved in 0.2-0.5 ml of buffer A and injected onto a Vydac RP-HPLC column (C4,300-/~ pore size; the Separa-

210

II

I M M U N O L O G I C A L AND B I O C H E M I C A L APPROACHES

m

~

~

~

~

~

~

m

|BIIIBBNNIIIIIINBIN ~n, A&~n'An'~Ik,~AJIk~IkAn

Z

n

-'0

0

~

10

~

20

~

~

30

40

~

50

~

n

60

70

80

"rime (min)

FIG. 1 Fractionation of peptides by RP-HPLC. Various combinations of 1-2 p,g of each peptide were dissolved in HPLC-grade H20 containing 0.1% TFA and injected onto a Vydac (Hesperia, CA) RP-HPLC column (C4, 300-,~ pore size). A Waters HPLC system with a fixed-wavelength UV detector (214 nm) was used to determine the elution positions of POMC-derived peptides. The heavy solid line shows the linear gradient of 0.1% TFA in HPLC-grade H20, and 80% CH3CN containing 0.1% TFA, that was used to elute peptides. The initial loading conditions were 17% solvent B with a brief "step up" to 23% solvent B. The flow rate was 1 ml/min. The marker peptides were as follows: (1) deacetyl-a-MSH, ACTH(1-13)NH2; (2) monoacetyl-a-MSH, a-N-acetyl-ACTH(1-13)NH2; (3) diacetyl-ct-MSH, a-N,O-diacetylACTH(1-13)NH2; (4) CLIP, human ACTH(18-39); (5) human ACTH(1-39); (6) fl-endorphin(1-31); (7) fl-endorphin(1-27); (8) a-N-acetyl-fl-endorphin(1-31); (9) fl-endorphin(1-26); (10) a-N-acetyl-fl-endorphin(1-27); (11) a-N-acetyl-fl-endorphin(1-26). All fl-endorphin peptides were the camel amino acid sequences, which correspond to rat fl-endorphins. tions Group, Hesperia, CA) and a Waters (Milford, NJ) HPLC system with a fixed-wavelength UV detector (214 nm) is used to fractionate the peptides. A linear gradient (shown in Fig. 1) of acetonitrile (CH3CN) in 0.1% trifluoroacetic acid (TFA) is used to elute peptides. The flow rate is 1 ml/min and 1-min fractions are collected. Buffer A is 0.1% TFA in HPLC-grade H20 (Baker, Phillipsburg, NJ); buffer B contains 80% CH3CN and 0.1% TFA. Synthetic peptides (1-2 g) are obtained from both Peninsula Laboratories (Belmont, CA) and Bachem (Torrance, CA) and used to determine the elution times of the POMC peptides shown in Fig. 1.

Radioimmunoassay of Preproopiomelanocortin Peptides All basic assay procedures have been described (1-3, 16). The 125I-labeled peptides used in the immunoassays are generated by either the hypochlorite

211

[11] ONTOGENY OF PROCESSING ENZYMES 1.2 A,

1.0 :E 0.8

.2

-~ 0.6

~ ~o., ~ "~o., o

~

0.0

I,LI 0,.

-. E

3

m

~

2

<

o

0

Z ~ X

~ =

4

1-ACTH(1-39) 2-? 3-? 4-POMC

2

0

20

40

TIME

60

80

(MIN)

FIG. 2 Further analysis of POMC processing patterns in developing rat pituitary tissues, as shown in Fig. 2. The fractions shown in Fig. 1 were assayed with ACTH midportion and ACTH C terminal-specific RIAs. (A) Whole pituitary (embryonic day 17); (B and C) anterior lobe (birth).

or the chloramine-T method. Synthetic peptides are obtained from Bachem and Peninsula. RP-HPLC fractions can be assayed directly up to 100 ~1, except for the C terminus-specific ACTH radioimmunoassay (RIA). In this RIA the samples require freeze drying before being diluted in assay buffer, owing to a small background interference by acetonitrile. Sensitivities of all POMC peptide RIAs are approximately 10-20 pg/tube. The recovery of all input immunoactivity is greater than 90%.

Preproopiomelanocortin Peptide Antiserum Specificity All POMC-specific antisera are generated in rabbits, using standard techniques (1). The /~-endorphin(1-31) antiserum used is midportion specific

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

and recognizes all molecules containing/3-endorphin, including POMC. The ACTH(1-39) antisera were midportion (Fig. 2A) and C-terminus (Fig. 2C) specific. Further, the ACTH midportion antiserum has an absolute requirement for an uncleaved KKRR sequence in the middle of ACTH(1-39). The /3-MSH antisera is C-terminus specific. The a-MSH antiserum is acetyl specific and demonstrates approximately 15% cross-reactivity with deacetyl-c~MSH and less than 0.01% cross-reactivity with ACTH(1-39) (1).

Primary Cell Culture Pituitaries are dissected on el9 as described in Allen et al. (4) and primary cultures are prepared as described previously (2). After 48 hr the cells are harvested in the extraction buffer described above, treated in the same way, and the extracts fractionated by RP-HPLC. Results

fl-Endorphin and ~-Melanocyte-Stimulating Hormone-Containing Peptides In the present study extracts of developing rat pituitary tissues taken at different stages of development were fractionated by RP-HPLC and the fractions assayed for POMC peptide immunoactivity. Here we demonstrate a striking example of processing pattern changes that take place in a 48- to 72-hr developmental time period. Figure 3 shows the processing patterns for/3-MSH (Fig. 3A) and/3-endorphine (Fig. 3B) found in whole pituitary tissue taken on e 17. Large amounts of unprocessed POMC were detected by both of the immunoassays. It should be noted that the difference in molar amounts between the /3-MSH and /3-endorphin RIAs is due to a species-specific amino acid sequence (rat vs monkey) in/3-melanotropins. Thus, at this stage of development/3-endorphin(1-31) was cleaved efficiently from/3-LPH; however, no appreciable amounts of carboxy-shortened /3-endorphins were detected. It has been thought that both of these cleavages are directed by PC2 (31). Four days later in development, different processing patterns begin to emerge. By birth, the anterior and neurointermediate (NIL) pituitary can be separated by dissection. Figure 3C and D shows that in developing anterior lobe corticotropes a large amount of POMC remains unprocessed as recognized by both RIAs. In marked contrast, as shown in Fig. 3E and F, NIL extracts contained small, relative amounts of unprocessed precursor, thereby indicating much more efficient cleavages of POMC in melanotropes. Further,

213

[11] ONTOGENY OF PROCESSING ENZYMES e17 WHOLE PiT

1.0

BIRTH; ANTERIOR LOBE C

0.8-

0.8-

0.8~. in

BIRTH; INTERMEDIATE LOBE

1.0

4

1

4

3

E

1

0.6-

0.4-

0.41

0.2-

3

0.2-

23

4

1

oO

0

......................................

o

20

4o

e0

oo

_!.!!.!!!'i!::!!!.! ..... .

0

20

40 TIME (MIN)

~

~

o

2o

4o

-~-

FIG. 3 Processing patterns of POMC-derived peptides in developing rat pituitary tissues as determined by RP-HPLC and RIA. Pituitary extracts were fractionated as described in Methods and fractions were assayed for/3-endorphin and/3-MSH immunoactivity. (A, C, and E)/3-MSH C-terminal immunoactivity; (B, D, and F) /3-Endorphin midportion immunoactivity. The elution positions of authentic synthetic and/or known final forms of these peptides found in adult pituitary tissues are shown.

at this stage, as in adult pituitary tissues,/~-LPH is completely processed to /3-endorphins in melanotropes while in corticotropes about 50% remains uncleaved.

Adrenocorticotropin-Containing Peptides Figure 4 shows the processing patterns for ACTH-containing POMC peptides at e 17 and birth in anterior lobe tissue, using a midportion-specific (Fig. 4A and B) and a C terminal-specific antiserum. The unidentified peaks eluting from 30 to 37 min have been identified as 13-kDa, N-glycosylated ACTH by HPLC studies performed by other laboratories (6). Further, this change in glycosylation state coincides with the apparent doublet, increasing in the region that POMC elutes (70-80 min, Fig. 4).

a-Melanocyte-Stimulating Hormone Peptides We then analyzed the processing patterns of a-MSHs in e 17 whole pituitary tissue, separated NILs at birth, 48-hr fetal cultures started on day 19 (equivalent to separated lobes isolated at birth) (Fig. 4), and adult NIL tissue (Fig.

214

_.,.A'[ ._:..,.,

II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES 0.4

9

,.

. . . . . . . . . . .

, . : . _

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ii1B , =

E

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1.cb,M,oo~

0.4

0.3

0.2

0.t

O0

0

20

40

60

..........................................

80

TIME (MIN)

FIG. 4 Further analysis of POMC-processing patterns in developing rat pituitary tissues, as shown in Fig. 2. The fractions shown in Fig. 2 were assayed for a-MSH immunoactivity, using an acetyl-specific RIA. This antiserum cross-reacts with deacetyl-a-MSH by approximately 15% and, as shown above, does not react with POMC at all. (A) Intermediate lobe (birth); (B) whole pituitary (embryonic day 17); (C) 48-hr fetal cultures.

5). Mature a-MSHs require N-terminal acetylation and C-terminal amidation. When compared to adult NIL tissue, all of the above tissue sources contain much larger relative amounts of deacetyl-a-MSH. Further, it is interesting to note that there is a small amount of deacetyl-MSH on e17 (Fig. 4B). PC2 is thought to participate in liberating a-MSHs from ACTH(1-39) and even though fl-endorphin(1-3 l) is efficiently cleaved from fl-LPH in el7 tissue, very small amounts of a-MSH-related material were deleted. Conversely, the predominant form of ct-MSH in adult NIL tissue is diacetyl-a-MSH, and

215

[11] ONTOGENY OF PROCESSING ENZYMES

c 0 (,,t 0

F:

60

W I-13. W 13. uJ

_~ I--

4o

Z

1 0

o

20

40

6O

80

TIME (MIN)

FIG. 5 Processing patterns of c~-MSHin adult NIL, as shown in Fig. 2.1, deacetyla-MSH; 2, mono actyl-c~-MSH; 3, diacetyl-a-MSH. this is demonstrated by the processing pattern shown in Fig. 5. Thus, as we and others have demonstrated, even though POMC-processing patterns are well established at birth, they continue to change throughout early postnatal development (26). This is apparent when comparing the acetylation state of a-MSHs and/3-endorphins.

Discussion In Fig. 6 we have labeled the paired basic endoproteolytic cleavage sites from the amino to carboxyl terminus of POMC as a-g for the purpose of discussion. Many laboratories have used a variety of approaches to elucidate the basis of cell- and tissue-specific POMC processing and at present there

216

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES PC2

PC 1

Iy3MSHI

PC 1/PC 2

JP

I

~

PC2

I CLIPACTH

PC 1

I

PC 2

13LPH

I

i a

PC 2

I~ENDO

I

II b

c

d

9

f

g

FIG. 6 The POMC precursor with its known prohormone convertase cleavage sites, depicting the biologically active peptide domains as described in Thomas et al. (24, 28). The cleavage sites are designated a-g, from N terminus to C terminus.

is some agreement regarding which PC is responsible for which cleavage (Fig. 6 caption). However, as we have shown, the expression of the different PCs during pituitary development cannot completely account for the processing patterns found at those stages. It is interesting to relate our findings shown here to the ontogeny studies addressing the expression of the mRNAs encoding PC1 and PC2 during development, carried out in both the mouse (20) and rat (see [3] in this volume). In adult pituitary tissues PC2 has been postulated to account for the major difference between corticotrope and melanotrope endoproteolytic processing patterns (31). Studies of adult pituitary show that PC2 is predominantly expressed in the IL, while PC1 is predominantly expressed in the AL. In e17 pituitary POMC mRNA and protein are expressed almost exclusively in anterior lobe locations (23). Confoundingly, in both the rat and mouse, the predominant PC expressed at this stage is PC2, which is predominantly expressed in the IL of adult pituitary tissues. As just one example, the data shown here demonstrate that there is efficient cleavage at site f, but not at sites g and d, which are extremely efficient cleavages in adult IL (24, 28, 31). There are several others, including our studies of the ontogeny of pituitary POMC-processing patterns in the monkey, showing that by midgestation (e80) the ACTH pathway is completely established while the IL melanotropin pathway matures much later (1). Here, then, is another example of PC2 cleavage expression exhibiting a developmental and temporal aspect. These observations taken together suggest that factors other than the PC specificities may account for cell-specific POMC processing. Further, the fact that ACTH(1-39) is not efficiently processed to melanotropins in corticotropes, even when PC2 is the only convertase expressed in the developing rodent pituitary, also supports this notion. Immunocytochemical studies were the vanguard of experiments directed at discovering the appearance of certain proteins during pituitary development. Numerous immunocytochemical demonstrations of POMC-related peptides in fetal, avian, and mammalian pituitaries have been reported (5, 7-10, 22). These studies could not resolve the extent of POMC processing in the

[11] ONTOGENY OF PROCESSING ENZYMES

217

developing corticotrope/melanotrope. Also, because the antisera employed in these studies might react with POMC, as well as the multiple forms of the immunodeterminants, these studies were difficult to interpret from the viewpoint of understanding POMC processing during development. We have made significant progress in biochemically characterizing POMC peptide expression during development (1, 16, 26) and have demonstrated that the combination of R P - H P L C and specific immunoassay is a powerful method with which to address posttranslational processing of neuroendocrine precursor prohormones.

References

,

5. 6. 7. 8.

9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.

R. G. Allen, J. M. Hatfield, and J. Stack, Dev. Biol. 126, 156 (1988). R. G. Allen, E. Herbert, M. Hinman, H. Shibuya, and C. B. Pert, Proc. Natl. Acad. Sci. U.S.A. 75, 4972 (1978). R. G. Allen, E. Orwoll, J. W. Kendall, E. Herbert, and H. Paxton, J. Clin. Endocrinol. Metab. 51, 376 (1980). R. G. Allen, J. E. Pintar, J. Stack, and J. W. Kendall, Dev. Biol. 102, 43 (1984). M. Begeot, M. P. Dubois, and P. M. Dubois, Cell Tissue Res. 193, 413 (1978). H. P. J. Bennett, C. A. Browne, and S. Solomon, J. Biol. Chem. 257, 10096(1982). A. Chatel.ain, M. P. Dubois, and J. P. Dupuoy, Cell Tissue Res. 169, 335 (1976). A. Chatelain, J. P. Dupouy, and M. P. Dubois, Cell Tissue Res. 196, 409 (1979). A. Chatelain and J. P. Dupuoy, Neuroendocrinology 33, 148 (1981). H. Coffigny and H. P. Dupuoy, Gen. Comp. Endocrinol. 34, 312 (1978). J. Douglass, O. Civelli, and E. Herbert, Annu. Rev. Biochem. 53, 665 (1984). B. A. Eipper and R. E. Mains, Endocr. Rev. 1, 1 (1980). B. A. Eipper and R. E. Mains, J. Biol. Chem. 256, 5689 (1981). B. A. Eipper, D. A. Stoffers, and R. E. Mains,Annu. Rev. Neurosci. 15, 57 (1993). L. D. Fricker, B. Das, R. S. Klein, D. Greene, and Y. K. Jung, NIDA Res. Monogr. 111, 171 (1991). J. M. Hatfield, R. G. Allen, J. Stack, and O. Ronnekleiv, Dev. Biol. 126, 164 (1988). D. T. Krieger, Science 222, 975 (1983). Y. P. Loh and H. Gainer, "Brain Peptides," p. 76. Wiley (Interscience), New York, 1983. R. E. Mains and B. A. Eipper, J. Biol. Chem. 256, 5683 (1981). M. Marcinkiewicz, R. Day, N. G. Seidah, M. Chr6tien, Pror Natl. Acad. Sci. U.S.A. 90, 4922 (1993). T. L. O'Donohue and D. M. Dorsa, Peptides (N. Y.) 3, 353 (1982). R. Y. Osamura and J. K. Nakane, Acta Histochem. Cytochem. 15, 294 (1982). L. E. Pintar and D. I. Lugo, Ann. N.Y. Acad. Sci. 512, 218 (1987). C. J. Rhodes, B. A. Thorne, B. Lincoln, E. Nielsen, J. C. Hutton, and G. Thomas, J. Biol. Chem. 268, 4267 (1993).

218

II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES 25. J. L. Roberts, M. Phillips, P. A. Rosa, and E. Herbert, Biochemistry 17, 3609 (1978). 26. S. M. Sato and R. E. Mains, Endocrinology (Baltimore) 117, 773 (1985). 27. G. Thomas, B. A. Thorne, L. Thomas, R. G. Allen, D. E. Hruby, R. Fuller, and J. Thorner, Science 241, 226 (1988). 28. L. Thomas, R. Leduc, B. A. Thorne, S. P. Smeekens, D. F. Steiner, and G. Thomas, Proc. Natl. Acad. Sci. U.S.A. 88, 5297 (1991). 29. M. O. Thorner, Clin. Endocrinol. Metab. 6, 201 (1977). 30. M. Uhler and E. Herbert, J. Biol. Chem. 258, 257 (1982). 31. A. Zhou, B. T. Bloomquist, and R. E. Mains, J. Biol. Chem. 268, 1763 (1993).

[12]

Measurement, Distribution, and Subcellular Localization of Peptide-Amidating Activity Rebecca A. Lew and A. Ian Smith

Introduction Peptide hormones are derived from larger precursor molecules that are processed by specific enzymes to produce mature bioactive peptides. These enzymes include endopeptidases, which cleave at paired or single basic amino acids; exopeptidases, which remove specific residues from either the amino or carboxyl terminus; and other modifying enzymes, such as Nacetyltransferase, glycosyltransferases, and phosphorylases. One common modification is a-amidation of the carboxyl terminus, which is mandatory for the bioactivity of a host of peptides. These include the hypothalamic releasing factors (corticotropin-releasing factor, gonadotropin-releasing hormone, growth hormone-releasing hormone, and thyrotropin-releasing hormone), neurohypophyseal peptides (vasopressin and oxytocin), other neuropeptides (neuropeptide Y, calcitonin gene-related peptide, and substance P), and gastrointestinal peptides (cholecystokinin, gastrin, vasoactive intestinal peptide, and bombesin). The a-amide is derived from a C-terminal glycine residue present in the immediate precursor (1); conversion is initiated by hydroxylation of the glycine, followed by formation of the peptidylamide and glyoxylic acid from the a-hydroxyglycine intermediate (2, 3). Although originally considered to be catalyzed by a single enzyme, known as peptidylglycine a-amidating monooxygenase (PAM) (4), studies indicate that separate enzymatic domains expressed within the PAM precursor catalyze the two reaction steps, although formation of the amide from the a-hydroxyglycine intermediate can occur spontaneously at neutral to alkaline pH (5-7). Most methods used to identify sites of peptide synthesis (e.g., immunohistochemistry, in situ hybridization) require prior knowledge of the peptide and/or its mRNA structure. However, tissues that are potential sources of novel peptides may be identified by screening for enzymes involved in their synthesis. For example, one could examine expression of prohormone convertase mRNA (8, 9) (see [1] in this volume). Alternatively, direct assays of enzyme activity could be used to screen a variety of tissues rapidly. Although assays of prohormone convertase activity are hindered by the complex specificity of these enzymes, assays for amidating activity are relatively simple to perform, and can provide valuable information regarding the Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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tissue distribution of amidated peptides, as well as their possible regulation. Once a potential site of amidated peptide synthesis is identified, the peptide(s) themselves may be isolated by techniques that exploit the presence of the C-terminal amide (10).

General Features and Considerations for Measurement of a-Amidating Activity Amidating activity can be readily monitored using a radiolabeled synthetic tripeptide substrate, usually [~25I]Ac-Tyr-Val-Gly. Of course, the C-terminal glycine is necessary as the nitrogen donor for amidation, while the tripeptide is the minimum length required by the enzyme (11). The tyrosine provides a site for iodination, and is N-acetylated to increase the stability of the peptide. The D isomer of tyrosine has also been frequently used, although we find that the acetylated form is more readily extracted into the organic phase. The second residue of the substrate can be varied; structure-activity studies indicate that the amino acids that are most readily amidated are sulfurcontaining, aromatic, and nonpolar residues, while more polar or charged residues, as well as glycine, react more slowly (1 1). Despite this preference, there are a few examples of naturally occurring peptides terminating in amidated charged residues (e.g., joining peptide, derived from proopiomelanocortin, ends with a glutamate amide), and a greater number of peptides that are amidated at a glycine residue (vasopressin, oxytocin, gonadotropinreleasing hormone, and pancreastatin). The a-amidation assay is based on the incubation of [~25I]Ac-Tyr-Val-Gly with tissue samples containing amidating enzyme, resulting in conversion of the peptide to [125I]Ac-Tyr-Val-NH2. The product can be separated from the substrate by selective extraction into alkaline ethyl acetate (facilitated by the absence of charge at high pH), and following separation the radioactivity in both the organic and aqueous phases can be determined (this separation is illustrated in Fig. 1). This rapid separation of substrate and product compares favorably with alternate methods, most notably ion-exchange chromatography, which was used extensively in early studies of the PAM enzyme (1, 12). Although chromatographic separation of substrate from product is straightforward, it is too time consuming for the screening of multiple samples, especially if performed in duplicate. The advent of a rapid separation technique, first described by Mizuno et al. (13, 14) and later adapted by our laboratory (15, 16) and others (17), facilitated comparisons of amidating activity between many different tissues or subcellular fractions of tissues within the same assay, which typically takes only 1 day to perform. The following features, however, should be carefully considered.

[12] PEPTIDE-AMIDATINGACTIVITY

221 Add 1 N NaOH + Ethyl Acetate

Incubation

i~::.:

:..~:..

~- Organicphase

v

Vortex and Centrifuge *:~**~

Aqueousphase

Substrate + Tissue Sample

o Substrate= iodo[acetyl-125I]Try-Val-Gly 9 Product= iodo[acetyl-125I]Try-Val-NH 2

Separate phases and count radioactivity FIG. 1 Schematic representation of amidation assay. Incubation of radiolabeled substrate (e.g., [125I]Ac-Tyr-Val-Gly) with sample results in partial conversion to amidated product. Separation of product from substrate is achieved by a selective extraction of the former into alkaline ethyl acetate. Conversion is quantitated by measurement of the radioactivity present in the aqueous (substrate) and organic (product) phases.

1. Tissue preparation: Tissues are normally homogenized, although PAM activity can also be measured in plasma, cerebrospinal fluid, cell culture medium, and so on. Localization of the enzyme to membrane or soluble compartments can be determined using subcellular fractions, as demonstrated below. When homogenization is necessary, it is advisable to include general protease inhibitors to prevent destruction of PAM by lysosomal enzymes. However, chelators of divalent cations (EDTA, EGTA, and phenanthroline) should be avoided, because these will also inhibit the amidating

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enzyme, which is dependent on copper ions. Once homogenized, samples should be aliquoted and stored at -70~ (storage at 4~ or repeated freezing and thawing, leads to a rapid decrease in activity). 2. Cofactors: Amidation of peptides by PAM requires the presence of three cofactors; molecular oxygen, reduced ascorbate, and copper ions. The requirement for oxygen reflects the monooxygenase nature of the enzyme, and can be met simply by using normal incubation conditions. The optimal concentrations of ascorbate and copper should be determined for each system, as described below; generally, ascorbate concentrations in the low millimolar range, and copper (normally CuSO4) concentrations in the low micromolar range, are sufficient. The enzyme catalase is also added to restore oxidized ascorbate to its reduced form. 3. Substrate: The nature of the synthetic substrate has already been discussed. In most instances, only a trace amount (<25,000 cpm) of labeled substrate is used in the assay. However, if kinetic studies are performed, known quantities of unlabeled substrate are also added to the incubation mixture. Although only amidation of the labeled substrate is actually measured in the assay, the extent of conversion can be extrapolated to the cold substrate, because the presence of an iodine does not alter enzyme affinity for the substrate (12). 4. Incubation time: Optimal incubation time should be determined by performing a preliminary time course experiment. Ideally, somewhere between 5 and 25% of the substrate should be amidated at the end of incubation. We routinely use an incubation period of 4 hr, which allows the entire assay to be completed in 8 hr. Shorter incubation times can be used, but the quantity of sample must be increased, such that the length of incubation might be dictated by sample size limitations. 5. Separation of product and substrate: Although simple to perform, this step of the assay is the most time consuming. Prior to the actual separation, ethyl acetate must be mixed thoroughly with an approximately equal volume of 1 M NaOH. This saturates ethyl acetate with the aqueous phase, and reduces the transfer of nonreacted substrate into the organic phase. The two phases separate within 5-10 min, and the appropriate solvent can be drawn from either the top (organic) or bottom (aqueous) with a glass pipette. After incubation, 3 vol of 1 M NaOH is added to the sample, followed by 8 vol of ethyl acetate, giving twice the volume of organic to aqueous solvent. The samples are then vortexed thoroughly, and the phases separated by a lowspeed centrifugation. Phase separation will occur if the samples are simply left on the bench, but centrifugation prevents the formation of pockets of one solvent within the other. The upper organic phase is then carefully removed from the aqueous layer, using either a glass Pasteur pipette or an automatic, variable-volume pipettor with a plastic tip. The latter method

[12] PEPTIDE-AMIDATING ACTIVITY

223

usually allows greater control, and setting the volume to be withdrawn helps prevent accidental removal of any aqueous phase. This is a crucial step in the assay and care must be taken to avoid removing any of the aqueous phase with ethyl acetate. Given the larger volume and the lower total radioactivity present in the organic phase, it is far better to leave a small amount with the aqueous phase than to risk contaminating the ethyl acetate with even a tiny volume of the aqueous fraction. It is also advisable to run a set of controls containing tracer minus tissue to account for any transfer of nonamidated substrate into the organic phase (normally 1-2% total). In the following sections of this chapter, we briefly describe two specific applications of the amidation assay to examine the distribution of PAM in the heart, pituitary, and central nervous system of the sheep.

S u b c e l l u l a r D i s t r i b u t i o n o f P e p t i d y l g l y c i n e c~-Amidating M o n o o x y g e n a s e in S h e e p H e a r t Our interest in amidating activity in the heart was prompted by the finding by Braas et al. (18) that the cardiac atrium of the rat contains by far the highest concentration of PAM activity of any tissue tested, including the hypothalamus and anterior pituitary. This was surprising, because the only peptides known to be produced in the atria are those of the atrial natriuretic peptide (ANP) family, none of which are amidated. In addition, the ANP precursor does not contain any sequences indicative of a possible amidated peptide (i.e., a dibasic cleavage site preceded by a glycine residue). Thus we (and others) felt it was likely that the atria produce another, amidated peptide unrelated to ANP. As a first step toward isolating such a peptide, we wanted to verify the existence of PAM in the atria of sheep, which given the availability of slaughterhouse material, may prove more useful than the rat in the purification of an atrial amidated peptide. The methods detailed below and published elsewhere (16) may prove particularly useful as a guideline for optimizing the amidation assay in any given system.

Methods Tissue Samples Ovine atrial and ventricular tissue are derived from 6 sheep slaughtered at a local abattoir, while the rat tissues are obtained from 35 male SpragueDawley rats. Left atria, right atria, and ventricular apices from each species

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are pooled and processed in parallel. All tissues are immediately frozen in liquid nitrogen following dissection, and are kept at -70~ until homogenization.

Tissue Homogenization Tissue samples are weighed and homogenized at 4~ [Polytron (Brinkmann, Westbury, NY) homogenizer] in 20 mM N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid (TES) buffer, pH 7.4, containing 10 mM mannitol, phenylmethylsulfonyl fluoride (300/xg/ml), leupeptin (2/zg/ml), and benzamidine (16/xg/ml) (19). Homogenates are frozen and thawed three times to rupture cellular membranes, then centrifuged at low speed (4000 rpm) for 5 min at 4~ in a tabletop microfuge to sediment debris. The homogenates are then centrifuged at 100,000 g for 60 min at 4~ the supernatants collected, and the pellets resuspended in fresh homogenization buffer. The supernatants from a second ultracentrifugation (100,000 g, 60 min, 4~ are pooled with the first to constitute the soluble fraction of each homogenate. The remaining pellets are resuspended in homogenization buffer containing 1% (v/v) Triton X-100 and shaken at 4~ for 60 min to solubilize membrane-associated protein. Supernatants from a third ultracentrifugation (100,000 g, 60 min, 4~ of this material are designated as membrane-associated fractions. Aliquots of both soluble and membrane-associated fractions from each sample are immediately frozen at -70~ until assay for amidating activity. An aliquot from each sample is also measured for protein content by the Lowry method (20). Amidation Assays Homogenate is incubated in a final volume of 100/zl of 0.2 M Tris-HC1 buffer at 37~ with radiolabeled synthetic substrate 125I-labeled o-Tyr-Val-Gly (o[125I]Tyr-Val-Gly) or [125I]Ac-Tyr-Val-Gly in the presence of required cofactors (CuSO4, ascorbate, and catalase) with or without addition of unlabeled substrate. The concentrations of these components in each experiment are detailed in Results (below). Following incubation, amidated product is separated from substrate by selective extraction into alkaline ethyl acetate. To each sample, 300/xl of 1 M NaOH and 800/xl of ethyl acetate are added, the sample is vortexed for 30 sec, and then centrifuged at 4000 rpm for 5 min in a tabletop microfuge. The organic phase is then carefully separated from the aqueous phase and the radioactivity in both phases counted in a y counter (Packard, Meriden, CT). Results from duplicate.samples are averaged (individual results differ from the mean by < 10%), and the transfer of iodinated substrate into the organic phase in the absence of tissue homogenate (<5%) is subtracted to give a net percent conversion. In experiments in

[12]

PEPTIDE-AMIDATING

225

ACTIVITY

10 .~ 10 o=

8

~o=

6

8 rJ 6

rj

~,

0

0

J

lb

1~

210

215

310

Incubation Time (hr)

315

4

ir

5.5

~

6~5

~ pH

7.15

~

8~5

FIG. 2 (A) Amidation of substrate by sheep atrial membranes is linear with incubation time. Tissue protein (1 /zg) was incubated with D-[125I]Tyr-Val-Gly (100,000 cpm ~ 6.6 nM) in the presence of cofactors [5/zM CUSO4-0.5 mM ascorbate-catalase (22,000 U/ml) (11) or 2/~M Cu804-2 mM ascorbate-catalase (22,000 U/ml) (O)] in 0.2 M Tris-HC1 buffer, pH 7.0, for 0.5, 1, 2, 4, 8, 16, 24, or 32 hr. (B) Effect of pH on amidation by sheep atrial membranes. Tissue protein (1/zg) was incubated with o-[125I]Tyr-Val-Gly (100,000 cpm ~ 6.6 nM) for 16 hr in the presence of 3 /zM CuSO4-0.1 mM ascorbate-catalase (22,000 U/ml) at the pH indicated. Because conversion was optimal at pH 7.5, subsequent experiments were performed at this pH. which only tracer is included, the data are expressed as percent conversion, which at such low substrate concentrations would be proportional to rate. When unlabeled substrate is added, as in experiments examining enzyme kinetics, then the data are expressed as picomoles of substrate converted per microgram of protein per hour.

Materials D-Tyr-Val-Gly and Ac-Tyr-Val-Gly are purchased from Peninsula (Belmont, Ca) and iodinated by the Iodogen method (Pierce, Rockford, IL) (21). Catalase and leupeptin are obtained from Boehringer Mannheim (Melbourne, Australia). All other chemicals are obtained from either Sigma (Melbourne, Australia) or Ajax Chemical (Melbourne, Australia).

Results Optimization of Assay Conditions Incubation Time The conversion of substrate to product was shown to increase linearly with time (Fig. 2A) by incubating sheep atrial membrane-associated protein (1 ~g) with 125I-labeled D-Tyr-Val-Gly (100,000 cpm ~ 6.6 nM) in the presence

226

II

IMMUNOLOGICAL

AND

BIOCHEMICAL

APPROACHES

A

B

8-

4-

0.1 mM 30 ~tM 6

7-.

2

3

3 ~tM

1

0 0

I 0.2

I i 0.4 0.6 I/S (I.tM)-l

i 0.8

i 1

0 0

~ 0.2

~

, ~ ~ 0.4 0.6 I/S (l.tM)-1

0.8

I 1

FIG. 3 Effect of (A) ascorbate and (B) copper concentration on amidating enzyme kinetics. Sheep atrial membrane-associated protein (2/zg) was incubated with []25I]AcTyr-Val-Gly (100,000 cpm = 6.6 nM) and unlabeled substrate (1-20/zM) for 4 hr. Cofactors were added at the following concentrations: (A) 3/zM CuSO4, catalase (22,000 U/ml) and 0.1, 1.0, or 10 mM ascorbate; (B) 1 mM ascorbate, catalase (22,000 U/ml), and 0.3, 3.0, or 30/zM CuSO4. Reaction velocity was determined on the basis of the percent conversion of tracer and the concentration of substrate present.

of cofactors [5/zM

Cu804-0.5

mM ascorbate-catalase (22,000 U/ml) or 2

/zM Cu504-2 mM ascorbate-catalase (22,000 U/ml)] in 0.2 M Tris-HC1 buffer, pH 7.0, for 0.5, 1, 2, 4, 8, 16, 24, or 32 hr. Although conversion of substrate to product was low in this early experiment, manipulation of the cofactor concentrations and the pH (detailed below), as well as using the acetylated rather than the D-Tyr substrate, resulted in improved rates of conversion in subsequent experiments. pH Optimum When the effects of buffer pH were tested over the range of pH 6.0-8.0 [sheep atrial membrane-associated protein (1/~g) incubated at 37~ with D[~25I]Tyr-Val-Gly (100,000 cpm ~ 6.6 nM) in the presence of 3 ~M CuSO4-0.1 mM ascorbate-catalase (22,000 U/ml) in 0.2 M Tris-HCl buffer at pH 6.0, 6.5, 7.0, 7.5, or 8.0 for 16 hr], conversion was found to be optimal at pH 7.5 (Fig. 2B); this pH was used for all subsequent experiments. Ascorbate Optimum The optimal concentration of ascorbate was determined by kinetic analysis of samples incubated in the presence of 0.1, 1, or 10 mM ascorbate at 3 /~M CuSO4 (Fig. 3A). Sheep atrial membrane-associated protein (2/~g) was incubated at 37~ with [125I]Ac-Tyr-Val-Gly (100,000 cpm ~ 6.6 nM) and

[12]

P E P T I D E - A M I D A T I N G ACTIVITY

227

unlabeled substrate (1-20 /xM) in the presence of 3 ~M CuSO4, catalase (22,000 U/ml), and 0.1, 1.0, or 10 mM ascorbate in 0.2 M Tris-HC1 buffer at pH 7.5 for 4 hr. The K m and Vmaxvalues observed were 4.1/zM and 0.7 pmol/p~g/hr, respectively, at 0.1 mM, 12.8/zM and 4.5 pmol//xg/hr at ! mM, and 55.6 /xM and 12.5 pmol//xg/hr at 10 mM. Thus, increasing ascorbate concentration from 0.1 to l0 mM enhances the velocity of the amidation reaction, yet also lowers the apparent affinity of the enzyme for its substrate. An intermediate ascorbate concentration of 1 mM was used in subsequent experiments. Copper Ion Optimum Similar kinetic analysis was used to determine the optimal copper concentration. When sheep atrial membrane-associated protein (2/xg) was incubated at 37~ with [125I]Ac-Tyr-Val-Gly (100,000 cpm ~ 6.6 nM) and unlabeled substrate (1-20/xM) in the presence of 1 mM ascorbate, catalase (22,000 U/ml) and 0.3/xM CuSO4 in 0.2 M Tris-HC1 buffer at pH 7.5 for 4 hr, the K m and Vmax values were indeterminable (data not shown). However, at 3 or 30/~M CuSO4, the K m and Vma• values observed were 27.0/zM and 8.8 pmol//xg/hr, and 10.8/~M and 3.6 pmol/~g/hr, respectively (Fig. 3B). These results indicate that the amidating activity in our atrial preparations requires a minimal copper concentration greater than 0.3/~M, but increasing CuSO4 concentration above 3/xM only marginally increases enzyme-substrate affinity and reduces reaction velocity. Subsequent experiments were performed using 3/~M CuSO4.

Distribution of Peptidylglycine ~-Amidating Monooxygenase Activity in Sheep and Rat Heart Samples from both soluble and membrane fractions of left atria, right atria, and ventricle from both species were compared. Each sample (0.5-8/zg of protein) was incubated with [~25I]Ac-Tyr-Val-Gly (100,000 cpm ~6.6 nM) for 4 hr at pH 7.5 in the presence of 3/xM CuSO4, 1 mM ascorbate, and catalase (22,000 U/ml). Membrane samples were diluted at least 20-fold in the assay, such that the final concentration of Triton X-100 (less than 0.05%, v/v) would not inhibit enzyme activity (22). Kinetic studies of the right and left atrial membrane-associated amidation activity of both species were performed by incubating 2/xg of sample protein under the same conditions with the addition of 1-100/~M unlabeled substrate. In both the sheep and the rat, the highest concentration of amidating activity was measured in the membrane-associated fraction of the atria (Fig. 4). The left and right atrial membrane fractions contained similar amounts of activity in both species, although the levels in the left atrium tended to

228

II

IMMUNOLOGICAL

AND

BIOCHEMICAL

APPROACHES

A

B 60 -

60-

50-

50-

40-

Membrane Soluble Left atria Right atria Ventricle

=o 4 0 -

O

F, o

30 = Dr

Left atri a

9

Rightatria

20-

9

Ventricle

9

[]

9

0

10-

[] & 0

30-

ble

O o

9 A 9

O

~" 2 0 -

10.

0

o.. 0

r 2

i 4 Protein (lxg)

, 6

r 8

0 0

2

4

g

8

Protein (Jig)

FIG. 4 PAM activity in the soluble and membrane-associated fractions of left and right atria and ventricle of the rat (A) and sheep (B). Sample protein (0.5-8/xg) was incubated for 4 hr with [125I]Ac-Tyr-Val-Gly(100,000 cpm ~ 6.6 nM) in the presence of 3/xM CuSO4-1 mM ascorbate-catalase (22,000 U/ml). be higher. The soluble fractions of the rat atria also contained measurable amidating activity (5- to 10-fold lower than the membrane fraction at 8 ~g of protein), while the levels of enzyme in the sheep atrial soluble fractions were minimal. Low levels of activity were also observed in the membrane and soluble fractions of the ventricle in both species. Enzyme activity was approximately linear with respect to protein concentration up to 4 tzg (Fig. 4). The plateau of activity above this concentration may result from rising concentrations of inhibitors of PAM (e.g., copper-binding proteins) found in the crude preparations; such endogenous inhibitory activity has been reported in rat anterior pituitary (4). Kinetic analyses of the membrane-associated fraction of both atria in both species were performed to examine possible tissue or species differences in enzyme affinity and reaction velocity. In the rat, the apparent Km and Vmax values for the left and right atria were 17.4/xM and 43.9 pmol//xg/hr, and 15.6/xM and 40.5 pmol//zg/hr, respectively (Fig. 5A). In the sheep, these values were 15.6/xM and 15.1 pmol//~g/hr for the left atrium, and 16.7/xM and 12.8 pmol//xg/hr for the right atrium (Fig. 5B). Thus, while the K m values were similar between atria and between species, the Vmaxvalues were approximately threefold higher in the rat than in the sheep.

Effect of Chronic Corticosteroid Treatment on Sheep Heart Amidation Activity Because tissue levels of several peptide-processing enzymes have been shown to be altered by corticosteroids (23), we also examined the potential

[12]

PEPTIDE-AMIDATING

229

ACTIVITY

A 1.4 -

0.5 u

0.4 =

9 Right atria KM= 15.6 BM

|

1.O-

Vmax = 40.5

__ p m o l / ~ g / h r /

0.3-

1.2 =

9 Right atria KM= 16.7 BM Vmax= 12.8 __ _ _ P m ~

~ /"

/

:t 0.80.6..

0.2-

/

0.1- f

-

KM= 17ABM Vmax = 4 3 . 9 pmol/gg/hr

J

/ ' /

-~ 0.40.2-

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FIG. 5 Kinetics of left and right atrial membrane-associated PAM activity in the rat (A) and the sheep (B). Sample protein (2/xg) was incubated with [125I]Ac-TyrVal-Gly(100,000 cpm ~ 6.6 nM) and unlabeled substrate (1-100/zM) in the presence of 3/zM CuSO4-1 mM ascorbate-catalase (22,000 U/ml) for 4 hr. regulation of cardiac PAM activity in sheep by chronic glucocorticoid administration. Four ovariectomized Corriedale ewes were given 1 mg of dexamethasone in 1 ml of saline intramuscularly twice a day for 10 days, while four other ovariectomized ewes receiving saline served as controls. The left and right atria and ventricular apices from these animals were processed individually and in parallel. Blood was also collected at the time of sacrifice, and plasma was assayed for cortisol by radioimmunoassay. Both soluble and membrane-associated fractions of the left atrium, right atrium, and ventricular apex of each sheep were tested for amidation activity by incubating 10 /xg of sample protein with [~25I]Ac-Tyr-Val-Gly (20,000 cpm ~ 1.3 nM) for 4 hr at pH 7.5 in the presence of 3 txM CuSO4, 1 mM ascorbate, and catalase (22,000 U/ml). Kinetic studies of the amidation activity in the membraneassociated fraction of atrial tissue were performed by incubating 2 txg of sample protein under the same conditions, with the addition of 1-20 txM unlabeled substrate. As in the previous experiments, the highest level of amidating activity was found in the membrane-associated fractions of the atria; however, treatment with dexamethasone did not significantly affect the levels measured in any tissue fraction (data not shown). In addition, there was no effect of dexamethasone on the apparent Km and gmax values for the atrial membrane fractions, and the values were similar to those found using pooled abattoir sheep material (K m values of 16.1-19.9 txM, gmax values of 10.7-29.5 pmol/txg/hr). Plasma cortisol levels at slaughter in the dexamethasone-treated sheep were all below the level of detection (< 10 nmol/liter), while cortisol concentrations in the control sheep ranged from 31 to 177 nmol/liter.

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Conclusions The study of atrial amidating activity described above illustrates several uses of the amidation assay. First, one should optimize the incubation time, pH, and cofactor concentrations for the system being studied. Once optimized, the assay can be used to compare levels of amidating activity between tissues, between subcellular fractions of a single tissue, and between species. One can compare not only absolute levels of activity, but also specific activities and kinetic parameters. In addition, although no significant effects of dexamethasone treatment on PAM activity were seen in this experiment, the amidation assay can be used to examine the possible regulation of PAM activity.

Distribution of Peptidylglycine c~-Amidating Monooxygenase Activity in Sheep Brain The majority of amidating activity in rat anterior and neurointermediate pituitary has been localized to secretory granules isolated by density gradient centrifugation (22), and amidation is therefore considered to occur within mature secretory vesicles. Significant activity (12-18% of the total), however, was also found in the rough endoplasmic reticulum and Golgi fractions (22), suggesting that amidation could potentially occur much earlier in the secretory pathway. In this study (15), we examined the distribution of PAM activity in the sheep brain, with particular attention to the hypothalamohypophyseal system, in which the site of secretory granule formation (the hypothalamic cell bodies) is distinct from the sites of vesicle storage and peptide release (the median eminence and neurohypophysis). In addition, we have examined the possible corelease of PAM from hypothalamic nerve terminals by measuring amidating activity in hypophyseal-portal blood before and after hypoglycemic stress.

Methods Tissue Homogenization Eleven brain regions are dissected from 4 age- and sex-matched sheep at a local abattoir and immediately frozen in liquid nitrogen. The brain regions collected are anterior pituitary (AP), neurointermediate lobe of the pituitary (NIL), median eminence (ME), medial basal hypothalamus (MBH), preoptic anterior hypothalamus (POA), hippocampus (HIP), pineal (PIN), cerebral

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cortex (COR), cerebellum (CER), pons medulla (PONS), and brainstem (BS). Individual whole tissues (AP, NIL, ME, MBH, POA, PIN, and HIP) or tissue samples (COR, CER, PONS, and BS) are then weighed and homogenized at 4~ as described above. Membrane and soluble fractions of each tissue are prepared by ultracentrifugation as before, and aliquoted and frozen at -70~ until assayed.

Hypophyseal-Portal and Jugular Blood Collection Four mature Corriedale ewes are fitted with hypophyseal-portal and jugular cannulas 1 day prior to experimentation, according to the method of Clarke and Cummins (24). On the day of experimentation, the sheep are heparinized, and both portal (1-2 ml) and jugular (10 ml) blood samples are taken every 10 min. Following a 20- to 30-min basal sampling period, the animals are injected with insulin (100 U) to induce hypoglycemic stress. Ninety minutes later, glucose (25 ml of a 50% solution) is given to restore blood sugar levels. Glucose levels are determined in each jugular blood sample, and portal and jugular plasma frozen at -70~ Assays for amidating activity and protein content are performed on alternate samples. Where sample volume permits jugular plasma samples are also measured for cortisol by radioimmunoassay.

Amidation Assays Homogenate (0.5-10/xg of protein) or plasma [2.86/zl (~200/zg of protein)] is incubated at 37~ with radiolabeled synthetic substrate [~25I]Ac-Tyr-ValGly (20,000-25,000 cpm ~ 16 pmol) in the presence of cofactors [3 /xM CuSO4, 1 mM ascorbate, and catalase (22,000 U/ml)] in 0.2 M Tris-HC1 buffer, pH 7.6, for 4 hr (final volume, 100/xl). For kinetic studies, unlabeled substrate (0.3-16.7/xM) is added. Following incubation, amidated product is separated from substrate by selective extraction into alkaline ethyl acetate, as described above. The mean net percent conversion (_+SE) of the four sheep is calculated for each tissue, except for the plasma samples; for these the data for each sheep are examined individually.

Results Tissue Distribution of Amidating Activity Measurable amidating activity was present in nearly all brain regions assayed, but by far the greatest concentrations were found in the medial basal and preoptic anterior areas of the hypothalamus, where conversion of substrate

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AND BIOCHEMICAL

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1412

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8

L"

6

<

4

0

9 CER

PIN

BS

ME

NIL PONS HIP

COR

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MBH POA

Tissue (10 I.tg protein)

FIG. 6 Tissue and subcellular distribution of PAM activity in the sheep brain. Data are expressed as the percent conversion of radiolabeled substrate by 10/zg of tissue protein in the absence of unlabeled substrate. Each value is the mean - SE of four tissue samples from different sheep, assayed in duplicate. The tissues assayed were cerebellum (CER), pineal (PIN), brainstem (BS), median eminence (ME), neurointermediate pituitary (NIL), pons medulla (PONS), hippocampus (HIP), cerebral cortex (COR), anterior pituitary (AP), medial basal hypothalamus (MBH), and preoptic anterior hypothalamus (POA). Black bars, membrane; hatched bars, soluble.

was more than threefold greater than in any other tissue (Fig. 6). The hypothalamus is the site of synthesis of vasopressin, oxytocin, and the hypothalamic releasing factors, as well as other amidated neuropeptides. Intermediate levels were found in most other brain regions, while little or no activity was found in the cerebellum and pineal gland. In most tissues, the membraneassociated fraction contained 40-75% more PAM activity per microgram protein than the soluble fraction; the exceptions to this pattern were the median eminence and neurointermediate lobe of the pituitary, where the concentrations of amidating activity were approximately equivalent in the two fractions. As expected, conversion of substrate was linearly related to sample protein content (data not shown).

Kinetics of Hypothalamic Amidating Enzyme Both the soluble and membrane-associated PAM activities of the medial basal and preoptic anterior areas of the hypothalamus displayed typical Michaelis-Menten kinetics (Fig. 7). The affinity constants (Km) for all four sample types were similar (soluble MBH, 12.9/zM; soluble POA, 12.7/zM; membrane-associated MBH, 13.3 /~M; membrane-associated POA, 12.3 /xM). The maximal velocities (Vmax) for the membrane-associated fractions were 4.7 pmol//xg/hr for the MBH, and 4.8 pmol//zg/hr for the POA. Maximal

[12] PEPTIDE-AMIDATING ACTIVITY

233 MBH (Soluble) POA (Soluble)

.-.i

10 MBH (Membrane) POA (Membrane)

..= 4 2

0

0.5

1 1/S (I.tM)

1.5 -1

2

2.5

FIG. 7 Kinetic studies of hypothalamic PAM activity. Sample (10 p~g of protein) was incubated with radiolabeled substrate in the presence of increasing concentrations (0.3-16.7/xM) of unlabeled substrate. Enzyme velocity was calculated on the basis of the mean percent conversion (n = 4 samples, each assayed in duplicate) of labeled substrate and the total substrate present, and then plotted against substrate concentration in a double-reciprocal plot. See text for Km and Vmaxvalues obtained.

velocities for the soluble fractions were 2.6 pmol//xg/hr for the MBH, and 2.9 pmol/~g/hr for the POA.

Amidating Activity in Plasma To examine the possibility of secretion of active amidating enzyme from hypothalamic neurons into the hypophyseal-portal system, we assayed PAM activity in portal plasma samples from four conscious sheep, and compared the activity with that in jugular vein plasma. Measurable levels of PAM activity were present in plasma (Fig. 8), but the specific activity (i.e., percent conversion of substrate per microgram of protein) was 20- to 25-fold lower than in hypothalamic extracts. In three of four sheep, basal hypophyseal-portal and jugular plasma samples contained similar levels of PAM activity; in a single sheep, basal portal levels were 58% higher than jugular levels (Fig. 8A). As expected, injection of insulin induced severe hypoglycemia in all animals, which was followed by a rise in plasma cortisol. Administration of glucose immediately restored blood sugar, and prompted the return of cortisol levels to baseline. During the course of the experiment, there was no significant change in systemic PAM activity, and no consistent change in portal activity (Fig. 8). It therefore appears unlikely that significant amidating activity is cosecreted with hypothalamic amidated peptides in either the basal or stressed state.

234

II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES A

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FIG. 8 Time course of PAM activity in hypophyseal-portal (0) and jugular ( 9 plasma in four conscious sheep (A-D) in response to insulin (I; 100 U at time 0), followed by glucose (G; 25 ml of a 50% solution at time 90 min). Plasma PAM activity was assayed using 2.86 txl of plasma per duplicate sample, and expressed as percent conversion per milligram protein (A-D, top). Also shown (A-D, bottom) are the plasma glucose (solid line) and cortisol (dashed line) levels (in sheep B, there was insufficient plasma for cortisol assays in samples taken between times -10 and 70 rain).

[12] PEPTIDE-AMIDATINGACTIVITY

235

Conclusions The distribution of PAM activity in the sheep brain was readily determined by the amidation assay. This study revealed some interesting differences in levels of PAM in different brain regions, as well as variations in the subcellular compartmentation (i.e., membrane vs soluble) of the enzyme. Most interesting to us, the concentration of PAM in the hypothalamus was much higher than in the median eminence or neurointermediate pituitary, suggesting that amidation of hypothalamic releasing hormones, at least in the sheep, occurs prior to axonal transport of secretory granules. Thus the amidating enzyme is either removed from the granules or, more likely, is inactivated before the granules reach the nerve terminals. This is supported by the observation that portal plasma did not contain a higher level of amidating activity than peripheral plasma, and that this level did not increase with hypoglycemia, a stressor known to elicit secretion of hypothalamic corticotropin-releasing factor and vasopressin into portal blood.

Summary The amidation assay described in this chapter is a rapid and straightforward method for the estimation of amidating activity. Such a method is useful for the screening of tissues for amidating activity that may suggest the presence of novel amidated bioactive peptides, as may occur in the atrium of the heart. Studies of the distribution of PAM activity may also reveal the subcellular sites of peptide amidation; in the sheep hypothalamopituitary system, levels of amidating activity were much greater at the site of peptide synthesis than at the site of storage and release. Finally, amidation assays can also be used to study the effects of endocrine or other interventions (such as chronic glucocorticoid treatment or acute hypoglycemia) on the levels of amidating activity. Regulation of amidating activity is likely to parallel that of the peptide products, and identification of PAM regulation in tissues such as the heart may suggest physiological roles for as yet unidentified amidated peptides.

References 1. A. F. Bradbury, M. D. A. Finnie, and D. G. Smyth, Nature (London) 298, 686 (1982). 2. M. Tajima, T. Iida, S. Yoshida, K. Komatsu, R. Namba, M. Yanagi, M. Noguchi, and H. Okamoto, J. Biol. Chem. 265, 9602 (1990).

236

II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES K. Takahashi, H. Okamoto, H. Seino, and M. Noguchi, Biochem. Biophys. Res. Commun. 169, 524 (1990). C. C. Glembotski, B. A. Eipper, and R. E. Mains, J. Biol. Chem. 259, 6385 (1984). I. Kato, H. Yonekura, M. Tajima, M. Yanagi, H. Yamamoto, and H. Okamoto, Biochem. Biophys. Res. Commun. 172, 197 (1990). A. G. Katopodis, D. Ping, and S. W. May, Biochemistry 29, 6115 (1990). S. N. Perkins, E. J. Husten, and B. A. Eipper, Biochem. Biophys. Res. Commun. 171, 926 (1990). N. G. Seidah, M. Marcinkiewicz, S. Benjannet, L. Gaspar, G. Beaubien, M. G. Mattei, C. Lazure, M. Mbikay, and M. Chr6tien, Mol. Endocrinol. 5, l l l (1991). S. P. Smeekens, A. S. Avruch, J. LaMendola, S. J. Chan, and D. F. Steiner, Proc. Natl. Acad. Sci. U.S.A. 88, 340 (1991). 10. K. Tatemoto and V. Mutt, Proc. Natl. Acad. Sci. U.S.A. 74, 4115 (1978). 11. A. F. Bradbury and D. G. Smyth, in "Biogenetics of Neurohormonal Peptides" (R. Hakanson and J. Thorell, eds.), p. 171. Academic Press, London, 1985. 2," B. A. Eipper, C. C. Glembotski, and R. E. Mains, Peptides (N. Y.) 4, 921 (1983). 13. K. Mizuno, J. Sakata, M. Kojima, K. Kangawa, and H. Matsuo, Biochem. Biophys. Res. Commun. 137, 984 (1986). 14. J. Sakata, K. Mizuno, and H. Matsuo, Biochem. Biophys. Res. Commun. 140, 230 (1986). 15. R. A. Lew, I. J. Clarke, and A. I. Smith, Endocrinology (Baltimore) 130, 994 (1992). 16. R. A. Lew and A. I. Smith, Clin. Exp. Physiol. Pharmacol. 20, 231 (1993). 17. S. N. Perkins, E. J. Husten, R. E. Mains, and B. A. Eipper, Endocrinology (Baltimore) 127, 2771 (1990). 18. K. M. Braas, D. A. Stoffers, B. A. Eipper, and V. May, Mol. Endocrinol. 3, 1387 (1989). 19. L. Ouafik, V. May, H. T. Keutmann, and B. A. Eipper, J. Biol. Chem. 264, 5839 (1989). 20. O. H. Lowry, N. J. Rosebrough, A. L. Farr, and R. J. Randall, J. Biol. Chem. 193, 265 (1951). 21. P. R. P. Salacinski, C. McLean, J. E. C. Sykes, V. V. Clement-Jones, and P. J. Lowry, Anal. Biochem. 117, 136 (1981). 22. V. May, E. I. Cullen, K. M. Braas, and B. A. Eipper, J. Biol. Chem. 263, 7550 (1988). 23. A. I. Smith, C. A. Wallace, I. J. Clarke, and J. W. Funder, J. Neuroendocrinol. 1, 357 (1989). 24. I. J. Clarke and J. T. Cummins, Endocrinology (Baltimore) 116, 2376 (1985).

[13]

Methods for Studying Carboxypeptidase E Lloyd D. Fricker

Introduction Carboxypeptidase E (CPE) (EC 3.4.17.10, carboxypeptidase H) plays a key role in the biosynthesis of numerous peptide hormones and neurotransmitters. This enzyme, which is also known as enkephalin convertase and carboxypeptidase H, removes basic amino acids from the C terminus of peptides (1, 2), a step usually necessary for the formation of bioactive peptides (3). In contrast to the variety of endopeptidases that initially cleave the peptide hormone precursors at specific cleavage sites, thereby generating the intermediates with the C-terminal basic residues (4, 5), only a single carboxypeptidase has been implicated in the intracellular processing of mammalian peptides (1, 2). In addition to mammals a CPE-like enzyme is also present in many organisms, including fish, frog, shark, and Aplysia (6, 7). However, in Saccharomyces cerevisiae (yeast), an unrelated enzyme, Kexl, performs the same function in the processing of the c~-mating factor peptide (2). Although the focus of this chapter is on CPE, the same enzymatic assays can be used to detect Kexl and other carboxypeptidase B-like enzymes (8-10). In concordance with its important role in the production of numerous peptides, CPE has been detected in all peptide-producing neuroendocrine tissues (1, 2). However, in some tissues it is difficult to measure CPE activity owing to the low levels of this enzyme relative to other enzymes present in those tissues (11). A variety of methods can be used to overcome this problem and it is possible to measure accurately CPE activity in many tissues and cell lines. Several assays have been described for carboxypeptidases that remove basic C-terminal amino acids (9, 12-16). The most widely utilized CPE assays are described in this chapter. These assays make use of a change in solubility incurred when substrate is converted into product. The assays differ in their reporter groups; one uses the dansyl fluorescent group, while the other uses a radioactive label. Both assays are rapid and sensitive, but are not specific for CPE. Thus, additional controls must be performed to ensure that the measured activity is due to CPE. These controls, which are discussed below, include using selective inhibitors and activators at several pH values. In addition to measurements of CPE enzymatic activity, it is often important to quantitate levels of CPE mRNA and protein. Standard methods, Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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such as Northern blots, in situ hybridization, and Western blots, have been used to detect CPE mRNA and protein (17-19). Autoradiography with [3H]guandinioethylmercaptosuccinic acid (GEMSA), a selective inhibitor of CPE (20), has been used to detect the membrane-bound form of CPE in brain and other tissues (21-23). In addition to these standard techniques, the rate of CPE biosynthesis has been measured by following the incorporation of radiolabeled amino acids into CPE, which is subsequently isolated by affinity chromatography (24). This technique is described below, following a description of the enzymatic assay.

General Carboxypeptidase Assay The first assay described for CPE used the fluorescent substrate dansyl-PheLeu-Arg, and measured the formation of dansyl-Phe-Leu by extraction of this product into chloroform (13). The substrate is not very soluble in chloroform, especially if the aqueous phase is acidic, whereas the product is highly soluble in chloroform. Because the wavelengths for the excitation and emission of the dansyl group are within the visible range (approximately 350 and 500 nm, respectively), standard borosilicate glass test tubes can be used for both the initial enzyme reaction and subsequent extraction with chloroform. This permits the rapid measurement of enzyme activity. Since the original description of the enzyme assay using dansyl-Phe-LeuArg for CPE, it was found that this general assay can be used to detect other carboxypeptidases, as well as some endopeptidases (9, 10, 25, 26). A variety of substrates have been tested with CPE and other enzymes. For CPE, the optimal substrate is dansyl-Phe-Ala-Arg, which is cleaved with a higher Vmax and lower Km than the original substrate (27). Thus, dansyl-Phe-Ala-Arg gives a higher "signal" for CPE; this is especially important when assaying CPE in crude tissue homogenates, which can contain other enzymes such as lysosomal carboxypeptidase B (CPB). The lysosomal CPB cleaves both substrates with comparable rates, and the ratio of CPE activity to lysosomal CPB is higher with dansyl-Phe-Ala-Arg than with dansyl-Phe-Leu-Arg (27). The dipeptide "dansyl-Ala-Arg" is a poor substrate for CPE, compared to the other substrates, and the ratio of CPE activity to lysosomal CPB activity is extremely low with this substrate. Dansyl-Ala-Arg has been reported to be a superior substrate for plasma carboxypeptidase N (CPN) and the membrane-bound carboxypeptidase M (CPM) (9). These other carboxypeptidases are maximally active at neutral pH, and therefore the assay conditions are different for CPE and these other carboxypeptidases (9). At the neutral pH assay conditions, endopeptidases that presumably cleave the dansyl-

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Phe-Ala-Arg into dansyl-Phe contribute significantly to "product" formation (dansyl-Phe is soluble in chloroform). Because dansyl-Ala-Arg is a poor substrate for these endopeptidases, the "signal-to-noise" ratio is greater with this substrate for assaying CPM and CPN. However, the endopeptidases in most tissue homogenates do not contribute substantially to substrate hydrolysis under the assay conditions for CPE (i.e., acidic pH). Because the lysosomal CPB is a significant problem under the acidic pH conditions of the CPE assay, the optimal substrate for detecting CPE is dansyl-Phe-Ala-Arg. A similar type of assay has been used to detect a dipeptidyl carboxypeptidase and an endopeptidase. Angiotensin-converting enzyme, a dipeptidyl carboxypeptidase, cleaves dansyl-Phe-Arg-Trp into dansyl-Phe, which is readily extracted into chloroform (25). The substrate dansyl-Phe-Leu-ArgArg-Ala-Ser-Leu-Gly is cleaved at the Arg-Ala bond by an endopeptidase present in adrenal chromaffin granules; the product of this reaction is converted by carboxypeptidase B into the chloroform-soluble dansyl-Phe-Leu (26). This general approach of solubility change can be used to detect a wide variety of proteases, and has even been used to detect the formation of Cterminal amide residues (28).

S y n t h e s i s of D a n s y l - P h e - A l a - A r g Several methods can be used to synthesize dansyl-Phe-Ala-Arg. The original report describing the assay with dansyl-Phe-Leu-Arg produced this substrate by reacting dansyl chloride with the dipeptide Phe-Leu, and then coupling this reaction product to arginine (13). However, there are several disadvantages with this strategy" some commercial preparations of dansyl chloride contain fluorescent impurities that are difficult to remove from the reaction product, the second reaction is difficult owing to large differences in solubility between the reagents, and the overall yield is low because two separate coupling reactions are required. A simpler and more efficient strategy is to couple dansyl-Phe to the dipeptide Ala-Arg (27). Commercial preparations of dansyl-Phe are very pure, and the reagents reasonably compatible in their solubilities, providing a more productive reaction. Also, the reaction is economical in that an excess of the inexpensive dansyl-Phe can be used to drive the reaction with the more expensive Ala-Arg. The major disadvantage with this single coupling reaction is that dansyl-Phe-Ala is not produced; this peptide is useful to generate standard curves for quantitation of the enzymatic reaction. However, this is not a serious limitation because dansylPhe-Ala can be produced enzymatically from dansyl-Phe-Ala-Arg by us-

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ing an excess of purified CPE, or commercially available pancreatic CPB.

Procedure The following procedure describes the large-scale synthesis of dansyl-PheAla-Arg, which can be reduced if required. In general, it is best to avoid exposing the dansyl material to bright light, although the reaction does not have to be performed in complete darkness. Wrapping the reaction flask in aluminum foil is sufficient precaution. Two grams of dansyl-Phe (Sigma, St. Louis, MO) is dissolved in 200 ml of acetonitrile in a flask with a stir bar. An equimolar amount of Nhydroxysuccinimide (0.66 g; Aldrich, Milwaukee, WI) is then added and the solution cooled in an ice bath. A solution of 1.0 g of dicyclohexylcarbodiimide (Aldrich) in 10 ml of acetonitrile is then added slowly over 1 min. (Note: Dicyclohexylcarbodiimide is toxic, and should be handled with the necessary precautions.) The reaction is stirred for approximately 1 hr in the ice bath, and then removed from the ice bath and stirred at room temperature overnight. The flask is then cooled in ice, without stirring, and the precipitate removed by filtration through Whatman (Clifton, NJ) filter paper. The filtered liquid is evaporated to dryness using a rotary evaporator, yielding approximately 2.6 g of a pale yellow solid: dansyl-Phe-N-hydroxysuccinimide. This material is stable for months when stored in a dry environment protected from light. The solvent composition for the coupling reaction to produce the dansylPhe-Ala-Arg is critical, and it is important to maintain the ratio of the two solvents when scaling the reaction up or down. The optimal method is to make two separate solutions: 2.6 g of dansyl-Phe-N-hydroxysuccinimide is dissolved in 40 ml of dimethylformamide (DMF), and 1.0 g of Ala-Arg (Bachem Bioscience, Philadelphia, PA) is dissolved in 8 ml of water. The two solutions are then combined in the reaction flask with a stir bar, being careful to maintain the 5"1 ratio of DMF-water. A precipitate should not form during this step, although if it does, continue adding the reagents and the precipitate may dissolve. Stir the reaction overnight at room temperature. In addition to dansyl-Phe-Ala-Arg product, the major components of the reaction mixture include unreacted dansyl-Phe-N-hydroxysuccinimide, dansyl-Phe, and N-hydroxysuccinimide. Because an excess of dansyl-Phe-Nhydroxysuccinimide is used, there should not be much unreacted Ala-Arg. The dansyl-Phe and dansyl-Phe-N-hydroxysuccinimide are removed by adding the reaction product to 500 ml of 0.1 M HCI and 500 ml of chloroform in a separatory funnel, mixing gently, and then, once settled, removing the

[13] CARBOXYPEPTIDASE E

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chloroform layer (which is discarded). The desired product, dansyl-Phe-AlaArg, is retained in the water layer. This procedure is repeated until little fluorescent material is recovered in the chloroform (approximately three to five extractions). The aqueous layer is then frozen and lyophilized, or neutralized and dried with a rotary evaporator. Further purification of the dansyl-Phe-Ala-Arg can be accomplished by several methods. The extent of purification partly depends on the intended use of the substrate. For some studies, further purification may not be necessary because the major contaminant, N-hydroxysuccinimide, does not interfere with the enzyme reaction. However, for most studies, some additional purification is desirable. High-performance liquid chromatography (HPLC) may provide the highest purity, although most HPLC columns are relatively low in capacity, and other approaches are more suitable to the large-scale preparation of substrate. Preparative thin-layer chromatography (TLC) could also be used, as described for the synthesis of dansyl-Ala-Arg (9). The method routinely employed in this laboratory is chromatography on a silica column. The dried aqueous phase is dissolved in a small volume of methanol (10 ml), and then combined with 4 vol of chloroform (40 ml). This solution is applied to a 4 x 50 cm column containing silica (Sigma), and the column is eluted with approximately 1 liter of a 4:1 ratio of chloroform-methanol, and then with several liters of a 1:1 mixture of these solvents. A small aliquot (1-5 /xl) of each fraction is analyzed by TLC on silica plates (Sigma), using chloroform-methanol (1 : 1). Product is detected by ultraviolet (UV) light. Fractions containing dansyl-Phe-Ala-Arg are pooled and dried with a rotary evaporator. This procedure should yield approximately 2-3 g of dansyl-PheAla-Arg (as the hydrochloride salt, Mr 660), which is stable for several years when stored in a desiccator, protected from light, at either 4 or -20~ Solutions of the substrate are stable for several weeks at 4~ and presumably for much longer when frozen. Some batches of substrate dissolve readily in water at the concentration needed for the enzyme assay (0.5 to 1 mM stock). However, some preparations do not dissolve immediately in water, and it is necessary first to dissolve the substrate in a small volume of 0.1 M HCI (usually 0.1% of the final volume) and then dilute the solution with the required volume of water. For example, to make 20 ml of a 0.5 mM stock solution, dissolve 6.6 mg of dansyl-Phe-Ala-Arg in 20/xl of 0.1 M HC1, and then dilute with 20 ml of water.

Standard Fluorescent Assay The volumes of the various reagents depend on many factors, and the following protocol can be modified as required. One important factor depends on the type of fluorimeter to be used; this will influence the size of the test tube

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DansyI-Phe-Ala-Arg (insoluble in CHCI3) I

Carboxypeptidase E

L IGHT BEAM

--AQUEOUS

CHCI

3

DansyI-Phe-Ala (soluble in CHCI3) + Arg

FIG. 1 The principle of the assay for carboxypeptidase E involves a solubility change incurred when the water-soluble (chloroform-insoluble) dansyl-Phe-Ala-Arg is converted into the chloroform-soluble dansyl-Phe-Ala. The reaction is performed in the absence of chloroform; following the reaction the solution is acidified and a sufficient amount of chloroform is added so that the aqueous phase is above the path of the spectrofluorimeter of the light beam. Following mixing and centrifugation, the amount of product is proportional to the fluorescence of the chloroform phase, which is determined by inserting the tube directly into a fluorimeter.

and the volume of chloroform used for the extraction. Another important factor is the purpose of the experiment; the following protocol uses a substrate concentration of 100/xM, which is two- to fivefold higher than the K m (for CPE). This concentration of substrate will detect changes in either the Km or Vmaxof CPE; both have been found to change under conditions that alter neuropeptide production (29, 30). To detect changes primarily in the Vmax, a higher concentration of substrate is optimal.

Procedure The enzyme reaction is performed in the same test tubes as the subsequent extraction with chloroform and measurement of the fluorescence of the chloroform phase (see Fig. 1). A typical protocol, using either 10 x 75 or 12 x 75 mm borosilicate glass test tubes, is presented. 1. Combine the following reagents in a test tube" Enzyme/tissue sample (150/xl maximum) Sodium acetate (pH 5.5), 1 M Either 10 mM CoC12 o r 10/zM GEMSA (discussed below) Water

25/xl 25/xl 25/xl 125/xl (to 200/zl)

Typically, four to six tubes are used for each sample: two or three with

[13] CARBOXYPEPTIDASE E

243

COC12, which activates CPE, and two or three with GEMSA (guanidinoethylmercaptosuccinic acid; Calbiochem, La Jolla, CA), an inhibitor of CPE. Owing to small variations in the glass test tubes, two or three replicates of each point are recommended, which will typically vary less than 10%. Of course, "blank" tubes should be prepared, containing all reagents except enzyme. 2. Incubate the mixture either on ice, at room temperature, or at 37~ for 5-15 min. This provides maximal activation with the CoCI: and inhibition with the GEMSA (discussed below). 3. Start the enzymatic reaction by the addition of 50/zl of 0.5 mM dansylPhe-Ala-Arg. 4. Incubate the reaction at 37~ for 10 min to 16 hr, depending on the required sensitivity. Although CPE is stable for days at 37~ under some conditions, long incubation times are not always linear, reflecting the presence of proteases that degrade CPE, and the linearity must be determined under the conditions used. 5. Stop the reaction by the addition of 100/zl of 0.5 M HCI. Alternatively, 150/zl of 1 M sodium citrate, pH 3.1, has been recommended as a superior alternative to the HC1 for the dansyl-Ala-Arg assay (9); this has not been tested with the present assay. In addition to stopping the enzyme reaction, the acidification is important for the proper partitioning of the substrate and product. Excess acid can decrease the fluorescence yield; if the assay is being modified, the optimal amount of acid should be determined. 6. Add chloroform to the reaction tube in a fume hood. The volume depends on the size of test tube and the type of fluorimeter. Basically, enough chloroform should be added so that the upper aqueous phase is well above the light beam of the fluorimeter. For a Perkin-Elmer (Norwalk, CT) LS3 fluorimeter, which can hold 12 x 75 mm glass tubes (without a test tube adaptor), 2.2 ml of chloroform is optimal. Amounts of chloroform in excess of this are difficult to keep in the tube during the mixing step (see step 7). For fluorimeters that use 10 x 75 tubes, volumes of chloroform are typically 1 ml. For most fluorimeters, it is easy to check if enough chloroform has been added to push the aqueous phase and the interface between the phases above the light beam; setting the excitation wavelength to 400-500 nm permits detection by eye. Mix the tubes on a vortex mixer for at least 5 sec. This step is important to extract the product, dansyl-Phe-Ala, and should also be performed in a fume hood. Following this step, the subsequent procedures do not need to be performed in a hood because the aqueous phase substantially reduces the rate of evaporation of the chloroform phase. At this point, the fluorescence is stable, and the tubes can be stored overnight if required. 8. Centrifuge the tubes at room temperature for 1-5 min at low speed

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(several hundred g) in a swinging bucket centrifuge. The centrifugation conditions (speed and force) should be sufficient to produce complete separation of the aqueous and chloroform phase, but not so extensive that any precipitate (which can form when using crude tissue homogenates) is pushed into the chloroform phase. 9. Determine the fluorescence of the tubes, without transferring the chloroform to a new tube. (Besides requiring extra effort, separating the two phases would greatly increase the evaporation of the chloroform; this could be hazardous to laboratory personnel unless the fluorimeter was in a fume hood.) The wavelength of excitation is approximately 350 nm, emission 500 nm, although individual fluorimeters tend to vary, and the wavelengths of peak fluorescence should be experimentally determined. Definition of C a r b o x y p e p t i d a s e E A c t i v i t y The relative fluorescence units can be converted to nanomoles of dansylPhe-Ala using a standard curve, generated by incubating various amounts of dansyl-Phe-Ala-Arg with an excess of purified CPE or CPB (Sigma). Carboxypeptidase E activity is defined as the difference in fluorescence between the samples containing CoC12 and the samples containing GEMSA. However, other metallocarboxypeptidases that can cleave the substrate are also activated by CoC12 and inhibited by GEMSA, and additional controls need to be performed (9). Because these other carboxypeptidases are maximally active at neutral pH, an important control is to perform the assay at several pH values. If the activity is due to CPE, the difference between CoC12 and GEMSA should be much lower at pH 6.5 and undetectable at pH 7.0, compared to pH 5.5. Although CPM and CPN are maximally active at neutral pH, the stimulation of these enzyme by Co 2+ is severalfold greater at pH 5.5 than at pH 7 (9). Thus, if some activity is detected at neutral pH, it is likely that CPM or CPN is present, and may be responsible for some of the activity detected at pH 5.5. To increase the selectivity for CPE, the assay can be performed at pH 5.0 in the absence of CoG12 (9), although under these conditions the lysosomal CPB could be a significant problem. One method of reducing the lysosomal CPB involves homogenizing the tissue at pH 8.0, and then centrifuging the sample for 1 hr at 4~ this treatment has been reported to inactivate the lysosomal CPB (31), although it has not been adequately tested. Radioactive Carboxypeptidase E Assays To increase the sensitivity of the assay, it is possible to use a substrate with a radiolabeled group in place of the fluorescent dansyl group. Several

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radioactive substrates have been developed for the detection of CPE activity (10, 15). The general principle of the radioactive assay is the same as for the fluorescent assay; the substrate is insoluble in an organic solvent, whereas the product quantitatively partitions into the organic solvent. Two radioactive substrates for CPE are ~25I-labeled acetyltyrosyl-Ala-Arg ([125I]AcYAR) and [3H]benzoyl-Phe-Ala-Arg ([3H]BzFAR). Of all of the substrates for CPE, [~25I]AcYAR is the most sensitive, being able to detect picogram amounts of CPE after a 1-hr incubation (10). This substrate is also easy to prepare because it can be iodinated by standard methods; in contrast, the [3H]BzFAR substrate is difficult to prepare, although it is commercially available (New England Nuclear, Boston, MA). For both radioactive substrates, the product can be extracted into chloroform, as with the assay with dansyl-Phe-AlaArg. However, it is easier to remove the organic phase if it is above the aqueous phase (chloroform forms the lower phase), and therefore ethyl acetate is used in place of the chloroform. An elegant modification of the assay with [3H]BzFAR, which requires scintillation cocktail, is to use a scintillation fluid that is nonmiscible with water; this fluid is combined directly with the aqueous reaction mixture, and only the product partitions into the organic phase of the scintillation fluid (31). The sensitivity of the assay with [~25I]AcFAR is due to the higher specific activity of the 125I, compared to 3H or to the sensitivity of the dansyl group. However, to make full use of this greater specific activity, one would have to use 100 ~M concentrations of [~25I]AcYAR, which would be approximately 109 cpm/assay tube. The concentration of radiolabeled substrate when realistic amounts are used (10,000 to 100,000 cpm/tube) in a small reaction volume (10-100 ~1) is in the nanomolar range, well below the K m for CPE. At this low substrate concentration, the rate of substrate hydrolysis is far below the maximal rate obtainable with higher substrate concentrations. Thus, the theoretical limit for the increase in sensitivity for the [~25I]AcYAR assay, relative to the other assays, is not easily attainable. The following procedure is a compromise between sensitivity and practicality; if the assay volume is increased, or the amount of labeled substrate is decreased, sensitivity will be less. However, for most purposes, sensitivity is not a primary concern, and the assay can be modified to maximize convenience.

Procedure The assay is performed in 0.5-ml microfuge tubes with tight-fitting caps. AcTyr-Ala-Arg is iodinated by any standard procedure, such as the chloramine-T method (10). The iodinated peptide is separated from unincorporated iodine by a reversed-phase Sep-Pak cartridge (Waters, Milford, MA).

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The iodination mixture is diluted with 1 ml of 0.1% (v/v) trifluoroacetic acid (TFA) and applied to the Sep-Pak cartridge. The cartridge is first washed with 5 ml of 0.1% TFA, sample is loaded, followed by a further wash with 5 ml of 10% (v/v) acetonitrile in 0.1% (v/v) TFA. The iodinated peptide is then eluted with 5 ml of 20% (v/v) acetonitrile in 0.1% (v/v) TFA. This eluate is combined with 5 ml of water and then extracted three times with 10 ml of ethyl acetate. The aqueous phase is stored in aliquots at -20~ until use. 1. An aliquot of iodinated peptide is dried in a vacuum centrifuge, and then resuspended in 0.1 M sodium acetate buffer, pH 5.5, to approximately 10,000 cpm/ixl. 2. The enzyme/tissue sample, in 0.1 M sodium acetate buffer, pH 5.5, is placed in a microfuge tube, along with either CoC12 (final concentration, 2 mM) or GEMSA (final concentration, 2 IxM) in a final volume of 10 txl. 3. The reaction is initiated by the addition of 10 txl substrate to the enzyme/ tissue mixture, and the tubes are incubated at 37~ for the desired time. 4. The reaction is stopped by the addition of 25 txl of 0.5 M HC1. 5. Ethyl acetate (250 txl) is added to each tube. 6. The samples are mixed on a vortex mixer and then centrifuged for 1 min in a microfuge. 7. A 200-txl aliquot of the upper phase (ethyl acetate) is transferred to a new tube, which is then quantitated in a 3, counter. Total enzyme activity is the difference between the samples containing enzyme/tissue and the control tubes ("blanks") containing only buffer and substrate. For crude tissue homogenates, CPE activity is defined as the difference in activity in the presence of 1 mM CoC12 vs 1 txM GEMSA, as discussed above. Because the substrate is hydrolyzed by other carboxypeptidases that are activated by CoG12 and inhibited by GEMSA, it is important to verify that the activity is due to CPE. As discussed above for the fluorescence assay, this can be accomplished by measuring the activity at several pH values; an optimum in the pH 5-6 range is indicative of CPE.

Measurement of Carboxypeptidase E Protein Biosynthesis In addition to measuring levels of CPE activity by the above-described assays, it is useful to measure the rate of biosynthesis of CPE. Because CPE is secreted from neuroendocrine cells along with peptide hormones, the cellular level of CPE activity represents a balance between synthesis and secretion. Likewise, the amount of CPE secreted from a cell can increase owing either to elevated secretion of preexisting protein or to elevated synthe-

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sis of new CPE. To address these possibilities, it is useful to quantitate the rate of CPE protein synthesis by metabolic labeling techniques. While the basic technique of labeling proteins with radioactive amino acids is well known, the usual method of isolating the protein of interest is immunoprecipitation. Although immunoprecipitation can be used to detect CPE, a method of isolating the radiolabeled CPE by affinity chromatography has been found to be more reproducible and sensitive than the immunoprecipitation procedures (18). A single column of the affinity resin p-aminobenzoyl-L-arginine-Sepharose 6B has been used to purify CPE to homogeneity from a variety of tissues and cell lines (18, 27). The high degree of purification attained by the affinity procedure comes from both the selectivity of the binding and the selectivity of the elution conditions. Carboxypeptidase E is specifically eluted from the resin by a shift in pH; other nonspecifically bound proteins remain attached to the column during this pH shift. This leads to purification of 1000- to 10,000-fold in a single step.

Procedure 1. Cells are labeled with [35S]Met, or another labeled amino acid, according to standard techniques. 2. Cells are extracted in 10 mM sodium acetate buffer, pH 5.5; if the soluble and membrane-bound forms of CPE are to be analyzed together, the buffer should also contain 1% (w/v) Triton X-100 and 1 M NaC1. Cell homogenates are centrifuged, and the supernatant applied to the affinity column (step 3). If medium is to be analyzed, the pH must be adjusted to <6 so that the CPE will bind to the affinity column. This is easily accomplished by diluting the medium with 0.1 M sodium acetate buffer, pH 5. 3. Cell extracts or diluted media are applied to the 10-ml capacity polypropylene columns (Bio-Rad, Richmond, CA) each containing an approximately 0.5-ml bed volume of p-aminobenzoyl-Arg-Sepharose 6B (Thomas H. Plummer, Jr., New York State Department of Health, Albany, NY). 4. The flow-through of the columns is collected and passed through the columns again. This can be repeated several times to increase the amount of CPE bound to the column. 5. The columns are washed with 16 ml of 1 M NaC1-1% Triton X-100-0.1 M sodium acetate, pH 5.5. Care should be taken to wash the sides of the column; typically the 16 ml is applied as eight washes of 2 ml each. 6. The column is rinsed with 4 ml (2 x 2 ml) of 10 mM sodium acetate, pH 5.5. This step reduces the concentration of salt and Triton in the column, which could interfere with the analysis of the eluted material on polyacrylamide gels.

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7. Carboxypeptidase E is eluted with 1 ml of 50 mM Tris chloride, pH 8.0, containing 100 mM NaCI and 0.01% Triton X-100. 8. The affinity column eluates are frozen and then dried in a vacuum centrifuge. The dried material is then typically resuspended in 100/A of gel loading buffer and applied to a denaturing 10% polyacrylamide gel, prepared by standard procedures. The analysis of the affinity-purified protein on denaturing gel electrophoresis is important because in some cases minor contaminants are present. These contaminants can often be removed by additional washing steps. In one case, we found that contaminants could be eliminated by applying the samples to the affinity column in buffer containing 1% Triton X-100 and 1 M NaC1, compared to buffer containing only the sodium acetate. One advantage of the affinity column procedure over immunoprecipitation is that the affinity column isolates only forms of CPE that bind substratelike compounds. Inactive forms of CPE would not be detected by this method. Interestingly, the precursor form of CPE (pro-CPE) is enzymatically active, and binds to the affinity column (32). Radiolabeled pro-CPE and CPE are difficult to resolve by gel electrophoresis owing to their similar molecular sizes (approximately 56 and 54 kDa, respectively). Although these proteins can be resolved when gels are analyzed by Coomassie or silver-staining methods, the resolution of the autoradiography process is lower. One method to enhance the resolution of pro-CPE and CPE is to digest the proteins with 70% formic acid overnight at 40~ this procedure cleaves proteins at AspPro sequences (33), which generates CPE fragments of approximately 30 and 20 kDa. This method has been useful to differentiate the 30.3-kDa N-terminal fragment of pro-CPE from the 28.5-kDa N-terminal fragment of CPE (34). In summary, the techniques described in this chapter enable the detection of CPE activity and protein biosynthesis. When combined with other standard techniques, such as Northern and Western blots, these methods provide information regarding the level at which CPE is regulated in various systems. Regulation of CPE has been found to occur at many levels, including secretion (30, 35-37), storage (30, 38, 39), protein biosynthesis (18, 37), and mRNA (39-41). In addition, the CPE gene is transcribed from several sites, some of which are differentially regulated (40). Thus, there are many levels at which CPE production and enzymatic activity can be controlled.

Acknowledgments The development of the procedures described in this chapter was supported in part by NIDA Grant DA04494, NIDA Research Scientist Development Award DA00194, and an Irma T. Hirschl Career Scientist Award.

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References

.

7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28.

L. D. Fricker, Annu. Rev. Physiol. 50, 309 (1988). L. D. Fricker, in "Peptide Biosynthesis and Processing" (L. D. Fricker, ed.), p. 199. CRC Press, Boca Raton, FL, 1991. D. F. Steiner, in "Peptide Biosynthesis and Processing" (L. D. Fricker, ed.), p. 1. CRC Press, Boca Raton, FL, 1991. I. Lindberg and J. C. Hutton, in "Peptide Biosynthesis and Processing" (L. D. Fricker, ed.), p. 141. CRC Press, Boca Raton, FL, 1991. L. Devi, in "Peptide Biosynthesis and Processing" (L. D. Fricker, ed.), p. 175. CRC Press, Boca Raton, FL, 1991. R. B. Mackin and B. D. Noe, Endocrinology (Baltimore) 120, 457 (1987). L. D. Fricker and E. Herbert, Brain Res. 453, 281 (1988). L. Latchinian-Sadek and D. Y. Thomas, J. Biol. Chem. 268, 534 (1993). R. A. Skidgel, Methods Neurosci. 6, 373 (1991). L. D. Fricker and L. Devi, Anal. Biochem. 184, 21 (1990). L. D. Fricker, S. Supattapone, and S. H. Snyder, Life Sci. 31, 1841 (1982). J. E. Folk, K. A. Piez, W. R. Carroll, and J. A. Gladner, J. Biol. Chem. 235, 2272 (1960). L. D. Fricker and S. H. Snyder, Proc. Natl. Acad. Sci. U.S.A. 79, 3886 (1982). V. Y. H. Hook, L. E. Eiden, and M. J. Brownstein, Nature (London) 295, 341 (1982). G. Stack, L. D. Fricker, and S. H. Snyder, Life Sci. 34, 113 (1984). B. G. Grimwood, A. L. Tarentino, and T. H. Plummer, Jr., Anal. Biochem. 170, 264 (1988). L. D. Fricker, C. J. Evans, F. S. Esch, and E. Herbert, Nature (London) 323, 461 (1986). L. D. Fricker, J. P. Adelman, J. Douglass, R. C. Thompson, R. P. von Strandmann, and J. Hutton, Mol. Endocrinol. 3, 666 (1989). L. D. Fricker, B. Das, and R. H. Angeletti, J. Biol. Chem. 265, 2476 (1990). L. D. Fricker, T. H. Plummer, Jr., and S. H. Snyder, Biochem. Biophys. Res. Commun. 111, 994 (1983). D. R. Lynch, S. M. Strittmatter, and S. H. Snyder, Proc. Natl. Acad. Sci. U.S.A. 81, 6543 (1984). D. R. Lynch, S. M. Strittmatter, J. C. Venable, and S. H. Snyder, J. Neurosci. 6, 1662 (1986). D. R. Lynch, S. M. Strittmatter, J. C. Venable, and S. H. Snyder, Endocrinology (Baltimore) 121, 116 (1987). R. S. Klein, B. Das, and L. D. Fricker, J. Neurochem. 58, 2011 (1992). M. S. Kapiloff, S. M. Strittmatter, L. D. Fricker, and S. H. Snyder, Anal. Biochem. 140, 293 (1984). S. Supattapone, S. M. Strittmatter, L. D. Fricker, and S. H. Snyder, Mol. Brain Res. 3, 173 (1988). L. D. Fricker and S. H. Snyder, J. Biol. Chem. 258, 10950 (1983). A. F. Bradbury and D. G. Smyth, in "Peptide Biosynthesis and Processing" (L. D. Fricker, ed.), p. 231. CRC Press, Boca Raton, FL, 1991.

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II IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES 29. V. Y. H. Hook, L. E. Eiden, and R. M. Pruss, J. Biol. Chem. 260, 5991 (1985). 30. L. D. Fricker, B. J. Reaves, B. Das, and P. S. Dannies, Neuroendocrinology 51, 658 (1990). 31. J. Rossier, E. Barres, J. C. Hutton, and R. J. Bicknell, Anal. Biochem. 178, 27 (1989). 32. D. Parkinson, J. Biol. Chem. 265, 17101 (1990). 33. G. Allen, in "Sequencing of Proteins and Peptides" (R. H. Burdon and P. H. Knippenberg, eds.), p. 81. Elsevier, Amsterdam, 1989. 34. L. D. Fricker and L. Devi, in "Innovations on Proteases and Inhibitors" (F. X. Aviles, ed.), pp. 259-277. de Gruyter, Berlin, 1993. 35. R. E. Mains and B. A. Eipper, Endocrinology (Baltimore) 115, 1683 (1984). 36. V. Y. H. Hook and L. E. Eiden, Biochem. Biophys. Res. Commun. 128, 563 (1985). 37. B. Das, E. L. Sabban, E. J. Kilbourne, and L. D. Fricker, J. Neurochem. 59, 2263 (1992). 38. S. M. Strittmatter, D. R. Lynch, E. B. De Souza, and S. H. Snyder, Endocrinology (Baltimore) 117, 1667 (1985). 39. L.D. Fricker, R. J. Rigual, E. J. Diliberto, Jr., and O. H. Viveros, J. Neurochem. 55, 461 (1990). 40. D.R. Smith, C. J. Pallen, D. Murphy, and L. Lim, Mol. Endocrinol. 6,713 (1992). 41. O. Grigoriants, L. Devi, and L. D. Fricker, Mol. Brain Res. 19, 161-164 (1993).

[14]

Characterization of Endothelin-Converting Enzymes Terry J. Opgenorth, Sadao Kimura, and Jinshyun R. Wu-Wong

Introduction Much research is focused on endothelium-derived factors with the expectation that these factors represent a new level of cardiovascular regulation with potential importance in circulatory homeostasis. Biomedical researchers have witnessed an explosion in activity related to the potent endothelialderived constrictor peptide endothelin (ET). Many aspects related to the biological activity of ET have been thoroughly reviewed (1-4). Understanding ET biosynthesis remains an important area of research. This chapter reviews the efforts aimed at identifying and characterizing the putative endothelin-converting enzyme (ECE), with special focus on the principal methodologies employed. Although ET has primarily been of interest to researchers in the cardiovascular area, a significant amount of literature has also been generated relating to the potential role of ET in nervous system function. This subject has been reviewed by Gulati and Srimal (5). In brief, there is some suggestion that the ET system in the brain may be different from that characterized in relation to systemic vascular function. The ET isoforms, receptor subtypes, and ECE activity are uniquely distributed within specific brain regions. Available data suggest that ET may have a role as a neuroregulatory factor in the central nervous system (CNS) and may function as a pathogenic agent in a number of neurological conditions.

History Following the discovery of endothelium-derived relaxing factor (EDRF) (6), the endothelium was also shown to produce constricting factors (EDCFs) (7). Yanagisawa et al. (8) identified 1 EDCF as a 21-amino acid bicyclic peptide and named it endothelin. The synthetic peptide was found to be a powerful constrictor of coronary arteries in vitro and a potent hypertensive agent in vivo. The same investigators (9) subsequently demonstrated that Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES PREPROENDOTHELIN

N H 2 - ~ E [ [ ~ ~ 1

17-18

~

]-COOH

52-53 / 73-74 90-91 T

212

PROENDOTHELIN

NH-,/11-COOH ~

endothefin converting enzyme

B|(; ENDOTHELIN ,.,

I

T ENDOTHELIN (ET-I) M.W. = 2492 I , , I NH2-CYS-SER-CYS-SER-SER-LEU-MET-ASP-LYS-GLU-CYS-VAL-TYR-PHE-CYS-HIS-LEU-ASP-ILE-ILE-TRP-COOH

1

3

11

15

21

FIG. 1 The proteolytic processing pathway proposed by Yanagisawa et al. for the generation of endothelin from preproendothelin. The preproform of human endothelin 1 contains 212 amino acids. The signal peptide sequence is represented by the crosshatched section and mature endothelin 1 is represented by the darkly shaded section. The 21-amino acid sequence of the ET-1 isoform and the decapeptide sequence encompassing the scissile bond on which endothelin-converting enzyme acts are both shown. Reproduced from ref. 1.

the human genome contains three ET genes encoding distinct but closely related products: ET-1, ET-2, and ET-3. Of phylogenetic interest is the remarkable homology of the ET sequences with a class of peptide toxins (sarafotoxins) isolated from the venom of the snake Atractaspis engaddenis (10). Yanagisawa et al. (8) cloned the porcine ET gene and, on the basis of cDNA sequencing, predicted the amino acid sequence for preproendothelin. The amino-terminal cysteine of ET is preceded by a dibasic amino acid pair, thereby anticipating cleavage by conventional intracellular hormoneprocessing enzymes to generate big ET (bET). As shown in Fig. 1, freeing the C terminus of the 21-residue ET requires the cleavage of a Trp2~-Va122 bond in bET, suggesting the participation of an endopeptidase with novel specificity. The carboxyl-terminal fragment (CTF) of bET [bET (22-39)], ET, and bET have all been detected in the culture supernatant of endothelial cells and in plasma, supporting the hypothesis that the mature form is generated from bET (1). Although the intracellular localization of ECE is unknown, the signal sequence revealed from the cDNA sequence suggests that at some point in the biosynthetic process the nascent precursor traverses a membrane, where it may be cleaved. The lack of secretory granules in endothelial cells and the finding that very little ET or bET can be identified intracellularly suggest that ET is made constitutively. This concept is supported by various studies demonstrating that induction of ET gene expres-

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sion, manifested by detection of an increase in ET precursor mRNA transcripts, is observed before detection of elevated ET in culture media (1). Several ECE-like enzyme activities representing different endopeptidase classes have been identified. In the original publication by Yanagisawa and co-workers (8), a chymotrypsin-like enzyme was suggested as the putative ECE. Chymotrypsin is a serine protease that cleaves peptide bonds primarily on the carboxyl side of aromatic residues, such as tyrosine, tryptophan, and phenylalanine. Therefore, it could be involved in the cleavage of the Trp 21Va122 bond in bET to yield ET and CTF. McMahon et al. (1 l) reported that the chymotrypsin cleaves not only the TI'pZl-ValZ2bond, but also the Tyr 3~Gly 32bond in the porcine bET molecule. Takaoka et al. (12) further elaborated by showing that the chymotrypsin initially cleaves the Tyr3~-Gly32 bond followed by cleavage of the TrpZl-ValZ2 bond. Chymotrypsin also hydrolyzes mature ET, suggesting that a chymotrypsin-like protease may be involved not only in the production but also in the degradation of ET. More recently, Wypij et al. (13) showed that chymase derived from rat lung mast cells displays ECE-like activity, suggesting that the putative ECE could be a serine protease. One study has identified an ECE-like thiol protease in porcine aortic endothelial cells (14). Regardless, most research has focused on ECE-like activities with either aspartyl protease or metalloprotease characteristics.

Aspartyl Endothelin-Converting Enzymes Aspartyl proteases are known to hydrolyze peptide bonds at the C-terminal side of aromatic amino acids. Takaoka et al. (15) found that preincubation of porcine bET with pepsin converted an inactive molecule into one with potent constricting activity in isolated rat aorta. Further characterization using high-performance liquid chromatography (HPLC) separation of the predominant products of the membrane mixture and amino acid sequencing verified that the two principal products were the 21-amino acid ET peptide and the CTF, thus suggesting that an aspartyl protease with pepsin-like activity may be ECE. The first evidence of an aspartyl ECE-like activity from endothelial cells was provided by Matsumura et al. (16). Incubation of porcine bET with an extract from cultured porcine aortic endothelial cells yielded immunoreactive ET, confirmed by HPLC, with optimal activity at pH 4.0. Formation of mature ET was completely inhibited by the universal aspartyl protease inhibitor pepstatin A [isovaleryl-L-Val-L-Val-Sta-L-Ala-Sta; Sta, (3S,4S)-4-amino3-hydroxy-6-methylheptanoic acid]. A number of laboratories, including our own, have since reported on both membrane- and cytosol-associated aspartyl protease-like ECE activities in a variety of cell types and tissues (17-21).

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This ECE-like activity has a pH optimum of 3.5-5.0 and is exclusively inhibited by pepstatin, with a 50% inhibitory concentration (IC50) in the low nanomolar range. Sawamura et al. (18, 19) found that this ECE has molecular weight characteristics similar to those of cathepsin D, and that cathepsin D can cleave bET to form ET. Furthermore, an antibody raised to bovine spleen cathepsin D absorbed this ECE activity and Western blotting showed the major fractions containing the ECE activity to have immunostainable components with molecular weights similar to that of cathepsin D. Wu-Wong et al. (20, 21) have characterized a pepstatin-inhibitable ECE-like activity from rat lung that may be different from cathepsin D. Both sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and gel filtration under reducing and nonreducing conditions showed the rat lung activity to reside in a single protein band with an apparent molecular mass of 90 kDa, in contrast to cathepsin D from rat kidney or liver, which has an apparent molecular mass of 42 kDa. Knap et al. (22) have also identified an aspartyl ECE in porcine aortic endothelial cells that is unrelated to cathepsin D or renin. While various investigators have established that ET can be formed by cathepsin D, it is also apparent that ET is degraded by cathepsin D (J. R. Wu-Wong, unpublished observations, 1992) (23-25), bringing into question the validity of this aspartyl protease as a physiologically relevant ECE. In contrast to human and rat cathepsin D, human and rat cathepsin E has been found to cleave bET exclusively to form ET and CTF in the absence of other peptidic degradation products, suggesting that this aspartyl protease may bear consideration as a candidate for the putative ECE (25, 26). The role of a cathepsin-like protease in ET biosynthesis has been further questioned because these enzymes are often contained in lysosomes and require a highly acidic environment for activity (pH <5.0). Also, pepstatin has been shown to have no effect on ET-1 release from cultured endothelial cells or plasma ET levels in vivo (27). In contrast, a meeting abstract (28) demonstrated that pepstatin reduced ET levels in an adenocarcinoma cell line, suggesting that this enzyme may be induced under selective, and as yet undefined, conditions.

Metalloprotease Endothelin-Converting Enzymes Ohnaka et al. (29) were the first to report the presence of an endothelial cellderived ECE active at neutral pH. Endothelin-converting activities with pH optima at both 3.0 and 7.0 were identified in a homogenate of cultured bovine endothelial cells. At pH 3.0 various HPLC-separable peptides other than ET or bET were visible, while at pH 7.0 only ET and bET could be identified. The pH 3.0 activity was completely inhibited by pepstatin while the pH 7.0

[14] ENDOTHELIN-CONVERTING ENZYMES

255

activity was insensitive to a wide variety of inhibitors except EDTA and EGTA, suggesting this ECE activity may be a metal-dependent endopeptidase. Another group reported on an ECE activity in bovine carotid artery endothelial cells that was inhibited by phosphoramidon [N-(a-rhamnopyranosyloxyhydroxyphosphinyl)-L-Leu-L-Trp], a neutral (metallo)endopeptidase (NEP 24.11) inhibitor, as well as EDTA and o-phenanthroline (30). Neither thiorphan, another NEP inhibitor, nor captopril, an inhibitor of the neutral metalloprotease angiotensin-converting enzyme (ACE), affected this enzyme, indicating that this ECE may be a unique neutral endopeptidase. Once inactivated with EDTA, the enzyme activity was restored most effectively with Zn 2§ , although addition of other divalent cations was effective. These investigators further determined that the enzyme was membrane bound and active in a narrow range between pH 6.6 and 7.6. When the crude enzyme preparation was separated by gel filtration all the ECE activity eluted in a single peak with an apparent mass of 100 kDa. Ahn (31) reported on similar ECEs from human umbilical and bovine aortic endothelial cells, although the molecular masses were estimated to be 300-350 kDa. Ohnaka (32) described an ECE with a mass of 120 kDa on SDS-PAGE from porcine aortic endothelial cells, purified by sequential lectin affinity and DEAE chromatographies, indicating that ECE may be a glycoprotein. Similarly, Sawamura (33) has shown that porcine lung contains two ECE-like neutral metalloproteases, one a glycosylated protein of 300 kDa very similar to the ECE in endothelial cells and the other a 65-kDa protein that resembles NEP 24.11. A number of additional laboratories have since described the existence of similar membrane-bound endothelial cell-derived ECE activities that are active at neutral pH and inhibited by phosphoramidon (ICs0 of 0.5-1.5/zM) and metal chelators but not by thiorphan (1). Neutral pH-active ECE-like activities have also been identified in the cytosol fraction of cultured endothelial cells, in vascular smooth muscle cells (1), and in the human serum lipoprotein fraction (34). Among the neutral pH-active ECEs described to date, there are a number of inconsistencies that only the ultimate availability of pure enzymes is likely to clarify. Ohnaka et al. (35) reported the successful purification of a phosphoramidon-sensitive ECE by sequential chromatography on DEAE-agarose, Ricinus c o m m u n i s agglutinin 120-agarose, peanut agglutinin-agarose, Mono Q, and TSK GS3000SWxL columns of a Lubrol PX-solubilized membrane fraction from porcine aortic endothelium. The ECE is a glycoprotein, has a molecular mass of 131 kDa, and has enzyme activity that is remarkably similar to that of an ECE purified from rat lung (36). These enzymes have activity profiles that closely agree with the general attributes of the phosphoramidon-sensitive ECE activity characterized by a variety of investigators using crude homogenates. While the purification of ECE is a major

256

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

accomplishment, it was reported at a meeting (Endothelin Inhibitors: Advances in Therapeutic Application and Development, Philadelphia, PA, June 9-10, 1994) that a phosphoramidon-sensitive ECE has been cloned (M. Yanagisawa, Howard Hughs Medical Institute, Dallas, TX). The cDNA predicts an amino acid sequence that is homologous to NEP 24.11 but its activity is different from NEP 24.11 and analogous to that characterized previously for the phosphoramidon-sensitive ECE. This represents a major advance and will certainly provide important new tools for the investigation of ET processing. The physiological relevance of a phosphoramidon-sensitive ECE activity is bolstered by the findings that phosphoramidon inhibits the release of ET from endothelial cells and appropriately alters the ET" bET ratio in media and intracellularly (1). Thiorphan does not inhibit ET release from cultured endothelial cells. Additional importance has been placed on this ECE because of the observation that the systemic and regional hemodynamic effects of intravenously administered bET can be abolished by phosphoramidon (35, 36). The implication is that phosphoramidon is blocking the conversion of the inactive precursor to the biologically important peptide, ET. Thiorphan, kelatorphan, captopril, enalapril, E-64, and leupeptin have little or no inhibitory effect on the in vivo responses to bET. Thus, phosphoramidon acts to inhibit an ECE-like enzyme in vivo that is apparently distinct from NEP 24.11 or ACE, and is not a serine or cysteine protease. It has been shown that phosphoramidon does not compete with ET in isolated rat aortic rings (37). These findings suggest that exogenous bET introduced into the circulation is activated by conversion to ET and that this conversion event is specifically inhibited by phosphoramidon. Because ET is thought to be a paracrine factor in vivo, studies with bET may be favored as an experimental approach to defining the potential physiological role of ET.

E n d o t h e l i n - C o n v e r t i n g E n z y m e S u b s t r a t e Specificity I s s u e s Described above are several enzyme activities that have been shown to cleave bET-1 to form ET-1. In most cases in which substrate specificity has been examined, the enzymes do not cleave the bET-3 isoform with nearly the efficiency that they cleave the bET-1 isoform (1, 26). In contrast, Matsumura et al. (38) reported that a phosphoramidon-sensitive ECE in endothelial cells converts bET-1 and bET-3 to their mature forms with comparable efficiency. Figure 2 shows human sequences for the three bET isoforms. The most important difference between bET-3 and bET-1 and the putative ECE would appear to be the isoleucine replacement of valine at the scissile bond. K. Yorimitsu et al. (unpublished results, 1993) in our laboratory have analyzed the pH dependencies of the conversion of bETs by ECE from

[14]

ENDOTHELIN-CONVERTING 3

257

ENZYMES

1

L, 11

3

21 22

15

21 22

OOH 38

1

11

3

15

"

37

OOH

1

NH 2

E,~EC' 11

15

21 22

HUMAN BIG ET-3 41

COOH

FIG. 2 The amino acid sequences of the three precursor forms (big ET) of human endothelin. Cysteine residues that form disulfide bonds are linked by cross-bridges. Shaded circles represent amino acids that differ from the big ET-1 isoform. In each case, an ECE is thought to cleave the precursor peptide between amino acids 21 and 22 to form the three endothelin isoforms: ET-1, ET-2, and ET-3. Reproduced from ref. 1.

bovine pulmonary artery endothelial cells and found that the optimal pHs for bETs are different: pH 6.8-7.0 for bET-l, and pH 6.4-6.6 for bET-2 and bET-3. Furthermore, the conversion efficiency depends on the amount of substrate (bETs) used because the ratio of immunoreactive ET-1/ET-3 reached nearly 1 at a low substrate concentration, suggesting the possibility that a single ECE may be responsible for the conversion of all bET isoforms. Supporting this idea, Yorimitsu et al. (39) reported that human renal carcinoma cells, producing only ET-2, contain an ET-2-converting enzyme with biochemical characteristics similar to those of ECE previously described in endothelial cells. Regarding substrate recognition by ECE, we found that a decapeptide substrate that contains five amino acids on both sides of the bET-1 scissile bond is not cleaved by either an aspartyl protease or a metalloprotease ECElike enzyme (40), suggesting that portions of the substrate sequence far removed from the scissile bond may be essential for enzyme recognition. Okada et al. (4 !) reported on studies utilizing a variety of truncated substrate sequences. It was found that the C-terminal residues 32-37 of bET-1 are important for conversion by a neutral active ECE from bovine endothelial cells. Of great interest is a molecular biological study reported by Fabbrini et al. (42). They designed and expressed in a frog oocyte a set of prepro-

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II IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

ET-1 mutants including the replacements of Trp 21 to Ala 21, Try 21, or Phe 21, and of Va122 to Asp 22. Interestingly, these four mutants affecting the TrpVal processing site always produced putative ET-1 at levels comparable to the wild type, indicating the importance of conformation rather than sequence for enzyme recognition of the scissile bond. In another molecular biological study on sarafotoxins, Ducancel et al. (43) cloned cDNAs encoding sarafotoxins and elucidated the sequences, finding 12 successive similar stretches of 40 residues, each containing 1 of 6 sarafotoxin isoforms, and all 6 isoforms contained within the precursor molecule. Furthermore, the deduced sequences revealed that only Leu-Cys and Trp-Arg/Lys bonds are found in common at the N and C termini of sarafotoxins, respectively, predicting that the biosynthetic pathways for sarafotoxins are completely different from those of the mammalian ETs, having a dibasic pair at the N terminus and Trp-Val/Ile at the C terminus. Additional studies exploring sequence changes in bET-1 and utilizing pure enzyme will be required to determine definitively the requirements for enzyme recognition. Another interesting observation, albeit with crude enzyme preparations, is that the endothelin turnover rates (Vmax) for the aspartyl ECEs versus the neutral ECEs appear to be different. Using bET-1 as a substrate for a phosphoramidon-inhibitable neutral ECE in bovine endothelial cells, values for Vmaxand K m were estimated by Takada et al. (44) to be 226 pmol of ET1/mg protein/hr and 10.7/xM, respectively. Ohnaka et al. (45), using a similar ECE, found the Vmaxand Km to be 7.9 pmol ET-1/5 x 104 cells/hr and 2.4 /zM, respectively. Ahn et al. (31) also reported similar values (Vmax, 160 and 46 pmol of ET-1/mg protein/hr; Km, 45.4 and 20.9/xM for Human Umbilical Vein Endothelial Cells (HUVEC) and Bovine Aortic Endothelial Cells (BAEC), respectively). The low Vmax values reported for the neutral pH ECEs are unprecedented for enzymes involved in synthetic processing and necessitate that additional caution be applied in considering the physiological importance of these ECEs.

Assay Methods Nearly all investigators have relied on reversed-phase HPLC (RP-HPLC) for quantitative characterization of ECE activity, either as a primary assay or as a means of verifying immunoassay results. Figure 3 is an RP-HPLC trace illustrating the essential peptides assayed (bET, ET, and CTF) to characterize ECE activity (20). Note that the conditions for RP-HPLC used were such that each ET peptide has a retention time that allowed it to be easily distinguished from the others. In this experiment, 3/~g of bET-1 was incubated with a crude homogenate of rat lung (Fig. 3A) or a partially purified fraction (Fig. 3B). It is also of interest to point out that many other peaks are visible, some of which can be identified as assay buffer constituents, but

[14] ENDOTHELIN-CONVERTING ENZYMES

R -9

Big E T

259

ET

Big Er

FIc. 3 Identification of ECE activity in the homogenate and plasma membrane fractions from rat lung: big ET was incubated with 0.1 mg of either homogenate (A) or membrane (B) in the presence of EDTA (10 mM) with a mixture of metal ions (BaC12, MnC12, ZnC12, MgC12, and CaC12, 10 mM each) as described in text. The reaction mixtures were analyzed by RP-HPLC. Reproduced from ref. 20.

a few are apparently fragments of bET-derived peptides. It should be obvious that these fragments are likely to be degradation products of the intact bET, ET, or CTF. Thus, characterization of these fragments is important for differentiation of specific cleavage from nonspecific or degradative cleavage events. Chiou et al. (45a) have described the characterization of two different ECE activities isolated from rat lung membranes. A compound found by screening for potent and selective inhibitors of an aspartyl protease-like ECE (20) was coupled to an affinity gel column (Affi-Gel) that could presumably be used to affinity purify this ECE. A RP-HPLC assay was utilized to identify a phosphoramidon-sensitive ECE in the fraction that passed through the column, and an aspartyl protease-like ECE in the fraction eluted from the column. Further characterization suggested that both enzymes may be different from those previously reported in the literature. Many studies have also employed immunoassays with selective antibodies for bET, ET, or CTF. These assay methods are much easier to perform but suffer from the concern that antibodies may have high cross-reactivity with degradation fragments, posing problems for interpretation of results. More often than not, investigators that rely on immunoassays to characterize ECE

260

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES 16

21

27

HIs-Leu-Asp-Ile-Ile-Trp--VaI-Asn-Thr-P ro-Glu.HIs "ECEdodecapeplide"

g

Ac-HIs-Leu-Asp-Ile-Ile-Trp--VaI-Asn-Th r-Pro-Glu(Edans )-HIs-Gly-OH Assay substrate 1

Ac-(Edans)Glu26-BigET[16-27]-Gly-OH ,~, ECE (or

Ac-His-Leu-Asp-Ile-Ile-Trp 2

tool w t -

838

chymotrypsin)

+ VaI-Asn-Thr-Pro-Glu(Edans).HIs-Gly-OH 3

molwt-

1001

FIG. 4 Substrate 1 is prepared by analogy to the big ET scissile site. Processing by an ECE occurs through hydrolysis of the Trp2~-Va122amide bond, generating fragments 2 and 3. Reproduced from ref. 47.

activity have verified their findings using RP-HPLC methods. A key advantage to immunoassays is the requirement for much less substrate (bET) because the sensitivity of this method is usually 100-fold greater than that of RP-HPLC. Attempts have been made to utilize pseudosubstrates that are amenable to sensitive, quantitative, high-throughput assay methods. On the basis of work by Ng and Auld (46) on fluorescent energy transfer in peptides, yon Geldern et al. (47) proposed that the T r p 21 residue of the bET cleavage site would provide a means of generating an assay utilizing fluorescence detection. The indole moiety of T r p 21 releases fluorescent energy (Yem = 360 nm) on excitation with 290-nm light. This fluorescence can be quenched in the intact substrate by addition of a suitable acceptor group, such as dansyl, in proximity to tryptophan. Physical separation of the fluorophore (tryptophan) and the quencher during enzymatic cleavage will produce an evolving fluorescence, allowing continuous monitoring of the ECE activity. Figure 4 illustrates the construction of a "fluorogenic substrate" for assaying ECE activity and Fig. 5 illustrates the change in fluorescence observed over time when this substrate was incubated with an ECE-like aspartyl protease from rat lung. A similar method has subsequently been employed by Kundu and Wilson (48) to characterize an ECE-like metalloprotease from bovine lung. Another method that has appeal because of its simplicity and suitability to high-throughput formats was described by Matsumura et al. (49). The method is based on the scintillation proximity assay (SPA) principle. The assay utilizes a product capture of 125I-labeled ET generated from 125I-labeled bET, with an ET-specific antibody. The ~25I-labeled ET : antibody complex is then detected in a single-step addition of SPA beads coated with protein

261

[14] ENDOTHELIN-CONVERTING ENZYMES

120-

I10-

I00-

I

0

I

10

I

20

I

30

I

4O

time (rain)

FIG. 5 Cleavage of substrate 1 by a crude rat lung preparation. The substrate (20 ~M in 0.3 ml of assay buffer, pH 7.4) was incubated with 20 ~1 of the crude ECE preparation at 37~ Fluorescence was recorded at 360 nm (hex = 290 nm). Reproduced from ref. 47. A. Only the 125I-labeled ET bound to protein A-coated SPA beads is able to generate a signal. Thus, the assay is homogeneous, requiring no separation steps. These investigators utilized this assay to characterize an ECE-like metalloprotease from porcine cultured endothelial cells and rat lung. Bioassays have played an important role in the characterization of ECE activities to date. The first report indicating that a chymotrypsin-like enzyme may be responsible for ET biosynthesis employed a bioassay exclusively. McMahon et al. (11) demonstrated simply that chymotrypsin activated bET. Because bET is less than 100-fold as potent as ET in contracting isolated tissue or producing an increase in blood pressure in vivo, this is a useful means of verifying that the putative ECE of interest is capable of producing a biologically relevant response. Adaptations of these bioassays have been employed by a wide range of investigators. However, the tissue bath contraction assay and in vivo blood pressure assays are not amenable to handling the large numbers of samples required to follow enzyme activity through purification protocols and to characterize enzyme kinetics. One bioassay method that may be more suitable for enzyme characterization studies is

262

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IMMUNOLOGICAL AND BIOCHEMICAL APPROACHES

described by Warner et al. (50), in which ET-stimulated elevation of cGMP in LLC-PK~ cells is utilized as a biological detector system for the conversion of bET to ET. Using this method ECE activities have been characterized in cultured endothelial cells and brain tissue (51, 52). Finally, although not directly relevant to assay methods, molecular biology techniques may be useful in the study of ECE. Kondo et al. (53) reported on studies in which preproET-1 cDNA was transfected into various cell lines. They were able to demonstrate that a human endothelial cell line was capable of synthesizing mature ET but a B cell line (lymphocyte derived) was not, despite high expression of prepro-ET mRNA. Thus, the B cells apparently lack the ability to translate or process prepro-ET. Employing similar methodology, Benatti et al. (54) demonstrated that transfection of Spodoptera frugiperda cells (insect) with human prepro-ET cDNA produced cells that released mature ET (quantitated by RP-HPLC and bioassay). Despite the fact that no ET or ET-related peptide has been reported to exist in insect cells, these cells apparently have the required processing enzymes to produce mature ET. Such studies may provide important information about the identity of the as yet uncharacterized ECE and, alternatively, may yield unique systems to study ECE that are devoid of other confounding activities.

Summary In conclusion, we have reviewed here the main ECE activities that have been identified and characterized. Presently, primary emphasis is on investigation of a metalloprotease that is inhibited by phosphoramidon but is not inhibited by a number of other inhibitors of neutral metalloproteases. Such an enzyme has now been purified and cloned. However, much work needs to be done to establish that this is the ECE of paramount importance. Advances in the area are likely to result from development of molecular probes that will enable further investigation of the endothelin system.

Acknowledgments The authors wish to acknowledge the many contributions made to our understanding of endothelin-converting enzyme, both in the laboratory and through discussion, by G. Budzik, W. Chiou, John DeBernardis, E. Devine, B. Divish, W. Holleman, C. Mauselle, M. Nishiyama, E. Novasad, D. Pollock, K. Shiosaki, O. Shinmi, A. Tasker, T. von Geldern, K. Yorimitsu, and R. Zaragoza. The authors thank Jan Rundgren for expert secretarial assistance. This work was supported in part by NIH Grant HL44512.

[14] ENDOTHELIN-CONVERTING ENZYMES

263

References

10. 11. 12. 13. 14. 15. 16. 17. 18. 19.

20. 21. 22. 23. 24.

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II IMMUNOLOGICALAND BIOCHEMICAL APPROACHES 25. W. E. Lees, S. Kalinka, J. Meech, S. J. Capper, N. D. Cook, and J. Kay, FEBS Lett. 273, 99 (1990). 26. P. S. Robinson, W. E. Lees, J. Kay, and N. D. Cook, Biochem. J. 284,407 (1992). 27. P. P. Shields, T. A. Gonzales, D. Charles, J. P. Gilligan, and W. Stern, Biochem. Biophys. Res. Commun. 177, 1006 (1991). 28. X. Y. Li, FASEB J. 7, A130 (1993). 29. K. Ohnaka, R. Takayanagi, T. Yamauchi, H. Okazaki, M. Ohashi, F. Umeda, and H. Nawata, Biochem. Biophys. Res. Commun. 168, 1128 (1990). 30. K. Okada, Y. Miyazaki, J. Takada, K. Matsuyama, T. Yamaki, and M. Yano, Biochem. Biophys. Res. Commun. 171, 1192 (1990). 31. K. Ahn, K. Beningo, G. Olds, and D. Hupe, Proc. Natl. Acad. Sci. U.S.A. 89, 8606 (1992). 32. K. Ohnaka, M. Mishikawa, R. Takayanagi, M. Haji, and H. Nawata, Biochem. Biophys. Res. Commun. 185, 611 (1992). 33. T. Sawamura, O. Shinmi, N. Kishi, Y. Sugita, M. Yanagisawa, K. Goto, T. Masaki, and S. Kimura, Biochim. Biophys. Acta 1161, 295 (1993). 34. T. Ohkawa, H. Sakai, and Y. Hirata, FEBS Lett. 320, 165 (1993). 34a. K. Ohnaka, R. Takayanagi, M. Nishikawa, M. Haji, and H. Nawta, J. Biol. Chem. 268, 26759 (1993). 34b. M. Takahashi, Y. Matsushita, Y. Iijima, and K. Tanzawa, J. Biol. Chem. 268, 21394 (1993). 35. E. G. McMahon, M. A. Palomo, W. M. Moore, J. F. McDonald, and M. K. Stern, Proc. Natl. Acad. Sci. U.S.A. 88, 703 (1991). 36. D. M. Pollock and T. J. Opgenorth, Am. J. Physiol. 261, R257 (1991). 37. D. M. Pollock, E. I. Novosad, B. J. Divish, N. S. Burres, and T. J. Opgenorth, FASEB J. 5, A399 (1991). 38. Y. Matsumura, Y. Tsukahara, K. Kuninobu, M. Takaoka, and S. Morimoto, FEBS Lett. 305, 86 (1992). 39. K. Yorimitsu, O. Shinmi, M. Nishiyama, K. Moroi, Y. Sugita, T. Saito, Y. Inagaki, T. Masaki, and S. Kimura, FEBS Lett. 314, 395 (1992). 40. J. R. Wu-Wong, K. Shiosaki, T. P. Dillon, T. R. Rockway, E. M. Devine, G. P. Budzik, C. A. Marselle, and T. J. Opgenorth, FASEB J. 5, A698 (1991). 41. K. Okada, J. Takada, Y. Arai, K. Matsuyama, and M. Yano, Biochem. Biophys. Res. Commun. 180, 1019 (1991). 42. M. S. Fabbrini, A. Vitale, E. Pedrazzini, G. Nitti, M. Zamai, M. Tamburin, V. R. Caiolfa, C. Patrono, and L. Benetti, Proc. Natl. Acad. Sci. U.S.A. 90, 3923 (1993). 43. F. Ducancel, V. Matre, C. Dupont, E. Lajeunesse, Z. Wollberg, A. Bdolah, E. Kochva, J. C. Boulain, and A. Menez, J. Biol. Chem. 268, 3052 (1993). 44. J. Takada, K. Okada, T. Ikenaga, K. Matsuyama, and M. Yano, Biochem. Biophys. Res. Commun. 176, 860 (1991). 45. K. Ohnaka, R. Takayanagi, T. Yamauchi, F. Umeda, and H. Nawata, Biochem. Int. 23, 499 (1991). 45a. W. J. Chiou, K. Shiosaki, A. S. Tasker, and J. R. Wu-Wong, Life Sci. 54, 1613 (1994). 46. M. Ng and D. S. Auld, Anal. Biochem. 183, 50 (1989).

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47. T. von Geldern, W. Holleman, and T. J. Opgenorth, Pept. Res. 4, 32 (1991). 48. G. C. Kundu and I. B. Wilson, Life Sci. 50, 965 (1992). 49. Y. Matsumura, T. Umekawa, H. Kawamura, M. Takaoka, P. S. Robinson, N. D. Cook, and S. Morimoto, Life Sci. 51, 1603 (1992). 50. T. D. Warner, J. A. Mitchell, P. D'Orleans-Juste, K. Ishii, U. FOrstermann, and F. Murad, Mol. Pharmacol. 41, 399 (1992). 51. T. D. Warner, H. H. H. W. Schmidt, J. Kuk, J. A. Mitchell, and F. Murad, Br. J. Pharmacol. 106, 505 (1992). 52. T. D. Warner, G. P. Budzik, T. Matsumoto, J. A. Mitchell, U. F6rstermann, and F. Murad, Br. J. Pharmacol. 106, 948 (1992). 53. M. Kondo, N. Ishida, M. Kobasyashi, and Y. Mitsui, Biochim. Biophys. Acta 1134, 242 (1992). 54. L. Benatti, L. Cozzi, M. Zamai, M. Tamburin, F. Vaghi, V. Caiolfa, M. S. Fabbrini, and P. Sarmientos, Biochem. Biophys. Res. Commun. 186, 753 (1992).

Approaches for Studying Peptide Processing

In Vivo

Arthur Shulkes

Introduction Because much of the processing of peptides occurs within various compartments of the cell and the secretory granules, an in vivo approach for examining processing has not been generally used. However, an important issue that is often overlooked is that there may be further processing between the cell of origin and the target site. This caveat applies to all forms of intercellular communications: autocrine, paracrine, endocrine, and neurotransmitter. In general an in vivo approach has been confined to measuring what has been termed postsecretory or extracellular processing and this is the emphasis of this chapter. A number of different patterns of processing and secretion can be classified. As reviewed by Carraway and Loh (1), the peptide can be stored in secretory granules and released by exocytosis as the bioactive, fully processed peptide. Alternatively, there may be constitutive release of the bioactive peptide. If the peptide is stored as an inactive or partially active precursor, the options include activation at the site of secretion, in the circulation or interstitial fluid, or at the target cell. Extracellular processing may result in the generation of fragments with quite different bioactivities from the bioactive parent peptide (2). As well as activation, such as the renin-angiotensin system, in which a series of proteolytic cleavages converts circulating renin substrate to the bioactive octapeptide angiotensin II, post- and parasecretory inactivation occurs. Examples include the enzymes endopeptidase (EP)-24.11 and EP24.15, which degrade a variety of substrates including enkephalin and neurotensin. In vivo approaches can aid in classifying these different processing systems. Another instance in which an in vivo approach is useful is in monitoring the effects on processing of changes in the secretory activities of cells. For instance, modifying gastric lumenal contents by starvation, refeeding, or pharmaceutical inhibition of gastric acidity changes gastrin biosynthesis, gastrin secretion, and gastrin processing (3, 4). The techniques described below have been concerned with the processing of gut regulatory peptides and have generally used sheep as the animal model. 266

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However, they are adaptable for characterizing the processing of any peptide for which sequence-specific analysis is available.

Metabolic Clearance Rate, Production Rate, and Disappearance Rate

Practical Considerations These studies can be performed in humans (5), although there may be ethical issues in infusing the peptide. A large animal such as a dog, pig, or sheep is best because it allows multiple blood samples during and after the infusion period (6). In addition, sufficient blood can be taken to allow chromatography. Ideally the structures and proportions of the peptides in the tissue of origin and in the circulation should be known. Differences in ratios suggest either preferential secretion, differential metabolism, or processing during secretion. These can be distinguished by the combination of clearance studies, organ-specific metabolism (Section III), and sampling close to the site of secretion (Section VIII). If the study is to be physiologically relevant, the peptide being infused should be identical with that detected in the circulation. For instance, modifications such as tyrosine sulfation (7) can affect clearance. The clearances for all the major circulating forms should be determined because conclusions about biopotency and secretory activity are critically dependent on relative clearances. An important issue that is often overlooked is the stability of the peptide in the sample tube between the time that the blood sample is drawn and when plasma is separated and frozen. If peptide added to plasma has an in vitro disappearance time of around 1 min, then methods to prevent postcollection losses are essential. Addition of proteolytic inhibitors or acid may be sufficient (8); alternatively, collection of blood directly into a denaturing agent such as ethanol has proved effective (9).

Animal Preparation The sheep is laid on a cannulating table, the neck shaved, and the area above the jugular vein infiltrated with local anesthetic. Cannulas are inserted through a 14-gauge needle into the left jugular vein (toward the heart for infusion) and into the right jugular vein (10 cm toward the head) for blood sampling. Although pointing the cannula in opposite directions is not essential, it is imperative that the infusion cannula be downstream from the sampling cannula, otherwise falsely high values will be obtained (6, 10).

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Peptides The peptides, if purchased, should be supplied with an amino acid analysis, and purity specification. The peptide is weighed out gravimetrically on a microbalance and dissolved in 0.05 M acetic acid containing 0.1% (v/v) serum albumin to prepare stock solutions of 50 to 100 nmol/ml. These are stored at -70~ until use. In general each stock is thawed only twice. Most peptides are soluble in acetic acid but for acidic peptides, such as gastrin, the stock solution should be prepared in 0.05 M NH4CO3, 0.1% serum albumin. If solubility remains a problem then the peptide should first be dissolved in dimethyl sulfoxide (DMSO).

Infusion To avoid losses by nonspecific adhesion to syringes and cannulas, the peptide should always be dissolved in a protein-containing solution: 0.1% (w/v) bovine serum albumin (BSA) in 0.9% (w/v) NaC1 is used for animal experiments and human serum albumin for experiments in humans. Additional sterile filtration is necessary for studies in humans. After mixing and drawing the infuseate into the infusion syringe, a sample is taken for radioimmunoassay. An additional sample is taken at the end of the infusion.

Experimental Design To ensure that equilibrium is attained, the infusion should continue for six half-lives. Thus for a peptide with a disappearance half-life of 10 min, a 60min infusion is desirable. For peptides with substantially longer half-lives, an initial loading dose can shorten the required time to achieve equilibrium. The loading dose equals (desired equilibrium concentration) x Vd x bodyweight, where Vd is the volume of distribution, in milliliters per kilogram. Of course, for peptides for which the half life or Vd is unknown, pilot studies are necessary. Separate experiments using high and low doses are useful in determining whether the clearance mechanisms are saturated. Figure 1 demonstrates that during the infusion of calcitonin-gene-related peptide (CGRP), equilibrium is achieved within 75 min. As an example, for a 60-min infusion, blood samples (6 ml) are taken at -20 and -1 min and at +25, +50, +55, and +60 min during infusion. To determine the half disappearance time (tv2) of the peptide, the infusion is

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l

400

o

300

E 13. n" 20O

L9

i

~

E rl

._

-10 0

30

60

90

120

150

Time (rnin)

FIG. 1 Plasma concentrations of calcitonin gene-related peptide (CGRP) in the pulmonary arteries of ewes during infusion of CGRP at 1 (0) or 5 (.) pmol/kg per minute. Values are means _+ SEM. [Reproduced with permission from Braslis et al. (11).] Reproduced by permission of the Journal of Endocrinology, Ltd.

stopped, the infusion line is clamped, and blood samples are taken at 1, 2, 3, 4, 5, 7, 10, 15, 20, 30, and 60 min. The catheter contents are cleared prior to and after each blood sample with a heparin (50 U/ml)-NaC1 (0.9%, w/v) solution. All blood samples are collected into lithium heparin tubes (or conditions appropriate to a particular assay) and stored on ice until the experiment is completed. The samples are centrifuged and the separated plasma stored at -30~ until assay.

Calculations

The metabolic clearance rate (MCR) is determined by the formula MCR = dose divided by equilibrium concentration, where dose is the concentration of infusate [measured by radioimmunoassay (RIA)] x infusion rate, equilibrium concentration is the mean plasma peptide concentration at 50, 55, and 60 min minus mean basal (samples taken at - 2 0 and - 1 min) concentration. Production rate (PR) is determined by the formula PR = MCR x mean basal plasma concentration, and gives a measure of the amount of peptide being secreted. The t~/2 value is calculated from the postinfusion plasma peptide values,

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corrected for basal concentration and normalized as a percentage of the 60min equilibrium sample concentration. The tl/2 value is given by the formula 0.693 divided by k, where k is the slope of the natural log of the normalized concentrations. The volume of distribution (Vd), which is informative in determining whether the peptide remains within the vascular, extracellular, or total body water compartments, is derived by the formula g d - (MCR x tl/2)/0.693. Organ-Specific Metabolism and Secretion The above-described protocol can be modified to allow determination of the contributions of specific organs involved in metabolism and/or processing. An animal operating theater is necessary, as some of the cannulas need to be placed surgically (6, 10).

Animal Preparation Food is withheld from mature merino cross wethers for 24 hr prior to surgery. The sheep are anesthetized by induction with 8 to 10 ml of 5% (w/v) pentobarbitone and maintained with a 1 to 1.5% (v/v) halothane-oxygen mixture. Catheters are inserted into a carotid artery (CA) and the left and right jugular veins, and secured with purse strings; the right jugular vein catheter is directed toward the heart and the left jugular vein catheter is directed toward the head (HJ). A Swan-Ganz catheter is inserted into the left jugular vein, secured with loose purse strings, and repositioned in the pulmonary artery (PA) before each experiment. With the sheep in the left lateral position, a right subcostal incision is made; catheters are inserted into the right renal vein (RV), right hepatic vein (HV), and portal vein (PV) and secured with purse strings. The abdomen is closed and the three catheters are brought out through a separate stab incision. All catheters are flushed with a heparin-NaC1 solution, wrapped in sterile gauze, and placed in a bag attached to the skin above the right kidney. A postoperative period of 3 to 4 days is allowed for recovery from surgery.

Calculations The extraction across each organ is calculated as Inflow equilibrium concentration - outflow equilibrium concentration Inflow equilibrium concentration

x

100%

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I N VIVO P E P T I D E P R O C E S S I N G 50, Q.

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40

0 '~

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,-

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o e-

20

Ifi ~o a

O,

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Kidney

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A~-P Gut

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PA-A Lung

FIG. 2 Percent disappearance of CGRP across kidney, liver, gut, brain, and lung of sheep during infusion of CGRP at 1 (open bars) and 5 (stippled bars) pmol/kg per minute. A, aorta; R, renal artery; P, portal vein; H, hepatic vein; J, jugular vein; PA, pulmonary artery. Values are means _+SEM. [Reproduced with permission from Braslis et al. (11).] *, p < 0.05; **, p < 0.01 for comparison of inflow and outflow concentrations. Reproduced with permission of the Journal of Endocrinology, Ltd. for head (CA-HJ), gut (CA-PV), liver (PV-HV), lung (PA-CA), and kidney (CA-RV). This procedure determines the extraction ratio across organs. To determine the relative contribution of particular organs, the organ blood flow must also be measured. Assuming that the infusion does not affect regional blood flow, published values are sufficient. However, if the peptide is vasoactive or infused at a vasoactive dose, then blood flow will need to be measured. From these experiments the contribution of specific organs to the overall metabolic clearance rate can be calculated. Figure 2 shows the percent difference of infused CGRP across kidney, liver, brain, gut, and lung and demonstrates that clearance occurs only across the kidney and liver (11). The difference between total clearance and the sum of the individual organ clearance is that attributable to that occurring in plasma. To determine the plasma clearance, the stability of the peptide in plasma should be measured.

Stability in Plasma Peripheral venous blood is collected into lithium heparin tubes from six sheep after an overnight fast. The peptide is added to the plasma from each sheep to give a final theoretical concentration of 300 pmol liter -1. The peptidecontaining plasma is aliquoted into two lots; one lot of plasma is incubated

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at 4~ and the other lot is incubated at 37~ Samples are taken from both incubation temperatures at 0, 15, 30, and 60 min, and at 2, 4, 8, and 24 hr. The incubation is stopped by placing the sample tubes into liquid nitrogen. The samples are kept frozen at -30~ until assay. This experiment determines the role of factors in plasma. However, many proteolytic enzymes are bound to capillary membranes, making this in vitro assessment an underestimation.

Parasecretory Processing For a number of regulatory peptides, the molecular profiles in the tissues and in the outflow differ markedly, implying that there is significant processing during or just after secretion. Alternatively, there may be preferential secretion from a heterogeneous granule population. Two examples and methods of approach are described. Gastrin in the antrum is more than 90% gastrin-17, whereas in the antral vein outflow it is less than 60%, with shorter C-terminal forms comprising the remaining immunoreactivity (12). This suggests a contribution of membranebound peptidases during secretion. Analysis of this phenomenon requires local sampling at the site of secretion and the use of inhibitors to block the putative enzyme. The antral vein is cannulated and blood sampled for assay and chromatography. An inhibitor is then infused and the sampling repeated. Parallel in vitro studies with the peptide incubated in plasma with and without added peptidase or inhibitor confirms the role of the particular peptidase. This type of study demonstrated that endopeptidase 24.11 or a related enzyme was involved in the parasecretory processing of gastrin (12). Comparison is also made with the molecular forms in the antrum. The assay system utilizes three different antisera: C-terminal directed, N-terminal directed, and one requiring the intact gastrin17. Ion-enchange and/or reverse-phase high-performance liquid chromatography (HPLC) is necessary to distinguish the different molecular forms. For gastrin, this postsecretory processing does not affect bioactivity because the C-terminal bioactive part is stable in the circulation. The major fragments have stability similar to the intact molecule, therefore the secretion rate and bioactivity (although not the profile) could be monitored by a single antiserum directed toward the C terminal. In contrast with the 13-amino acid peptide neurotensin, cleavage between Arg-8 and Arg-9 inactivates the peptide because the C-terminal bioactive part is unstable in the circulation, leaving only inactive, long-lasting N terminal metabolites (5, 6). The experimental approach is similar except that the major source of circulating neurotensin is the ileum; therefore the ileal vein is

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cannulated and the molecular profiles between the secreted and stored forms are compared. Because the C-terminal fragment is degraded rapidly to nonimmunoreactive forms, parasecretory processing has a marked effect on bioactivity. Furthermore, both C-terminal and N-terminal assays are required: the C terminal, which in effect measures intact bioactive neurotensin, and the N terminal-directed antiserum, which is a measure of the total production (intact plus metabolites).

Use of Proteolytic Inhibitors and Activators in Vivo The administration of inhibitors of processing enzymes can characterize the processing pathways and define physiological roles of particular peptides. An example of quantitation and regional specificity of processing has been reported by our laboratory (13). Neurotensin and the neurotensin analog kinetensin are metabolized by peptidyl-dipeptidase A (angiotensin-converting enzyme, EC 3.4.15.1). The MCR is determined (Section II). Converting enzyme (CE) inhibitor such as captopril (E. R. Squibb & Sons, Princeton, NJ) is infused into sheep for 48 hr (15 mg/kg every 24 hr) (13) and the clearance rate determined again. The change in MCR gives a measure of the role of CE in the metabolism of neurotensin. By measuring the clearance across specific organs, the role of membrane-bound versus circulating proteases can be assessed. High concentrations of CE are present in the lung and treatment with captopril completely abolished lung clearance without effect on clearance across other organs (13). Thus it is possible to determine the relative contribution of the soluble circulating form of CE, the CE in the general vascular endothelium, and the CE present in lung. It is essential that all measurements for calculation of the MCR be made with region-specific assays and confirmed by appropriate chromatography. In terms of revealing physiological function, an inhibitor of the peptidase and a specific inhibitor of the putative target peptide are required. For instance, inhibition of endogenous neutral endopeptidase 24.11 showed that lung bombesin is involved in the regulation of fetal lung growth (14). In this study an inhibitor of EP-24.11 is administered to pregnant mice for several days and measures of fetal lung growth then determined. Studies are repeated with the addition of a bombesin antagonist (14). In this way the regulatory role of a peptidase on a particular peptide can be determined.

Effects of Hypersecretion on Processing Increased secretory activity of particular cell types can reveal rate-limiting steps in processing and the interactions between synthesis, storage, and

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secretion. The example presented is the stimulation of antral gastrin by inhibition of gastric acidity (3, 4). The measured and derived parameters include plasma and tissue concentration of the various molecular forms, clearance rates, fractional release and conversion rates, and mRNA abundance. Using the same animal model as in Section II, the MCR of gastrin amide is measured. The sheep is then infused for 5 days with the gastric acid inhibitor omeprazole. Daily blood samples are obtained and the MCR is repeated on day 4. On day 5 the sheep is culled and the stomach removed and the antral mucosa quickly dissected, weighed, and frozen at -70~ for peptide and mRNA analysis. To obtain baseline antral tissue values, a group of animals that do not receive any infusion is also culled. Measurements: The concentration of the bioactive amidated gastrin, the precursor forms including carboxy- and amino-terminal extensions, and glycine-extended gastrin are measured in plasma and tissue by specific RIA. Chromatography on selected samples is performed to confirm the RIA specificities. The secretion rate is determined by the formula MCR x basal plasma concentration, and the fractional release rate is determined as the secretion rate divided by the total antral content. Gastrin-specific mRNA is measured by conventional techniques (4). These type of studies allow the balance between synthesis, storage, and secretion to be determined. In addition, the conversion rate of precursors is measured (e.g., the extent of amidation). The particular example also allows an assessment of the role of gastric lumenal factors on biosynthetic processing. For a complete understanding of the interactions, data on the clearances of the precursors as well as the bioactive forms are needed. If this information is not available, these clearance measurements should be performed in a separate control group (10).

Sampling at Site of Secretion In vivo microdialysis allows the local sampling of interstitial fluid. The major advantage of in vivo microdialysis is that it allows a dynamic measurement

from a anatomically discrete area. It can be performed in the conscious animal, there is no direct contact of perfusion fluid with the tissue, and comparison with an in vitro model system allows an estimation of the in vivo concentrations (15). Measurements of neurotransmitter and neuropeptide release, processing, and metabolism have been reported (16, 17). In vivo microdialysis is based on the principle that when a semipermeable

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dialysis fiber is inserted into a tissue and perfused with saline, movement of water and small molecules will occur in the direction of the lowest concentration until equilibrium is reached. Examples of the use of in vivo microdialysis include the measurement of dopamine and cholecystokinin release and metabolism in the central nervous system (CNS) (17, 18), serotonin from the spinal cord (19) and stomach (20), and catecholamines and enkephalins from the adrenal medulla (21). In the gut, in vivo microdialysis is useful for measuring the local release of prostaglandins and the metabolism of peptides (22, 23). The dialysis fibers can be used for administration of inhibitors or precursors as well as sampling. However, studies by this and other laboratories using dialysis fibers implanted in the stomach or intestine failed to detect any release of endogenous peptides. This is probably a reflection of the lower concentrations of neuropeptides in the gut compared to the brain. Nevertheless, the processing or metabolism of exogenous neuropeptides within the stomach or gut can be monitored by implanting two sets of fibers in parallel, infusing via one set and sampling from the other (23). The nature of the processing/degradative pathway can also be assessed by the concurrent infusion of inhibitors. The procedures for in vivo dialysis have been described extensively (15-17) and only some key issues that we have found to be critical are highlighted. To determine equilibrium times and the ratio of concentrations in the dialysate versus the outside, in vitro studies are performed. The fibers are immersed in a bath containing a known concentration of the substance under investigation. The fibers are perfused with saline and samples collected at 30min intervals for 2 hr. The time course and percent recovery are determined. Variables that need to be examined in vitro and repeated in vivo include (a) perfusate flow rate, (b) type, length, and number of dialysis fibers, and (c) osmotic pressure of the dialysate. 1. Rate of perfusion: Flow rates from 0.1 to 20/zl/min have been reported, with the most common being between 2 and 5/zl/min for small probes used in rats and 20/zl/min for probes in large animals. 2. Nature of probe: Either laboratory-fabricated or commercial microdialysis probes can be used. The commercial probe CMA/10 (Carnegie Medicin, Stockholm, Sweden) has a molecular cutoff of 20,000 Da and an in vitro recovery for a 1-mm probe ranging from 2% for neuropeptide Y (NPY) to 20% for angiotensin II (17). Recovery increases with the length of the dialysis fiber: a 5-mm probe has a threefold higher recovery than a 1-mm probe. A table of recoveries has been published (17). Although the membrane has a nominal cutoff of 20,000 Da, recoveries of peptides with molecular weights greater than 5000 is poor. A number of laboratories have described the construction of dialysis probes

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for intracranial and spinal cord use and these include U-shaped, I-shaped, and concentric probes (17, 24). The dialysis membranes used include Spectrapor cellulose fibers, with an approximate molecular weight exclusion of 6000 (Spectrum Medical Industries, Los Angeles, CA) (22) and Amicon (Danvers, MA) Vitafiber TM type 3 with a molecular weight cutoff of 50,000 (21). For studies in sheep we have used from 3 to 10 Spectrapor fibers bundled together and glued each end into a silastic tube. From 1 to 5 cm of fiber remains exposed. The assembly is implanted into the submucosa and flushed daily with saline. In vitro experiments show a 2% equilibrium ratio for gastrin and a 10% ratio for somatostatin. 3. Osmotic pressure of dialysate" For amines, increasing the osmotic pressure of the perfusate had no effect on the equilibrium concentration (20). However, increasing the osmotic pressure with up to 5% (w/v) BSA facilitated the equilibration of bombesin (22) and prostaglandins (23). The effects of protein are dependant on the peptide under investigation. For instance, the in vitro equilibrium concentration of somatostatin is higher with 0.15% albumin than with 5% albumin, whereas protein had no effect on gastrin equilibrium concentrations. In vivo microdialysis has proved to be a useful technique for sampling of amines, amino acids, and prostaglandins from the fluids surrounding their site of release. To achieve a similar success with peptides, recoveries need to be optimized by varying probe type and perfusion medium and the assay system must be at maximum sensitivity. However, the ability to monitor the processing that occurs at the site of release and to influence this by local administration of inhibitors makes in vivo microdialysis a powerful investigative tool. In summary, in vivo approaches are of particular use in determining extracellular processing. Moreover, they are assuming greater importance with the realization that extracellular processing of peptides involves activation as well as inactivation (2).

Acknowledgment Arthur Shulkes is a Principal Research Fellow of the National Health and Medical Research Council of Australia.

References 1. R. E. Carraway and R. P. Loh, Handb. Exp. Pharmacol. 106, 69 (1993). 2. C. J. Molineaux and S. Wilk, in "Peptide Biosynthesis and Processing" (L. D. Fricker, ed.), p. 251. CRC Press, Boca Raton, FL, 1991.

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10. ll. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.

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G. J. Dockray, C. Hamer, D. Evans, A. Varro, and R. Dimaline, Gastroenterology 100, 1187 (1991). M. Read, D. M. Read, M. Kapucscinski, and A. Shulkes, Regul. Pept. 40, 13 (1992). A. Shulkes, S. Bijiphala, J. K. Dawborn, D. R. Fletcher, and K. J. Hardy, J. Clin. Endocrinol. Metab. 58, 873 (1984). A. Shulkes, D. R. Fletcher, and K. J. Hardy, Am. J. Physiol. 245, E457 (1983). S. Pauwels, G. J. Dockray, and R. Walker, Gastroenterology 92, 1220 (1987). V. E. Eysselein, G. E. Eberlein, W. H. Hesse, M. V. Singer, H. Goebell, and J. Reeve, Jr., J. Biol. Chem. 262, 214 (1987). A. Shulkes, D. R. Fletcher, and K. J. Hardy, J. Gastroenterol. Hepatol. 4, 429 (1989). G. Ciccotosto and A. Shulkes, Am. J. Physiol. 263, G802 (1992). K. G. Braslis, A. Shulkes, D. R. Fletcher, and K. J. Hardy, J. Endocrinol. 118, 25 (1988). D. M. Power, N. Bunnett, A. J. Turner, and R. Dimaline Am. J. Physiol. 253, 633 (1987). D. Read, A. Shulkes, D. R. Fletcher, and K. J. Hardy, Agents Actions 38, 231 (1993). K. A. King, J. Hua, J. S. Torday, J. M. Drazen, S. A. Graham, and M. A. Shipp, J. Clin. Invest. 91, 1969 (1993). U. Ungerstedt, in "Measurement of Neurotransmitter Release in Vivo" (C. A. Marsden, ed.), p. 81. Wiley, Chichester, 1984. K. M. Kendrick, J. Neurosci. Methods 34, 35 (1990). B. H. C. Westerink, G. Damsma, H. Rollema, J. B. De Vries, and A. S. Horn, Life Sci. 41, 1763 (1987). M. Takita, T. Tsuruta, Y. Oh-hashi, and T. Kato, Neurosci. Lett. 100, 249 (1989). P. M. Pilowsky, V. Kapoor, J. B. Minson, M. J. West, and J. P. Chalmers, Brain Res. 366, 354 (1986). M. Pairet, O. Meirieu, T. Bardon, and Y. Ruchebusch, Gastroenterology 91, 1250 (1986). H. Jarry, E. M. Duker, and W. Wuttke, Neurosci. Lett. 60, 273 (1985). N. W. Bunnett, J. R. Reeve, Jr., and J. H. Walsh, Neuropeptides (Edinburgh) 4, 55 (1983). N. W. Bunnett, J. H. Walsh, H. T. Debas, G. L. Kauffman, Jr., and E. M. Golanska, Gastroenterology 85, 1391 (1983). L. Hernandez, B. G. Stanley, and B. G. Hoebel, Life Sci. 39, 2629 (1987).

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Section III

Identification and Characterization of Extracellular Processing Enzymes in the Central Nervous System

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[16]

Identification and Characterization of Central Nervous System Peptidase Activities J o h n R. M c D e r m o t t

a n d A l i s o n M. G i b s o n

Introduction Central nervous system (CNS) peptidases* mediate a variety of specialized functions in addition to their role in protein turnover, and much of the work on these enzymes has had its origins in identifying peptidases involved in the biosynthesis and catabolism of neuropeptides. More recently, interest has focused on the involvement of CNS peptidases in the pathogenesis of human neurodegenerative disorders, particularly Alzheimer's disease, where accumulation of extra- and intracellular protein aggregates (/3-amyloid protein and neurofibrillary tangles, respectively) are the pathological hallmarks. Experience indicates that few peptidases are likely to be specific to the CNS and furthermore, many "novel" peptidase activities will be the result of the action of known peptidases on new substrates. In this chapter we first summarize the main experimental approaches. We then describe methods using synthetic peptide substrates to identify and characterize peptidases in human brain. Specific examples are taken from studies (1, 2) to identify peptidases with the specificity to process the Alzheimer's/3-amyloid precursor protein (APP), and the methods were derived from our earlier studies on neuropeptide-degrading peptidases in human brain (3-5).

M e t h o d s to I d e n t i f y N o v e l C e n t r a l N e r v o u s S y s t e m Peptidase Activities

Degradation of Substrate The most direct method is to follow the degradation of the native substrate on incubation with an appropriate tissue fraction. For neuropeptides, the availability of large amounts of the synthetic peptide makes it relatively straightforward to isolate and characterize the degradation products and * We use the term peptidasefor all enzymes that hydrolyze a peptide bond; it thus includes endo- and exopeptidases and proteinases. Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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thus determine the cleavage points. The peptidase activities can then be characterized (sensitivity to inhibitors and activators, pH optimum, distribution, etc.) and purified from the tissue. This approach has led to the identification of several relatively broad-specificity peptidases in the CNS. Identification of peptidases that cleave at specific bonds in proteins presents a greater challenge. Sufficient quantities of pure protein may not be available to allow routine assays. Furthermore, incubation of the protein with crude tissue fractions exposes many bonds to a large number of peptidases, and unless some predictions can be made about the properties of the peptidase (e.g., localization, inhibitor sensitivity), it will be difficult to identify the specific peptidase activity under investigation. When the site of cleavage of the protein is known, synthetic peptides spanning the cleavage site can be used as substrates to identify candidate peptidase activities. The main perceived drawback of this approach is that the conformation of the intact protein may be important in directing the specificity of the peptidase and short peptide sequences from the protein may lack this information. However, many protein-degrading peptidases do appear to degrade short peptide sequences with the same specificity as for the protein, although the turnover number (a measure of the efficiency with which an enzyme acts on a substrate) is often reduced. A more general approach is to screen for peptidase activity against relatively nonspecific substrates. Amino acids or small peptides linked via a Cterminal amide bond to a chromogenic (e.g., p-nitroanilide, pNA) or fluorogenic (e.g., 7-amino-4-methylcoumarin, AMC) compound provide convenient substrates for the rapid spectrometric assay of large numbers of samples. The amino acid composition of the substrate confers some specificity but only through the residues upstream of the cleavage site. Denatured proteins (e.g., casein) have been widely used to screen for proteolytic activity; they are readily hydrolyzed into acid-soluble fragments by a wide variety of peptidases and convenient, sensitive assays can be constructed by using labeled proteins (e.g., azocasein). The power of methods based on substrate hydrolysis to identify individual peptidase activities is greatly increased by chromatographic or electrophoretic separation of the peptidases in the tissue fraction before the assay. As well as resolving individual peptidases, this can also separate endogenous inhibitors and substrates from the peptidase, which often interfere with peptidase assays in crude tissue extracts. Chromatographic methods are described below. The high resolving power of polyacrylamide gel electrophoresis is also useful and has been applied to human brain (6). The resolved peptidase activities can be detected in situ by degradation of a protein substrate copolymerized with the gel or after transfer to a second gel or membrane [see

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Sarath et al. (7) for methods]. These methods are obviously not applicable to enzymes composed of heteromeric subunits, or to enzymes that are irreversibly denatured under the electrophoretic conditions.

Binding of Peptidase to Substrate or Inhibitor Affinity chromatography can be used to isolate peptidases from crude tissue extracts by their retention on columns of immobilized substrate or inhibitor. An example of this technique was the isolation of a chymotrypsin-like peptidase from brain, using immobilized al-antichymotrypsin (8). A novel method to identify specific peptidases involves adding a cross-linking reagent to a mixture of radiolabeled substrate and tissue extract. Peptidases in contact with the substrate are covalently bound and thus radiolabeled, and can be isolated by chromatography and electrophoresis. This method was used to identify a brain peptidase capable of generating the N terminus of the Alzheimer's ~-amyloid protein from its precursor (9). A drawback of all methods relying on interaction of peptidase with substrate is that some enzymes are present in tissues as inactive proenzymes and require conversion, usually by limited proteolysis, to the active form before substrate interaction takes place.

Molecular Biological Approaches Molecular biological techniques have added a new dimension to the search for peptidases. Protein sequence information obtained from cDNA libraries has revealed that a great many proteins are processed by proteolytic cleavage after translation and has allowed the sites of cleavage within proteins to be pinpointed. This is the starting point for much of the current work on peptidase identification. The specificity of the peptidase activity can be defined by introducing single amino acid substitutions at the site of cleavage in target proteins, using site-directed mutagenesis in suitable cell lines. The sequences of many peptidases are now known and have revealed families related by sequence homology, mechanistic class, and tertiary structure (10). A profitable strategy to identify novel mammalian peptidases has been to use polymerase chain reaction (PCR) techniques to search for mammalian homologs of peptidases of known sequence from other organisms. Although these methods can help to identify novel peptidases, their purification and characterization still require the development of biochemical assays.

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Synthetic Peptide Substrates to Identify Central Nervous System Peptidases This section describes methods to identify peptidases that degrade proteins at specific bonds, using synthetic peptide substrates. Examples are taken from work aimed at identifying and characterizing (a) peptidases that hydrolyze APP (11) at two sites [the N-terminal/3-amyloid generating site, Met596-Asp 597, and the secretase site, Lys612-Leu 613 (12)] and (b) peptidases that degrade/3-amyloid protein (/3-AP).

General Considerations Peptides of 8-16 residues surrounding the cleavage site in the protein are used as substrates. To avoid degradation by aminopeptidases, the peptides are N-acetylated, and C-terminal amides are used to prevent degradation by carboxypeptidases. Peptides having amino acid substitutions associated with increased or decreased susceptibility of the protein to cleavage at the target site are useful. This information may be available from site-directed mutagenesis studies or from disease-related mutations. Analysis of peptide degradation is carried out using automated reversed-phase high-performance liquid chromatography (HPLC). While this is relatively slow, it has the advantage that cleavage at a specific bond can be followed by analyzing the degradation products. Internally quenched fluorogenic peptide derivatives [e.g., 2,4-dinitrophenyl-Pro-Leu-Gly-Pro-DL-2-amino-3-(7-methoxy-4coumaryl)propionic acid for endopeptidase 24.15] (13) offer the convenience of fluorimetric assays; however, they are more difficult to synthesize and assays based on them are not specific because cleavage of any peptide bond gives fluorescence. Fluorescent assays can be constructed using protected peptidyl derivatives in which cleavage at the target site releases an aminoacyl4-methyl-7-amidocoumarin; an aminopeptidase is then used to release the fluorophore. This method is used to follow cleavage at Lys-Leu in the substrate Z-Gln-Lys-Leu-AMC (Z, carbobenzoxy); released Leu-AMC is converted to fluorescent AMC with an aminopeptidase.

Synthesis of Substrates Peptides are synthesized with an Applied Biosystems (Foster City, CA) 431A peptide synthesizer on p-hydroxymethylphenoxymethyl polystyrene resin (free acids) or Rink amide resin (for amides) using Fmoc-amino acids

[16] IDENTIFICATION OF CNS PEPTIDASES

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(1 mmol) and either HBTU activation (FastMoc chemistry) or dicyclohexylcarbodiimide/HOBu / activation, according to the instructions of the manufacturer. The former activation method is recommended because of the shorter coupling times. Acetylation of the N-terminal c~-amino group is achieved using acetic anhydride (10%, v/v) in N-methylpyrrolidone for 20 min at room temperature. Crude peptides [30 mg in 2 ml of 21% (v/v) acetonitrile, 11 mM trifluoroacetic acid (TFA)] are purified by reversed-phase HPLC on an Aquapore ODS column (25 x 1 cm; Brownlee Laboratories) using Waters gradient HPLC equipment, eluting with an acetonitrile gradient (21-49%; 2 ml/min) containing 11 mM TFA. Fractions (0.5 ml) are collected around the main ultraviolet (UV)-absorbent (280 or 214 nm) peak and aliquots (5/zl) analyzed by reversed-phased HPLC (NovaPak C18 cartridge in RCM 8 x 10 unit), using a 2.9-56% (v/v) acetonitrile gradient over 20 min (2 ml/min), containing 11 mM TFA. Fractions containing pure peptide are pooled, dried, and reanalyzed by HPLC to ensure purity. Peptide identity is confirmed by amino acid analysis (Pico Tag method; Waters). Z-Gln-Lys-Leu-AMC is synthesized by standard solution methods as follows: Fmoc-Lys ( / B O C ) - O H (0.6 mmol) is coupled to Leu-AMC (0.6 mmol; HC1 salt plus 1 equivalent of triethylamine) in dichloromethane-dimethylformamide (DCM/DMF, 9: 1; 6 ml) using dicyclohexylcarbodiimide-N-hydroxybenztriazole (DCC1/HOBu/; 1.05 equivalents). The neutral, ethyl acetate-soluble product is treated with 20% (v/v) piperidine in DCM/DMF (4 : 1; 5 ml) to remove the Fmoc group. The resulting Lys(/BOC)-Leu-AMC (0.4 mmol) is coupled to Z-GIn in DCM/DMF (9: 1; 10 ml) using DCC1/HOBu/. The insoluble product is filtered off, washed with acetone, and dissolved in DMF (10 ml). After filtration, the product, Z-Gln-Lys(/BOC)-Leu-AMC, is precipitated with water and dried. To remove the/BOC group, the peptide (200 mg) is dissolved in TFA (2 ml); after 15 min, the TFA is removed under vacuum and the product, Z-Gln-Lys-Leu-AMC/TFA salt, is obtained as a white solid by adding ether. Reversed-phase HPLC shows a single peak, and the amino acid composition is Glu (1.0), Lys (1.0), Leu (1.0). Z-GlnLys-Leu-p-nitroanilide is synthesized by the same scheme (1).

Tissue and Subcellular Fractions We mainly use postmortem human brain for our studies on peptidases related to human neurodegenerative disorders. The possible instability of peptidases postmortem is the main drawback and this may be investigated by comparative studies on fresh rat brain. When the subcellular localization of the peptidase is unknown, the tissue can be separated into crude soluble and

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particulate fractions. It is an advantage to determine at an early stage whether the peptidase activity is maintained after freezing and thawing the tissue. If so (and most appear to be), frozen, neuropathologically assessed brains (cerebral cortex from nonneurological cases) are used from the Newcastle Brain Bank. Synaptosomes and synaptic membranes are used when investigating peptidases associated with neurotransmitter-related functions. They are also useful for studying APP-metabolizing enzymes because high concentrations of APP are present at nerve terminals. Cell lines having neuronal characteristics (e.g., rat phaeochromocytoma cell line PC12 or human neuroblastoma cell line IMR32) are also used, either as intact cells to study cell surface peptidase activity or as a source of subcellular organelles [see Storrie and Madden (14) for methods] that can be obtained more readily than from human brain.

Soluble and Particulate Fractions from Human Brain Postmortem cerebral cortex (2 g) is homogenized (Ultra Turrax homogenizer; Fisons, Loughborough, U.K.) in 5 vol of 50 mM Tris-HC1 (pH 7.4) and centrifuged (27,000 g, 15 min, 4~ to yield soluble and particulate fractions. The particulate fraction is washed twice in homogenization buffer, centrifuging as before, and reconstituted to the original volume in homogenization buffer. If lysosomal peptidases are of particular interest, a lower pH homogenization buffer should be used. As the characteristics of the peptidase activity under investigation become known, the buffer can be modified to optimize activity [e.g., addition of 1 mM dithiothreitol (DTT) for cysteine peptidases]. Preparation of Synaptosomes Synaptosomes are prepared from postmortem human brain or freshly removed rat brain by the rapid method of Dodd et al. (15), using discontinuous sucrose density gradients. Human cerebral cortex is homogenized (using a Teflon/glass homogenizer) in 10 vol of 0.32 M sucrose and centrifuged at 2000 g at 4~ for 10 min in a fixed angle rotor. The resulting supernatant S~ is layered onto 1.2 M sucrose (8 ml of S~ onto 4 ml) and centrifuged at 200,000 g at 4~ for 15 min. The resulting pellet is composed mainly of mitochondria, while synaptosomes, myelin, and some microsomes are retained at the gradient interface. This material is collected, diluted threefold with 0.32 M sucrose, layered onto 0.8 M sucrose (7 ml layered onto 4 ml), and centrifuged under the same conditions as the previous gradient spin. The resulting synaptosomal pellet is washed to remove the sucrose and resuspended in Krebs-Ringer bicarbonate (pH 7.3) at 2 mg of protein/ml for peptide degradation studies (16) or used to prepare membranes. Preparation of synaptosomes from postmortem human brain is best carried

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287

out on fresh tissue. However, frozen tissue can be used when it is first immersed in isotonic sucrose (cryoprotective agent) prior to slow freezing and storage at -70~ and then thawed rapidly in warm isotonic sucrose before use (17).

Preparation of Crude Synaptosomal Membranes Synaptosomes are lysed by resuspension in 5 mM Tris-HC1, pH 8.0, at 4~ for 1 hr. The suspension is centrifuged at 200,000 g at 4~ for 15 min. The resulting pellet is rehomogenized in l0 mM Tris-HC1 (pH 7.4)-0.2 M NaC1 and centrifuged as above (twice). The pellet is then rehomogenized in 10 mM Tris-HC1 (pH 7.4) and centrifuged as above (twice). The pellet is then resuspended (at ~5 mg/ml protein) in assay buffer for use immediately or stored frozen at -70~ for up to 1 month.

Identification of Peptidase Activity Peptide is incubated with tissue fraction and the products analyzed by HPLC. From the degradation products we can determine whether any of the primary sites of cleavage in the peptide correspond to the bond cleaved in the protein. A fluorimetric assay using Z-Gln-Lys-Leu-AMC is used to characterize LysLeu-cleaving activity (APP secretase).

HPLC Analysis of Peptide Degradation Peptides (0.5-mg aliquots) are stored freeze dried at -40~ in 1.5-ml Eppendorf tubes and dissolved in water (0.5 ml) immediately before use. Some peptides require acid (e.g., 0.01 M acetic acid), base [e.g., 0.1% (v/v) NH4OH], or organic solvent [e.g., 10% (v/v) dimethyl sulfoxide] to promote solubility. Peptide (100/zg), tissue fraction (100/zl; 5-10 mg/ml protein), and 0.5 M Tris-HC1, pH 7.4 (100/xl) in a total volume of 1 ml are incubated at 37~ with shaking. Alternative buffers are used as appropriate (e.g., lower pH for lysosomal peptidases). Bestatin (10 txM) and guanidinoethylmercaptosuccinic acid (GEMSA; 100/zM) are added to inhibit exopeptidases. Peptide and tissue blanks are included. Aliquots are removed at 10, 30, 60, 120, 240, and 300 min, acidified [100/xl of 1% (v/v) TFA] and centrifuged (15,000 g, 3 min, 4~ Supernatant (180/xl) is injected onto a NovaPak C~8 cartridge fitted with a guard column (Waters) as before, eluting with a 2.1-49% (v/v) acetonitrile gradient (20 min, 1 ml/min) containing 11 mM TFA. Peaks are detected at 214 nm. The tissue concentration may need to be adjusted if the peptide is degraded too rapidly or slowly. However, too concentrated a tissue fraction can give peaks that may interfere with the HPLC analysis;

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tyrosine, phenylalanine, and tryptophan are usually the main tissue-derived peaks. To identify peaks, incubations are repeated, choosing times to maximize the yield of degradation products, and subjected to HPLC as before. Fractions containing the peaks are collected by hand, evaporated to dryness, and subjected to amino acid analysis. We use precolumn derivatization with phenyl isothiocyanate and reversed-phase HPLC (Waters Pico Tag method). Amino acid composition is usually sufficient to identify the products; peptide sequencing is used to resolve ambiguities.

Fluorimetric Assay for Peptidase Activity The peptidase with the specificity to cleave the Lys-Leu bond was determined using Z-Gln-Lys-Leu-AMC as substrate in a two-stage assay. Substrate [7.5 /~1 of 2.5 mM stock solution in dimethyl sulfoxide (DMSO)] is incubated at 37~ with tissue extract (10-50/zl) in 50 mM Tris-HC1 (pH 7.4) (total volume, 150 ~1). After 1-5 hr, leucine aminopeptidase (LAP; porcine kidney type ClllCP, 0.4 units; Sigma, St. Louis, MO) and 25 mM MnC12 (40/zl) were added and the incubation continued for 2 hr to convert Leu-AMC to AMC. Ethanol (750/xl) was added and fluorescence measured (h~x 375 nm; hem445 nm). Blanks, where LAP was omitted, were taken through the whole procedure.

Preliminary Characterization of Peptidase Activity Mechanistic Class and pH Optimum Peptidases can usually be assigned to one of four mechanistic classes according to the chemical group mediating the catalysis and based on their sensitivity to inhibitors (Table I), although a few peptidases cannot be classified by this scheme (10). Tissue extract (10/zl) is preincubated for 15 min at 37~ with inhibitor (10/~1 of aqueous solution or 1/zl of DMSO solution) in appropriate buffer (total volume, 90 tzl). Peptide substrate (10 ~g in 10 txl) is added and the incubation continued. Ideally around 30-50% of peptide should be degraded in the absence of inhibitor. Trifluoroacetic acid (100/xl; 1%, v/v) is added to stop the reaction and the products analyzed by reversedphase HPLC as before. A rough estimate of the pH optimum for the peptidase activity is obtained by substituting appropriate buffers in the above scheme. The expected pH optima in Table I are only a rough guide. Stability of the activity at different pH values is determined by preincubating the peptidase at various pH values and then determining the activity remaining at the optimum pH.

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[16] IDENTIFICATION OF CNS PEPTIDASES TABLE I

Peptidase Inhibitors to Determine Mechanistic Class

Inhibitor

Concentration

Stock solutiona

3,4-Dichloroisocoumarin PMSF b

0.1 mM 1 mM

10 mM (DMSO) c 100 mM (DMSO) ~

E-64

0.1 mM

Class

Expected pH optimum

Serine

7-9

1 mM (water)

Cysteine

5-7

1,10-Phenanthrolined EDTA

1 mM 2 mM

100 mM (DMSO) 20 mM (water)

Metallo

7-8

Pepstatin

2/zM

200/xM (DMSO)

Aspartic

2-5

Stable at -20~ for months. b Phenylmethylsulfonylfluoride also inhibits some cysteine peptidases (reversible with 10 mM DTT). c Must be kept moisture free. d 1,10-Phenanthrolinegives a large peak on reversed-phaseHPLC, which can interfere with product identification. a

Other Inhibitors and Activators A large number of peptidase inhibitors of varying degrees of specificity are k n o w n a n d s o m e c o m m e r c i a l l y a v a i l a b l e o n e s a r e l i s t e d in T a b l e II. T h e e f f e c t o f p o t e n t i a l a c t i v a t o r s o n p e p t i d a s e a c t i v i t y (e.g., C a 2+ a n d o t h e r m e t a l i o n s , d i t h i o t h r e i t o l , A T P ) is a l s o d e t e r m i n e d . A c o m p r e h e n s i v e i n h i b i t o r a c t i v a t o r profile o f t h e a c t i v i t y is u s e d f o r c o m p a r i s o n w i t h t h e p r o p e r t i e s of known peptidases.

TABLE II Peptidase Serine Serine/ cysteine Cysteine Metallo Amino Carboxy

Commercially Available Inhibitors a Inhibitors Proteins b (al-antichymotrypsin, al-antiplasmin, antithrombin III, al-antitrypsin, aprotinin, soybean and lima bean trypsin inhibitor) Antipain, leupeptin (trypsin-like); chymostatin (chymotypsin-like); elastatinal (all 0.1 rnM) Iodoacetamide (0.1 mM), HMPS (0.1 mM); cystatin b Phosphoramidon, thiorphan (EP 24.11); captopril (angiotensin-converting enzyme); CPP-AIa-Ala-Phe-pAB (EP 24.15) Bestatin, amastatin GEMSA (0.1 mM)

Inhibitor concentration is 10/xMunless stated otherwise. Source of inhibitors: SigmaChemical Co. (Poole, Dorset, UK), Peninsula Laboratories Europe (Merseyside, UK), Calbiochem-Novabiochem(Nottingham, UK). HMPS, p-hydroxymercuriphenylsulphonate. b Protein inhibitors are active at equimolar concentrations with enzyme.

a

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Ac-E V H H Q K L V

,,....

E c

E D V G S

13

<

,r-

F

1

04 w Q 0 C

F

.13 L 0

1 3

.<

91o

7

--q 0

9

8

I

i

i

5

10

15

1

3

>

-

14

F

_..

2

~-

_

I

20

25

Elution time {min)

FIG. 1 HPLC analysis of degradation products from incubation of Ac-/3-AP (11-26) with human brain synaptic membranes. The numbered HPLC peaks correspond to the peptide sequence shown by the numbered arrows (fight). From the time course, peaks 7, 1l, and 13 [peptides(10-16), (1-9), and (1-11), respectively] were the earliest products detected. The proposed primary cleavage points are shown by large arrowheads; secondary (or minor) cleavage points by small arrowheads. Single-letter abbreviations: A, Ala; D, Asp; E, Glu; F, Phe; G, Gly; H, His; K, Lys; L, Leu; Q, Gln; S, Ser; V, Val. P, Parent peptide.

Applications 1. Using Ac-Glu-Val-Lys-Met-Asp-Ala-Glu-Phe-NH2, a human brain peptidase was identified that cleaves at the target site, Met-Asp. The ethylenediaminetetraacetic acid (EDTA) sensitivity of this activity showed that it was a metallopeptidase. Furthermore, inhibition by CPP-Ala-Ala-Phe-pAB indicated that endopeptidase 24.15 (EC 3.4.24.15, thimet oligopeptidase) could be responsible and this was confirmed after purification of the peptidase had allowed a more detailed characterization (2). The corresponding peptide (Ac-Glu-Val-Asn-Leu-Asp-Ala-Glu-Phe-NH2), having the two amino acid substitutions (Asn and Leu) associated with early deposition of/3-AP in a hereditary form of Alzheimer's disease, was cleaved at the same position (Leu-Asp) by this enzyme, and at three times the rate. 2. The second example illustrates a potential problem with using peptide substrates to identify bond-specific peptidases, namely the presence in the tissue extract of peptidases directed against other sites in the substrate. The peptide Ac-/3-AP (11-26) (Fig. 1) was originally used to search for the peptidase that cleaves Lys-Leu in APP (so-called a-secretase), but there were no products corresponding to cleavage at this site. The primary sites

[16] IDENTIFICATION OF CNS PEPTIDASES

291

with membrane preparations and intact cells were Phe-Phe and Ala-Glu. The same bonds were cleaved in the overlapping peptide Ac-fl-AP (16-29). Although not relevant to a-secretase activity, these peptidase activities are of great interest because they have the potential to degrade fl-AP itself, and may be involved in controlling extracellular concentrations of this protein. Analysis of the effect of inhibitors on the products of degradation showed that (a) peptide(1-11) formation was totally inhibited by 3,4-dichloroisocoumarin (3,4-DCI) but not by EDTA, E-64, or pepstatin; (b) formation of peptides(I-9) and -(10-16) was inhibited by 2 mM EDTA. Thus, a serine peptidase cleaves at Ala-Glu and a metallopeptidase cleaves at Phe-Phe. 3. Activity cleaving at Lys-Leu in Z-Gln-Lys-Leu-pNA was detected in human brain using HPLC analysis of products (1). However, the main activity was directed at the Leu-pNA bond. Formation of Z-Gln-Lys and Leu-pNA was inhibited only by EDTA, showing that a metallopeptidase was responsible. The main activity cleaving at Leu-pNA was inhibited by both chymostatin and HMPS, and was later shown to be multicatalytic endopeptidase (MCP). This demonstrates a hazard in using cleavage of the arylamide bond in small fluorogenic or chromogenic substrates to identify peptidases directed at specific peptide bonds. Although MCP cleaves the Leu-pNA bond in ZGln-Lys-Leu-pNA, it does not cleave the equivalent bond (Leu-Val) in the peptide substrate, Ac-fl-Ap (! 1-26). Because of the competing MCP activity, further characterization of the Lys-Leu cleaving activity was carried out after chromatographic separation of the two peptidases.

High-Performance Ion-Exchange and Size-Exclusion Chromatography Prior to a large-scale purification of the peptidase, preliminary characterization is continued by resolving multiple peptidase activities in the crude extracts, using high-performance ion-exchange chromatography (HPIEC). Endogenous inhibitors may also be resolved from the peptidase by this procedure. Soluble tissue extract (5-10 ml; in l0 mM Tris-HCl, pH 7.4) is injected onto a glass Protein Pak DEAE 5PW column (Waters; 8 x 75 mm), connected to a Waters 650 protein purification system, eluting at 0.5 ml/min with a 60-min gradient from 0.01 to 0.5 M Tris-HC1 (pH 7.4), and fractions are collected (0.5 ml). Membrane-bound activity is first solubilized with nonionic detergent [1% (w/v) Triton X-100 in 10 mM Tris-HC1, pH 7.4; 60 min; 4~ centrifuged, and the efficiency of solubilization checked by assaying peptidase in the supernatant. Triton X-100 (0.1%, w/v) is then included in the elution buffers. Fractions are analyzed for peptidase activity against peptide substrate, using automated reversed-phase HPLC. The elution condi-

292

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3

z

0.6

?

g

N

,

~

~

0.2 0.2

0

20

40

Fraction No.

60

0

10

20

30

40

50

Fraction No.

FIG. 2 High-performance anion-exchange chromatography (TSK-DEAE 5PW; left) of human brain soluble fraction. Peptidase activity (O) was detected by HPLC analysis using Z-Gly-Lys-Leu-pNA as substrate. Peak A (the Lys-Leu hydrolyzing activity) was subjected to size-exclusion chromatography (Superose 6; fight); the HPLC assay (0) and the two-stage fluorimetric assay with Z-Gln-Lys-Leu-AMC ( 9 detect the same peak.

tions are optimized to allow as short a run time as possible. Peptide (10/~g) is incubated with l0/A of fraction in appropriate buffer (total volume, 50/A) for 15-30 min, the reaction being stopped with 150/~1 of 1% (v/v) TFA. Because the HPLC assay is still relatively time consuming, the approximate location of the activity is first determined by analyzing pooled samples (10 /~1 each from groups of five tubes). Individual tubes are then analyzed in areas of activity. Figure 2 shows the resolution of Lys-Leu hydrolyzing activity (peak A, Fig. 2) from Leu-pNA-hydrolyzing activity (peak B, Fig. 2). Peak A (Fig. 2) gave only Z-Gln-Lys and Leu-pNA on incubation with Z-Gln-Lys-LeupNA, while peak B (Fig. 2) gave only Z-Gln-Lys-Leu and pNA. Pooled fractions from peak A (Fig. 2) were concentrated to 0.5 ml, using a centrifugal miniconcentrator (Centricon 10; Amicon, Danvers, MA) and applied to a size-exclusion column (1 • 60 cm, Superose 6; Pharmacia, Piscataway, NJ) eluting with 50 mM Tris-HCl, pH 7.4 (0.5 ml/min) (Fig. 2). Fractions were assayed for activity by the coupled fluorimetric assay. By comparing the elution time with that of known molecular mass standards, the molecular mass of the peptidase was estimated at 119 kDa. The peptidase activity was not pure at this stage but could be used for further characterization. It was shown that the metallopeptidase had an optimum pH in the range of pH

293

[16] IDENTIFICATION OF CNS PEPTIDASES

TABLEIII Purification of Peptidase Activity a ,,

Step 1. Tissue extraction

2. Ammonium sulfate precipitation 3. Anion-exchange chromatography 4. High-performance anion-exchange chromatography 5. High-performance hydrophobic interaction chromatography 6. Size-exclusion chromatography

Procedure Human brain (500 g): homogenized in 2.5 liters of buffer A, centrifuged (27,000 g; 15 min; 4~ and the supernatant removed Fraction precipitating between 25 and 50% saturation. Dissolve pellet in buffer A. Dialyze against buffer A DEAE-cellulose (3 x 30 cm). Wash with buffer A (2 column volumes). Elute with a 500-ml gradient to 0.5 M Tris-HC1 (pH 7.4) TSK-DEAE 5PW (8 x 75 mm). Elute with a 60-min gradient of buffer A to 0.5 M Tris-HC1 (pH 7.4), 1 mM EDTA; 0.5 ml/min TSK-phenyl 5PW (8 x 75 mm). Elute with a 60-min gradient of 1.8 to 0 M (NH4)2SO 4 in 10 mM Tris-HCl (pH 7.4), 1 mM EDTA Superose 6 (1 x 60 cm) or TSK G3000SW (7.5 x 60 cm). Elute with buffer A, 0.5 ml/min

a Assay method" Fluorimetric" Z-Gln-Lys-Leu-AMC~ Leu-AMC ~ AMC; buffer A, 50 mM Tris-HC1 (pH 7.4), 1 mM EDTA.

6.8-8 and that it was stimulated by 0.1 mM Mn 2+ (1). It had proteolytic activity against casein and, from its inhibitor profile, it appeared to be a novel peptidase.

Purification of Peptidase Activity While partial purification as outlined above is often sufficient to determine whether the activity is due to a known peptidase, a greater degree of purification is required to determine specificity, kinetic parameters, possible subunit composition, and sequence of the peptidase. For a comprehensive guide to protein purification procedures the reader is referred elsewhere (18). Buffers used in purification should contain stabilizers when appropriate. Cysteine peptidases are often protected by dithiothreitol and addition of glycerol to buffers may help stabilize some peptidases. EDTA is used in the buffers for purification of the Lys-Leu hydrolyzing peptidase to avoid inactivation by heavy metal ions; the activity is restored by adding Mn 2+ in the assay. A summary of the scheme used to purify the Lys-Leu hydrolyzing activity using Z-Gln-Lys-Leu-AMC as substrate is given (Table III). Ammonium sulfate precipitation or batch adsorption onto ion-exchange resins are

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often good initial steps to both purify and concentrate the peptidase from large volumes of tissue extract. The purification is followed by SDS-PAGE at each stage to determine the degree of purity. While there is little that is unique to CNS peptidases in the application of protein purification techniques, two aspects require some consideration: assay method and starting material. As noted earlier, HPLC is not convenient for the assay of large numbers of column fractions, although we have demonstrated that it is possible to use this method to purify a human brain peptidase (2). While it may be necessary to use HPLC analysis of the peptide substrate to follow early steps of the purification, the characteristics of the partially purified enzyme may allow more convenient assays for the later steps. For example, the Lys-Leu hydrolyzing activity has general protease activity against casein, is inactive in the presence of EDTA, but is reactivated with 0.1 mM Mn 2+. A simple and quick spectrometric assay can be constructed to measure Mn2+-stimulated degradation of azocasein. Second, the tissue or cell fraction in which the peptidase activity was identified may not be the most appropriate starting material for a large-scale purification. Subcellular fractions may be used in the initial search for the activity but a crude tissue extract (soluble or particulate) is often suitable for large-scale purification. It may also be worth assaying tissues outside the CNS for the activity in case it is present elsewhere in significantly higher concentration, which might facilitate purification.

Summary Synthetic peptides from/3-amyloid precursor protein were used as substrates to identify and characterize four peptidase activities in human brain, as follows: 1. A novel Mn 2+-stimulated metallopeptidase with the specificity to cleave at the Lys-Leu secretase site in APP (1). 2. Peptidase with the specificity to generate the N-terminus of/3-AP from APP by cleaving a Met-Asp (or Leu-Asp) bond: characterization showed this activity to be endopeptidase 24.15 (2). 3. Two membrane peptidases with the specificity to cleave Phel9-Phe 2~ and A l a Z l - G l u 22 in/3-AP; the former metal dependent and the latter serine dependent.

References 1. J. R. McDermott and A. M. Gibson, Biochem. Biophys. Res. Commun. 179, 1148 (1991).

[16] IDENTIFICATIONOF CNS PEPTIDASES .

.

10. 11.

12. 13. 14. 15. 16. 17. 18.

295

J. R. McDermott, J. A. Biggins, and A. M. Gibson, Biochem. Biophys. Res. Commun. 185, 746 (1992). J. R. McDermott, A. M. Gibson, A. E. Oakley, and J. A. Biggins, J. Neurochem. 56, 1509 (1991). J. R. McDermott, A. M. Gibson, and J. A. Biggins, Biochem. Soc. Trans. 16, 851 (1988). J. R. McDermott, D. Mantle, B. Lauffart, and A. M. Kidd, J. Neurochem. 45, 752 (1985). J. R. Backstrom, C. A. Miller, and Z. A. Tfk6s, J. Neurochem. 58, 983 (1992). G. Sarath, R. S. De La Motte, and F. W. Wagner, in "Proteolytic Enzymes: A Practical Approach" (R. J. Benyon and J. S. Bond, eds.), p. 25. IRL Press, Oxford, 1989. R. B. Nelson and R. Siman, J. Biol. Chem. 265, 3836 (1990). C. R. Abraham, J. Driscoll, H. Potter, W. E. Van Nostrand, and P. Tempst, Biochem. Biophys. Res. Commun. 174, 790 (1991). N. D. Rawlings and A. J. Barrett, Biochem. J. 290, 205 (1993). J. Kang, H. G. Lemaire, A. Unterbeck, J. M. Salbaum, C. L. Masters, K. H. Grzeschik, G. Multhaup, K. Beyreuther, and B. M011er-Hill, Nature (London) 325, 733 (1987). D. H. Small, G. Reed, S. J. Fuller, A. Weidemann, K. Beyreuther, and C. L. Masters, this volume [18]. C. G. Knight, Biochem. J. 274, 45 (1991). B. Storrie and E. A. Madden, in "Methods in Enzymology" (M. Deutscher, ed.), Vol. 182, p. 203. Academic Press, San Diego, 1990. P. R. Dodd, J. A. Hardy, A. E. Oakley, J. A. Edwardson, E. K. Perry, and J. P. Delaunoy, Brain Res. 226, 107 (1981). J. R. McDermott, A. I. Smith, P. R. Dodd, J. A. Hardy, and J. A. Edwardson, Peptides (N.Y.) 4, 25 (1983). J. A. Hardy, P. R. Dodd, A. E. Oakley, R. H. Perry, J. A. Edwardson, and A. M. Kidd, J. Neurochem. 40, 608 (1983). M. P. Deutscher, ed., "Methods in Enzymology," Vol. 182. Academic Press, San Diego, 1990.

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Strategies for Characterizing, Cloning, and Expressing Soluble Endopeptidases M a r c J. G l u c k s m a n a n d J a m e s L. R o b e r t s

Introduction Endopeptidases are peptidolytic enzymes that cleave small proteins (peptides) at internal peptide bonds and thus are extremely important in neuropeptide metabolism, both synthetic and degradative. For example, while the synaptic action of neurotransmitters is terminated to a large extent by specific high-affinity uptake systems, most neuropeptides have their extracellular action terminated by peptidases. Alternatively, peptidases can also be involved in the synthesis or activation of neuropeptides, either intracellularly as part of the vesicular secretory pathway, or extracellularly by modifying a previously secreted peptide. Thus, peptidases play a central role in neuropeptide action in the nervous system.

Peptidase Characteristics Proteases/peptidases are divided into four classes based on signature elements of their catalytic site: serine, cysteine, aspartic, and zinc metalloproteinases. Of importance to this treatise is that these enzymes can also exist in either a membrane-bound or soluble form. Endopeptidases involved in the intracellular biosynthesis of neuropeptides are sequestered within the classic endoplasmic reticulum/Golgi/secretory vesicle pathway with their substrates, the neuropeptide precursors; an example of membrane-bound enzymes would include the prohormone convertases, or PC enzymes. Another group of membrane-bound endopeptidases includes those involved in extracellular neuropeptide processing; these are synthesized through the vesicular pathway with one or more transmembrane segments anchoring the enzymes to the extracellular surface. Examples of this type of membrane endopeptidase are neutral endopeptidase (enkephalinase) and angiotensinconverting enzyme. The soluble endopeptidases are found in the cytoplasmic subcellular compartment and, by mechanisms not clearly understood, are secreted into the extracellular space, where they act on released neuropeptides. The major advantage in working with these endopeptidase proteins is 296

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their solubility, allowing for simple purification schemes and easy manipulation into and by prokaryotic systems.

Endopeptidase 24.15 as an Example While a soluble endopeptidase (EP) may belong to any of the four classes of proteases, we will illustrate our cloning strategies by considering EC 3.4.24.15 (EP 24.15, thimet oligopeptidase, endo A), a member of the zinc metalloproteinases. This enzyme is a soluble, nonglycosylated, highly abundant, 77,000-D endopeptidase present primarily in brain, pituitary, and testis (1). Endopeptidase 24.15 cleaves bonds on the carboxyl side of specific hydrophobic residues present in peptides less than 25 amino acids in length. The presence of another hydrophobic residue in the third amino acid position carboxyl to the peptide bond to be cleaved increases the substrate affinity of the enzyme (2). This enzyme has been isolated (2), and the full-length rat cDNA has been sequenced, used to direct the expression of protein, and enzymatically characterized (3). Endopeptidase 24.15 exists in two forms in both tissues and cell lines: a predominant soluble form constituting about 80% of the total activity and minor forms bound to the membrane and nucleus accounting for the remainder of the activity (4, 5). The two forms of EP 24.15 are similar if not identical, with respect to substrate specificity, immunological properties, and sensitivity to specific inhibitors. Initially, it was thought that the two forms might differ by the presence of a membrane-spanning segment in the membrane-bound form, as seen in several zinc-containing peptidases, such as neutral endopeptidase (6), or in their posttranslational modifications, which might segregate the enzyme into different compartments of the cell. However, one of the outcomes of the cloning and sequencing of the EP 24.15 cDNA was that the encoded enzyme was shown to lack a membrane-spanning segment, suggesting that the latter possibility for membrane association is most likely.

Extended Family of Endopeptidase 24.15 To identify and characterize novel enzymes related to EP 24.15 and family members from different species and tissues, we need to detect cognate proteins reported in the literature to build a database on these soluble endopeptidases. Several putative peptidases (which may be EP 24.15 or closely related

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by sequence comparisons) have been described on the basis of substrate cleavage to metabolize gonadotropin-releasing hormone, neurotensin, and other bioactive peptides for rat, rabbit, and human (5, 7-10). Slight differences have been noted with respect to inhibitor profiles, but these may be issues of purity. Protein purity is an issue when attempting to distinguish enzyme activity in the presence of substrates and inhibitors. Prior to molecular cloning, the characterization of these peptidase activities has been performed on semipurified or crude tissue preparations, which often contain more than one enzymatic activity. The development of an assay system including specific substrates and inhibitors is crucial for delineating the biochemical properties of an enzyme. Additionally, there is a need for the demonstration of comparable affinities and kinetics toward natural substrates of the metalloendopeptidase in the purified tissue form and in the recombinant protein (if there are no substantial posttranslational modifications; the case in the soluble endopeptidases) for further studies with the cloned form. Once cloned, and then expressed, the activity of the enzyme must be characterized compared to its purified activity. Phylogenetically, these metalloendopeptidases contain conserved sequences that reflect the mechanism of catalysis and elements of structural motifs (11, 12). Endopeptidase 24.15 enzyme activity has been characterized, by inhibitor and substrate studies, to be present in species from human to Xenopus laeois, Aplysia californica, and yeast, although its endogenous substrate in many species have yet to be discovered. A pig liver angiotensin II-binding protein with considerable homology to EP 24.15 has been cloned, sequenced (13), and shown to have properties almost identical to EP 24.15. On the level of the amino acid sequence, rat EP 24.15 has a 61% homology with rabbit liver microsomal endopeptidase (14) and 62% with the pig liver angiotensin II-binding protein (13), whereas the rabbit and pig proteins share 92% identity in sequence. This demonstrates that there are enzymes related to EP 24.15 that add to the information for cloning other soluble endopeptidases (see Table I). An EP 24.15-1ike enzyme was hypothesized to be encoded in the open reading frame of YCL57w, present in yeast chromosome III (15), as discovered by searching a "translated" nucleic acid database instead of a protein database for similarities where this would not have been detected, and subsequently cloned (16). Utilizing this information, a protein was subsequently isolated and a partial sequence (20 of 20 amino acids were identical) matching the molecular mass of the putative Saccharomyces cereoisiae EP 24.15 was discovered (16). This yeast enzyme was demonstrated to cleave bradykinin, fl-neoendorphin, and neurotensin as does the rat and human EP 24.15. Other

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C L O N I N G A N D E X P R E S S I O N OF E N D O P E P T I D A S E S TABLE I

Enzymes Functionally Equivalent or Related to Endopeptidase 24.15 Enzyme

Ref. a

Endo-oligopeptidase A (Endo A) (E) b Pz-peptidase (E) Thimet oligopeptidase (E) Rat mitochondrial intermediate peptidase Pig soluble angiotensin-binding protein Rabbit liver microsomal endopeptidase Yeast metalloendopeptidase (E)

1 2 3 4 5 6 7

a Key to references: (1) A. C. M. Camargo, H. Caldo, and M. L. Reis, J. Biol. Chem. 254, 5304 (1979); (2) A. J. Barrett and U. Tisljar, Biochem. J. 261, 1047 (1989); (3) A. J. Barrett and N. D. Rawlings, Biol. Chem. Hoppe-Seyler 373, 353 (1992); (4) G. Isaya, F. Kalousek, and L. E. Rosenberg, Proc. Natl. Acad. Sci. U.S.A. 89, 8317 (1992); (5) N. Sugira, H. Hagiwara, and S. Hirose, J. Biol. Chem. 267, 18067 (1992); (6) S. Kawabata, K. Nakagawa, S. Iwanaga, and E. W. Davie, J. Biol. Chem. 268, 12498 (1993); (7) C. A. Hrycyna and S. Clarke, Biochemistry 32, 11293 (1993). b E, Equivalent.

enzymes, including a rat mitochondrial intermediate peptidase (17) and a rabbit liver microsomal endopeptidase (14), have sequences similar to that of EP 24.15. Thus, it is likely that a family of soluble endopeptidases is present for peptide metabolism.

Strategies for Cloning For the purposes of this chapter, EP 24.15 is used as a prototype, because the strategy involved in its cloning and expression is general enough to be applicable to novel and related enzymes. However, this chapter does not attempt to review all of the possible protocols involved in obtaining recombinant DNA clones for the soluble metalloendopeptidases. A few specific instances in which a proven method has been demonstrated to be useful will illustrate the manner in which that approach can assist in the elucidation of specific characteristics regarding a specific metalloendopeptidase.

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First Considerations Four strategies are discussed that are dependent on information available to the researcher: (a) purifying the enzyme and obtaining a portion of the amino acid sequence, (b) obtaining specific antisera to the protein of interest, (c) cloning by homology to a nucleic acid sequence that is known, and (d) analyzing secondary structural characteristics. One of the primary considerations before embarking on a project is that of the tissue to be utilized as starting material. While one may be interested in a proteolytic activity in brain, the same enzymatic activity may be present in another tissue at a much greater concentration, dramatically aiding in enzyme purification. Indeed, because enzyme activity in the form of a crude tissue homogenate is a fair approximation of enzyme concentration, that tissue probably contains a higher concentration of the enzyme encoding mRNA for obtaining clones during a subsequent library screening. Endopeptidase 24.15 enzyme activity is 5-fold greater in testis than in brain, and the RNA is 35-fold more abundant in testis (5). This same principle can be further applied to subdivide specific areas of the brain wherein the message is more plentiful, such as the hippocampus or hypothalamus. Another choice lies between creating a library from RNA isolated in the laboratory and purchasing a commercially prepared library. To make a library from a specific source, any standard manual (18) can suggest a method for isolating RNA from tissue. Once pure RNA is obtained, a high-fidelity RNase H- reverse transcriptase such as Superscript (GIBCOBethesda Research Laboratories, Gaithersburg, MD) should be utilized to perform a first- and second-strand synthesis; adapters should be added on each end and the DNA packaged into an appropriate vector. The ZAP vectors and their derivatives, which can be made to express the inserts as well as form an excisable phagemid at the end of the procedure (Stratagene, La Jolla, CA), are a common choice. The excisable phagemid is an important consideration because a vexing problem in a cloning program is the subcloning of the cDNA out of the phage vector. If a commercially available library is purchased [major suppliers are Stratagene and Clontech (Palo Alto, CA)], there are several safeguards that should be followed. Note the number and sex of the specimens used as the starting material and the method for the isolation and treatment of the RNA if this is relevant. The method by which the library was prepared and the average size of the insert will determine the number of individual phage that will have to be screened for a statistical sampling of three to five genome equivalents. It may be optimal to have the RNA treated with mercury methyl hydroxide for longer stretches of 5' sequence by releasing

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RNA secondary structure and to attain the largest average cDNA size. Additionally, it is useful to employ oligo(dT) and random priming prior to the second-strand synthesis to assure a true representation of all of the mRNAs in creating a library.

Purifying the Enzyme and Obtaining a Portion of the Amino Acid Sequence In the absence of nucleic acid sequence data, but having the ability to obtain some highly purified protein by conventional biochemical techniques, one can begin a cloning project. A well-resolved band from electrophoresis on a polyacrylamide gel that is then transferred to polyvinylidene difluoride (PVDF) nitrocellulose can be microsequenced to yield amino acid sequence data. Fragments isolated by peptide mapping can, on the basis of homologous enzymes that are isolated, be proteolytically digested and examined for "fingerprints" identical to those obtained with a known protein. As an adjunct to nucleic acid sequencing it is crucial that the data are collected on peptide fragments obtained by protein sequencing, in addition to DNA sequencing of both strands. In the case of clones obtained from an expression library, an additional criterion can be the demonstration of enzymatic activity. Western immunoblotting may be misleading because there is a dependency on the antigenic determinants and the source of the antibody for immunoreactivity. Obtaining Specific Antisera to the Protein of Interest Screening a cDNA library can be done on the basis of the availability of antibodies to screen an expressed protein. If so, the choice of libraries must be made with cDNAs being expressed to synthesize their encoded patterns. In conjunction with enzyme assays, immunoblotting techniques (Western blots) are a quantitative way to assess the presence and amount of a protein antigen in order to aid in the choice of library source. This technique is described in detail elsewhere (18). When designing antibodies several considerations should be taken into account. First, polyclonal antibodies have greater utility than monoclonals, because there will be a greater number of epitopes expressed that can crossreact with related portions of homologous endopeptidases. Second, a choice should be made as to the state of the protein before presentation to the animal (usually rabbit) producing the antibodies. Either a native protein that is purified by conventional biochemical techniques can be used or a welldetermined band excised from a sodium dodecyl sulfate (SDS)-polyacrylamide gel can be pulverized and injected directly to produce antibodies. The

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original antibody to EP 24.15 was derived from a denatured antigen and proved useful for screening and expression library and subsequent quantitation by Western immunoblotting. Last, a choice must be made as to whether to use a fused or unfused enzyme as an antigen. Empirically, fused portions of proteins such as glutathione transferase or maltose-binding protein (described in the next section) hasten the time course and amplify the immune response to the enzyme of interest, which acts as a carrier, but may yield higher nonspecific backgrounds on detection. An application of this experimental approach utilizes polyclonal rabbit antiserum against EP 24.15 produced from the purified homogeneous enzyme. Native enzyme was eluted from a glutathione-Sepharose column, or EP 24.15 in a denatured form was obtained following preparative SDS slab gel electrophoresis and excision of the single band corresponding to the protein. Initial injections of 200/zg of protein were used for immunization, with three subsequent injections of 100/zg of EP 24.15 repeated at 2-week intervals. The enzyme was mixed with Freund's complete adjuvant and injected intradermally into New Zealand White rabbits. Blood was collected in heparinized tubes from the terminal ear vein 1 week after the injections. A high titer of the antibody was judged autocatalytically, by the presentation of the initial antigen. Antiserum was stored in 200-/zl aliquots at -20~ Western blots were performed that produced a strong signal with little background. The polyclonal antibody reacted with cloned and bacterially expressed enzyme, and with enzyme isolated from mammalian tissue. The original clones for EP 24.15 were obtained from a h ZAP expression library (3). Cloning by Homology to a Nucleic Acid Sequence That Is Known With the various peptidase enzyme activities described and sources of sequence information available, the goal is either to clone the homologous enzyme in a different species or to undertake the discovery of a novel enzyme activity in the family of soluble zinc metalloendopeptidases. Unfortunately, there is a paucity of structural information derived from X-ray crystallography concerning the mammalian endopeptidases, therefore other sources of data must be derived. It is often useful to define and search for patterns in these proteins (and nucleic acid sequences) in order to elucidate related or consensus motifs. Sequence comparisons and pattern matching have proved to be extremely effective in the study of the structural and functional properties of proteins. In this manner, the protein sequences of thermolysin, EP 24.15, angiotensin-converting enzyme, and neutral endopeptidase were compared using an alignment paradigm. FASTA, which allows a comparative search of one sequence against an entire sequence database (or a subset thereof) to screen all known metalloproteases (19). The algorithm BESTFIT in the GCG-Wisconsin package (20) was then employed, using a fixed anchor

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at one of the following functional domains: the active site (-H-E-x-x-H-), a carboxy-terminal four-helix bundle, or close invariant residues involved in the enzyme mechanism, searching for the highest amount of homology using several gap penalties and checking for consistency of alignments as reflected by the match score. Amino acids required for function and identified biochemically are additional patterns which can be recognized. The general spatial relationships between secondary structural elements remain highly conserved among related enzymes as well as between species. These defined hallmarks typical of metalloproteases are pursued in obtaining consensus features for the elucidation of novel related enzymes.

Novel Structure Analyses for Secondary Structural Characteristics A criterion to assess the relationship of a novel endopeptidase with previously described ones lies beyond weak homologies based solely on the primary sequence. There are motifs composed of secondary structure that are present in all of the metalloendopeptidases (12, 19), and this analysis can yield rigorous determinants to answer the question, "Is this a related soluble endopeptidase?" Furthermore, in designing inhibitors (one of the potential goals of the work) common structural elements have already been described (12, 19). The first step is to construct a compilation of alignments of related sequences of the soluble metaloendopeptidases to indicate the regions of greatest homology (evolutionary conservation), such as the active site H-Ex-x-H. A common approach to lcoating sites that are similar in a group of proteins is by searching for a consensus sequence that is common to all of the elements. There are other amino acid residues of biological interest reflecting conservation of the mechanism of substrate catalysis. With the algorithm MOTIFS, we have identified several signature protein sequences that are consensus sites for modification such as phosphorylation. Knowledge of these conserved amino acid residues may serve as the starting point for the design of synthetic oligonucleotides for screening cDNA libraries. Another approach was used to identify protein motif similarities that complemented standard methods. The best alignments obtained from the GCGWisconsin package were then used for hydrophobic cluster analysis (HCA) (21, 22). The invariance of secondary structural elements important spatially between the metalloendopeptidases has been demonstrated in comparisons of thermolysin (EC 3.4.24.27), Pseudomonas aeruginosa elastase (EC 3.4.24.26, pseudolysin), Bacillus cereus neutral protease (EC 3.4.24.28, bacillolysin), and EP 24.15 (12). The initial alignment of metalloprotease sequences by HCA begins at the active site and at the carboxy terminus. One can then compare domains by a matrix function and derive rudimentary

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evolutionary relationships between the proteins being assessed to compose a family. The sequences of EP 24.15, neutral endopeptidase, and angiotensin-converting enzyme, as well as the carboxy-terminal half of each enzyme alone, were also input to PROBE, a series of neural network programs that predicts secondary structure (23, 24). This neural network was of a tandem design to prevent a state of memorization that might compromise the results when compared to the basis set. The results, corroborating HCA analyses, indicated regions that were conserved with respect to secondary structural elements such as a four-helix bundle and an ~ helix at the active site as well as residues involved with substrate catalysis (19). These motifs must be preserved in this family of enzymes and go beyond the signature H-E-x-x-H sequence.

Library Screening Strategies Two methods are initially utilized for procuring an enzyme previously isolated and characterized from a new species or to ascertain members of a family of related enzymes. The methods are oligonucleotide screening of libraries with synthetic oligonucleotides or restriction endonuclease fragments from a clone previously obtained, and polymerase chain reaction (PCR) screening with oligonucleotides synthesized to a region homologous between other enzyme family members.

Oligonucleotide Screening of Libraries Oligonucleotide screening is illustrated by a description of our work on EP 24.15, in which human EP 24.15 cDNA clones are isolated and characterized. A human cDNA library is obtained that contains bacteriophage hgtl0 with an average insert size of 1.8 kb. This library is plated to allow three genome equivalents to be screened on duplicate filters. Rat EP 24.15 (3) is utilized as a probe to a stringency of, first, 1.25x SSC (1 x SSC: 150 mM NaC1, 23 mM sodium citrate) with 0.1% (w/v) SDS at 65~ and subsequently washed with 0.9x SSC with 0.1% (w/v) SDS at 65~ to produce low backgrounds while maintaining a tenable signal. An additional two rounds of screening with the full-length probe allow the initial putative positives to be isolated as single plaques. Southern blot analysis (25) searching for a full-length insert based on the rat sequence yields several clones of interest. A Southern blot is performed on isolates and individual phage DNA preparations are run in triplicate. Each of 11 positives from the library are digested with the restriction enzyme

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EcoRI to excise the human DNA inserts, electrophoresed on a 1% (w/v) gel, and after transfer to nitrocellulose is probed with a full-length 2.1-kb rat probe, a 5' probe containing a 210-base pair (bp) BamHI-BstEII restriction fragment, or a 3' probe containing a 265-bp KpnI-EcoRI restriction fragment. The full-length probe identifies all cDNAs homologous to EP 24.15 while only clones cross-hybridizing with both the 5' and 3'ends as well give a fulllength human EP 24.15. Polymerase Chain Reaction Polymerase chain reaction (PCR) is a technique common in the laboratory with proven published protocols (18). When designing an oligonucleotide probe, several steps must be taken. The degeneracy of the genetic code must be considered; all amino acids except methionine and tryptophan are encoded by two to six codons. Codon degeneracy involves only the third base in most cases, the exception being those amino acids encoded by six codons (arginine, leucine, and serine). Because of this feature, probe sequences have usually been based on a region of the protein sequence rich in amino acids responding to a restricted number of codons (particularly methionine and tryptophan), thereby limiting the method to detecting the presence of contiguous sequences with low codon degeneracy. However, in order to cover all codon possibilities, it is necessary to synthesize a set of alternative probes comprising from 4 to 256 different oligonucleotides. The caveat remains: the lower the degeneracy, the higher the specificity. Avoiding the use of amino acids with four or six codons is critical whenever possible. Primers or probes can then be designed adding sequences for restriction endonucleases on the 5' end to facilitate directional subcloning of the subsequent PCR products. These primers have an additional two 5' nucleotides to allow the restriction endonuclease to act under reasonable cutting efficiency. This approach has also been used to obtain clones for the human EP 24.15. Experimentally, pools of oligonucleotide primers representing all possible codon combinations for a small part of the given amino acid sequence are synthesized. Within this mixture must be one primer that has a sequence complementary to the DNA encoding the given amino acid, and will form a perfectly base-paired duplex with the DNA, while the other primers in the mixture will form mismatched duplexes under certain stringent hybridization conditions discriminatory against incorrectly base-paired primers. These mixtures are capable of locating specific gene sequences in the libraries. This can best be illustrated with the codons for the amino acids arginine, serine, and leucine. There are six codons for each of these amino acids and in the case of serine, a protein perfectly matched by the amino acid sequence may be misaligned on the nucleotide level. Two of the codons of serine are

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mismatched in all three sites (UCA vs AGU), not just in the third wobble position. Organismal or taxonomic bias in codon choice is a more complicated problem, but can be used to limit further the degeneracy of an oligonucleotide primer. For a given amino acid and a given organism there is a skewed chance of one triplet codon appearing more than the others. Using the example of glutamine, in yeast, CAA is preferred over CAG about three-quarters of the time, whereas in mammals the disposition is reversed (26). The GCGWisconsin computer package used to search the nucleic acid database contains an algorithm CODON PREFERENCE with information on codon usage by organismal criteria. The correct reading frame for a real gene may rise significantly above the background if a suitable codon frequency table is used; in addition this is a sensitive method for locating DNA sequencing errors. Polymerase chain reaction primers may contain as few as 15 to 21 nucleotides, which requires the identity of a conserved region to be 5 to 7 amino acids long. In some cases, PCR primers can be designed on as little as four amino acids by inclusion of six to nine 5' base extensions, including restriction endonuclease sites incorporated into the amplified DNA product, increasing stability of the priming duplex at all subsequent amplification cycles. This approach has been used successfully in identifying uncharacterized viruses by limited amino acid sequence similarity to the reverse transcriptases (27). It is vital to avoid a mismatch on the 3' end, because this would affect extension. Therefore, the sense primer should avoid a degenerate terminal nucleotide and end (if possible) with a G or C nucleotide. One microgram of a cDNA library, as the template containing the thermostable polymerase buffer, 100 pmol of each degenerate primer, and a 200 ~M concentration of each dNTP are added to a total volume of 50 ~1. After an equal volume of mineral oil is added, the sample is first placed in a thermocycler at 100~ for 10 min to denature all duplexes, then 1 unit of thermostable polymerase is added after ramping down the temperature to 80~ for 6 min. Following enzyme addition, the temperature is programmed for denaturation of 94~ for 1 min, then dropped to 55~ for 1.5 min for annealing and extending at 72~ for 1 min. After 30 cycles, 10 ~1 of the reaction is electrophoresed through a 1.2% (w/v) agarose gel and visualized with ethidium bromide. If appropriately sized bands are present, the remainder of the PCR reaction is then extracted with a one-half volume of phenol, chloroform extracted, precipitated with ethanol, resuspended in buffer, and digested for 5 hr with the appropriate restriction enzyme buffer. Electrophoresis as above follows. The PCR products are excised from the agarose gel and extracted using Mermaid or Geneclean (Biol01, La Jolla, CA), depending on the size of the oligonucleotide fragment. If low-melt agarose is employed, the PCR product

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can be extracted after agarose digestion by/3-agarase (New England BioLabs, Beverly, MA). The DNA fragments are then ligated into an appropriate vector with the use of previously added restriction site oligonucleotide linkers to the primers, adding an extension of two bases on the 5' ends. Alternatively, one can ligate the PCR products directly into a vector without purification or restriction digests, following the ligation with screening and excision (Invitrogen, San Diego, CA). This technique is based on the empirical observation that there is a one-base extension created by Taq polymerase. The products of these reactions are then ligated and transformed into bacteria and plated onto drug-resistant plates. Because inosine is a neutral base, it can pair with any of the four nucleotide bases in the sequence; in some cases the oligonucleotides were synthesized as a mix of inosine-replaced nucleotides. Deoxyinosine can be inserted in oligonucleotide probes at the ambiguous codon positions as determined by analysis of the peptide sequence, obviating the need to synthesize highly degenerate primers (28). This may be a significant improvement over the technique of utilizing mixed probes to compensate for the degeneracy of the genetic code.

Strategies for Protein Expression

Expression Systems for Enzyme Study Protein expression, in the context used here, refers to the particular synthesis of copious quantities of a specific protein directed by a designated cell. Historically, the genetic manipulation through recombination events and mutations of regulatory proteins in prokaryotes used terms such as overexpression or overproduction to describe alterations in gene expression that resulted in a few select proteins of interest. The knowledge gained as a result of the fluency in using recombinant DNA technology has permitted the engineering of vectors with promoters that direct large amounts of synthesis of mRNA corresponding to the gene of interest, drug selectivity, and sequences to aid in purification, and host bacteria or cells to allow high-yield synthesis of a particular gene product. In fact, there has been a paradigm shift from the genetic reconstruction of the vector carrying the gene to the choice and development of the host organisms, from bacteria to lactating cows. Whether intended for extensive biochemical and enzymatic characterization, or biophysical studies to provide three-dimensional information at atomic resolution [such as nuclear magnetic resonance (NMR) or X-ray

308

Ill

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS TABLE II

Applications Requiring Purified Enzymes

Application/technique

Amount required

Purity/activity of isolated protein

Crystallography Enzyme kinetics Antibody production Mutagenesis Multiprotein enzyme complexes

100 mg 10 mg 3 mg 2 mg 10/zg

Very high High Moderate Moderate High

crystallography], these methods have the ability to provide plentiful and pure protein. Prior to designing a strategy for purification, the amount and purity of the enzyme must be assessed for subsequent applications (see Table II). If a small amount of high purity protein is required, for instance in studying multiprotein complexes, a system other than enzyme kinetics or an attempt at crystallography may be employed. Purification and degree of expression can be monitored by use of polyacrylamide gel electrophoresis, whereby expression levels of >2% will be apparent as a new band, and by Western immunoblotting (if a specific antibody is available). Functionality can be evaluated if one has an assay for enzymatic activity to test a panel of samples. Common features are shared among expression vectors (see Fig. 1). 1. Selectable phenotype: This is a genetic utility for the large-scale screening of putative positive clones. Often a marker such as drug immunity is employed. The most commonly used is ampicillin resistance (AmpR). 2. Promoter: This element consists of an RNA polymerase-binding site for tight regulation and is one of the most important determinants of efficient transcription~the frequency of RNA polymerase initiation. These promoters are usually induced by the addition of IPTG (isopropyl-/3-D-thiogalactopyranoside) to the media; IPTG stimulates transcription of the fused lac operon. 3. Ribosome-binding site: The ribosome-binding site is involved in the initiation of mRNA translation in Escherichia coli by the Shine-Dalgarno sequence (complementary to a sequence within the 16S rRNA), bringing the ribosome in close proximity to the initiator codon and an appropriate AUrich translational spacer of four to nine nucleotides. The site is designed for optimum recognition and binding. 4. Purification aids (polylinker, fused affinity tag, protease site): The polylinker introduces restriction endonuclease sites for convenient directional

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[17] CLONING AND EXPRESSION OF ENDOPEPTIDASES

DNA

-~ veCK;F

MARKER

~DRUG I picillin) I

/

" At

[.TrGACANI2TATAAT.,~B ~

/

u G

!

iv

purif. ENDOPEI~IDASE Transcriptio ector Aids CODING SEQUENCE f~ Terminator - ~

transcription mRNA

5 I[I AGGA GG XsATG( C AT/C)5-8" Protease Cleavage Site-ENDOPEt~IDASE C RIBOSOMAL/ BINDINGSIT~[ (RnS)

/

translatiOnprotein

S~detHisHisHisHisHisHis- ProteaseCleavageSite -ENDOPEPTIDASE I

I

~/cleavage purification PURIFIED ENDOPEPTIDASE PROTEIN

FIG. 1 Anatomy of an expression system. Top: Elements involved in a plasmid construct designed to produce high amounts of a specific protein. RBS, Ribosomal binding site; AUG and UGA, translational start and stop codons, respectively. The other elements are described in text. All sequences are written 5' to 3' and correspond to the coding strand. Initiation of transcription and translation are controlled by the promoter and the ribosomal binding site, respectively. Middle: Elements comprising the mRNA. Bottom: Fused protein. Although this depiction is for a prokaryotic system there are analogous sequences in the eukaryotic vector, and the same principles apply. subcloning of enzyme DNA for expression in-frame. Depending on the system employed (described below), a fused affinity tag aids in the purification. To liberate a fusion sequence from the enzyme, a sequence encoding a cleavage site for a protease such as thrombin, factor X, or enterokinase is incorporated. 5. Coding sequence: This portion of the construct encodes, in-frame, the endopeptidase of interest from the first amino acid through the stop codon. 6. Transcription terminator" A transcription terminator is included in the vector to prevent unneeded read-through transcription. This element usually consists of a putative stem-loop structure in the transcript, which aborts transcription. 7. (HIS), tag: Six to eight histidines, synthesized using the two codons, can be placed on the amino or carboxy terminus of the protein of interest,

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Protein Expression Systems

System [prokaryotic (P)/ eukaryotic (E)]

trpE (P) GST/MalE (P) pET (P) Baculovirus (E) Pichia pastoris (E) CHO/COS cells (E)

Fused/unfused

Glycosylation

Fused Fused/unfused Fused/unfused Fused/unfused Fused/unfused Fused

No No No Yes a Yes a Yes

Glycosylation of the core sugars is performed, but is not the same as in mammalian systems. Subcellular targeting must be confirmed.

a

regardless of the vector utilized for expression. Once the protein is expressed, selective purification is achieved through a remarkable affinity of the tag to a resin containing nickel nitrilotriacetic acid (Ni-NTA). Many laboratories are equipped to perform molecular biology and rudimentary protein purification, which is adequate for a bacterial expression system. Lower eukaryotes such as Pichia will perform posttranslational modifications and are amenable to manipulations in a laboratory setting without the high startup costs required for tissue culture facilities. Many of these protein expression systems are available in the form of commercial kits. Representative systems that are most prevalent are summarized in Table III and described below.

trpE Fusion Vectors The prokaryotic trpE fusion vector system was one of the first used on a large scale. It is relatively easy to insert the gene into the polylinker region of the vector. The gene expressing the protein of interest is placed under the strict transcriptional and translational control mechanisms recognized by the bacterial host in order to synthesize fused proteins efficiently. There is often basal expression of the cloned gene owing to the enormous strength of the promoter. Induction can occur by a temperature jump (to 42~ or chemically, by tryptophan starvation or addition of fl-indole acetate. The product can be detected by enzyme assay or antibody to the inserted gene (if available), or by commercially available antibody directed toward the fused portion of the gene, acting as an antigenic marker (trpE) to monitor purification and conjugation. Often the prodigious amounts of protein pro-

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duced partition into inclusion bodies and are thus contained in the insoluble fraction of the cell when lysed. p E T Vectors

pET vectors (Novagen, Madison, WI) utilize the bacteriophage T7 RNA polymerase and the simpler promoter sequences have several advantages compared to E. coli. The T7 system can synthesize longer transcripts, and is more efficient in initiating transcription and translation. It can carry out this synthesis at a fourfold higher rate. Some of the plasmid constructs are inducible by IPTG. A yield of greater than 10% of the cloned gene translated into cellular protein is common, after a few hours of induction. Another feature of using this promoter is that the genes under control ofT7 polymerase are relatively transcriptionally silent, so that there is little protein production in the absence of induction. This is useful for potentially toxic genes, although soluble endopeptidases are not usually included in this class. The original EP 24.15 cloning and expression (3) were performed in a derivative of this system, the Bluescript vector (Stratagene). Sequences are available for cleavage by proteases to yield unfused product. The presence of a filamentous bacteriophage origin allows single-stranded plasmid DNA to be produced for DNA sequencing and site-directed mutagenesis. Glutathione S-Transferase or MalE

Glutathione S-transferase (GST) (Pharmacia-LKB, Piscataway, NJ) or Maltose binding protein (MalE) (New England BioLabs) systems are based on the same principle: fusion to a portion of a protein that binds tightly to a chemical moiety in an affinity resin to allow single-step purification. The pGEX system utilizes a chromatography matrix composed of glutathione coupled to Sepharose. The glutathione has a high affinity for the glutathione S-transferase/enzyme fusion protein. Thrombin is added and incubated in the column to yield free protein, which is then eluted. The MalE system fuses the protein of interest to maltose-binding protein, and signals cytoplasmic expression. The resultant malE-protein fusion binds with high afinity to amylose-conjugated resin. The plasmid utilizes a strong, inducible tac promoter (fused to the lacZ gene and thus inducible with IPTG) and malE initiation translation sequences to yield high amounts of expression. The vector also contains a sequence encoding specific cleavage by protease factor Xa, allowing for nonfused gene product. The vectors also include the filamentous bacteriophage DNA origin of replication, allowing production of single-stranded DNA for both sequencing and site-directed mutagenesis. Elution is conducted with free maltose. In both cases efforts must be made to avoid contamination with the protease utilized for the cleavage of the

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fused protein to the enzyme of interest. This is the current expression system employed for use with EP 24.15 (29). Baculovirus The baculovirus system (Invitrogen) has been reviewed comprehensively (30). Neutral endopeptidase (enkephalinase, EP 24.11), an endopeptidase closely related to EP 24.15 but found as an integral membrane protein, was expressed in a functionally soluble form utilizing an insect cell line infected with baculovirus (31). This eukaryotic expression system, in contrast with the bacterial systems, increases the likelihood of obtaining large quantities of target proteins requiring extensive post- and cotranslational modifications in a biologically active form. A "Baculo-GEX" system has been produced that combines this eukaryotic insect cell line (ligated to the polyhedrin promoter) with the advantages of the GEX system described above for a one-step purification coupled with specific proteolytic cleavage resulting in unfused protein (32). Pichia pastoris Pichia pastoris (Invitrogen) (33) is a eukaryotic expression system, inroduced in the last year and closely related to S. cerevisiae; it is reported to express up to 10 g/liter of a recombinant protein introduced via plasmid into a transformed spheroblast of this methylotropic (fermentation induction by methanol for high-volume, foreign protein productivity), yeastlike organism. An extremely efficient promoter (alcohol oxidase, AOX1) is utilized, which normally allows the organism to process methanol as the sole carbon source. Additionally, the vector contains a coupled HIS4 gene, allowing for screening of recombinants that will grow by plating on histidine-deficient medium. Expression is allowed in either the secreted or intracellular form. Because the medium is virtually protein free, and there are signal sequences in the vector for secretion, purification of the protein is simplified. Expression in yeast requires no additional equipment or media other than that present in common microbiological facilities. CHO or COS Cells In the study of endopeptidases, Chinese hamster ovary (CHO) or African green monkey kidney (COS-CV1) cells [American Type Culture Collection (ATCC), Bethesda, MD] have been used to express a soluble and fully active form of rabbit neutral endopeptidase, which was secreted utilizing transfection of a recombinant expression vector fusing the ectodomain of neutral endopeptidase (NEP) to a cleavable signal peptide (34).

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Hisn Protein Purification Many of the vectors described above include (or the user can add) the oligonucleotide encoding six to eight histidine residues (using a mixture of the two codons, CAC/CAT). The poly(His) can be inserted in-frame at the amino or carboxy terminus (to assure fully terminated proteins), and does not usually interfere with the activity of the cloned, expressed gene, because there is no net charge difference at physiological pH. This enables a rapid purification (--~95% purity achieved in one step) of the protein of interest by selective chelation on a nickel nitrilotriacetate resin by conventional chromatography under native or denaturing conditions. The protein is then eluted from this column under gentle conditions of imidazole as a competitor, or by reducing the pH to about 5.5. Column binding is unaffected by small concentrations of ionic/non-ionic detergents, reducing agents, or strong denaturants. Maximizing heterologous expression levels of recombinant proteins (proteins not native to the particular host organism) in prokaryotes or eukaryotes may present a problem with their solubility. Because high amounts of expression lead to cells that produce several percent of their total protein as foreign biomolecules, these foreign proteins are partitioned, at high concentration, into inclusion bodies (dense aggregates of insoluble, misfolded protein). Often, changing the host strain is all that is needed to change expression and increase partitioning into a soluble fraction. In certain cases this is desirable, because this may simplify obtaining a homogeneous product. The first step of purification involves washing away cytoplasmic proteins and the major contaminants that are misfolded, proteolyzed, or oligomeric forms of the protein of interest. One must be aware that it is possible to produce cytotoxic material deleterious to the viability of the organism. All of these points notwithstanding, the problems with the inclusion body approach deal with issues of purpose and purity. If the purpose is to study a native, folded, intact molecule by biophysical approaches such as NMR or X-ray crystallography, or by an elicited activity in an assay system, then inclusion bodies pose serious problems. With purity as a criterion, the nature of the contaminants as often poorly soluble, hydrophobic membrane components must be considered; variants of the expressed protein are difficult to purify. Refolding proteins quantitatively on a large scale proves a vexing problem. One simple way to avoid aggregates of a soluble recombinant protein in an intracellular system such as E. coli is to lower the temperature (35) to 28-30~ at which the majority of EP 24.15 was found in the soluble form (10). Other solutions involve the choice of expression vector coupled to a secretion system. The addition of protease inhibitors, or detergents, or highdensity growth (as well as optimizing the temperature), can improve secreted yields.

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It is important that after cloning and expressing an enzyme a comparison be made, utilizing enzymological criteria such as catalytic properties toward substrate and inhibitors, with the purified wild-type enzyme derived from a tissue source.

Endopeptidase 24.15 as Example A plasmid vector obtained via library screening and without previous modification for expression is usually not suitable if an unfused protein product is required. The insert containing the soluble endopeptidase must thus be subcloned into an alternative vector. The original recombinant EP 24.15 clone (3) lacked the coding potential for the first two amino acids and was modified by site-directed mutagenesis to yield the complete unfused EP 24.15 with a BamHI site. A 39-nucleotide oligomer was synthesized containing 14 nucleotides identical to the vector, 6 nucleotides identical to the BamHI site, and 19 nucleotides identical to the amino terminus of the protein. The construct was transformed and bacterial cultures containing either plasmid with or without cloned insert, was grown in 100 ml of medium with antibiotic selection and diluted into 1000 ml to an absorbance reading of 0.6 at 600 nm. Expression of the fusion gene was induced with 0.4 mM IPTG, and the growth of properly folded fusion gene product proceeded at 30~ for 3 hr. The bacterial suspension was centrifuged for 5 min at 3000 g and the pellet was resuspended in 20 ml of 50 mM Tris-HCl, pH 7.0. Bacteria were then lysed by two cycles of freezing and thawing followed by sonication. Bacterial debris was removed by centrifugation (10,000 g for 10 min). The bacterial supernatant was then incubated with glutathione-Sepharose beads, and incubated with thrombin to cleave at the junction of the two genes. Thrombin, which contaminates the column eluate by 0.02% (w/w), can be easily removed by exhaustive filtration with a Centricon 50 (Amicon, Beverly, MA) which quantitatively removes thrombin to the filtrate. This yielded pure protein (2.5 mg/liter culture) as assayed by native and SDS-polyacrylamide gel electrophoresis.

Conclusion The cloning of the cDNA encoding an endopeptidase becomes a crucial step in explicating the role that the peptidase plays in nervous system function. Ultimately, elucidating the function and structure of one such protease can aid in understanding the regulation of neuropeptide function by these enzymes as a class. The peptidases can be targeted for pharmacological inter-

[17] CLONING AND EXPRESSION OF ENDOPEPTIDASES

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vention through the use of specifically designed modulatory ligands, either agonistic or antagonistic. Examples using this rationale involve the development of inhibitors of the human immunodeficiency virus (HIV) aspartic protease as a treatment for human immunodeficiency virus (36), inhibitors of angiotensin-converting enzyme, such as captopril, to treat hypertension (37), and inhibitors of enkephalinase as a treatment for congestive heart failure and as a nonaddictive analgesic (38).

Acknowledgments M.J.G. thanks the Revson Foundation for Biomedical Research and a grant from the SEED program of Mount Sinai Medical Center for support of this work.

References M. Orlowski, C. Michaud, and T. G. Chu, Eur. J. Biochem. 135, 81 (1983). 2. T. G. Chu and M. Orlowski, Biochemistry 23, 3598 (1984). 3. A. Pierotti, K. W. Dong, M. J. Glucksman, M. Orlowski, and J. L. Roberts, Biochemistry 23, 10323 (1990). G. R. Acker, C. Molineaux, and M. Orlowski, J. Neurochem. 48, 284 (1987). 5. M. J. Glucksman, N. X. Barrezueta, A. Pierotti, N. S. Bengani, S. Greene, and J. L. Roberts, Endocrinology (1994) (in press). A. Devault, C. Lazure, C. Nault, H. Le Moual, N. G. Seidah, M. Chr6tien, P. Kahn, J. Powell, J. Mallet, A. Beaumont, B. P. Roques, P. Crine, and G. Boileau, EMBO J. 6, 1317 (1987). K. M. Carvalho and A. C. M. Camargo, Biochemistry 20, 7082 (1982). 8. U. Tisljar and A. J. Barrett, Arch. Biochem. Biophys. 274, 138 (1989). 9. J. R. McDermott, J. A. Biggins, and A. M. Gibson, Biochem. Biophys. Res. Commun. 185, 746 (1992). 10. Z. Zeng and M. J. Glucksman, in preparation (1994). 11. B. L. Valee and D. S. Auld, Proc. Natl. Acad. Sci. U.S.A. 87, 220 (1990). 12. M. J. Glucksman, M. Cascio, M. Orlowski, and J. L. Roberts, in press (1994). 13. N. Sugira, H. Hagiwara, and S. Hirose, J. Biol. Chem. 267, 18067 (1992). 14. S. Kawabata, K. Nakagawa, S. Iwanaga, and E. W. Davie, J. Biol. Chem. 268, 12498 (1993). 15. S. G. Oliver, Q. J. M. van der Aart, M. L. Agostini-Carbone, M. Aigle, L. Alberghina, D. Alexandraki, G. Antoine, R. Anwar, J. P. Ballesta, P. Benit, et al., Nature (London) 357, 38 (1992). 16. C. A. Hrycyna and S. Clarke, Biochemistry 32, 11293 (1993). 17. G. Isaya, F. Kalousek, and L. E. Rosenberg, Proc. Natl. Acad. Sci. U.S.A. 89, 8317 (1992). 18. F. M. Ausubel, R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. ~

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19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38.

Smith, and K. Struhl, eds., "Current Protocols in Molecular Biology," Vol. 2. Wiley, New York, 1992. M. J. Glucksman, M. Orlowski, and J. L. Roberts, Biophys. J. 62, 119 (1992). J. Devereux, P. Haeberli, and O. Smithies, Nucleic Acids Res. 12, 387 (1984). C. Gaboriaud, V. Bissery, T. Benchetrit, and J. P. Mornon, FEBS Lett. 224, 149 (1987). T. Benchetrit, V. Bissery, J. P. Mornon, A. Devault, P. Crine, and B. P. Roques, Biochemistry 27, 592 (1988). S. M. Muskal and S. H. Kim, J. Mol. Biol. 225, 713 (1992). S. R. Holbrook, I. Dubchak, and S. H. Kim, BioTechniques 14, 984 (1993). J. Sambrook, E. F. Fritsch, and T. Maniatis, "Molecular Cloning: A Laboratory Manual." Cold Spring Harbor Lab., Cold Spring Harbor, NY, 1989. T. Maruyama, T. Gojobori, S. Aota, and T. Ikemura, Nucleic Acids Res. 14, r151 (1986). D. H. Mack and J. J. Sinisky, Proc. Natl. Acad. Sci. U.S.A. 85, 6077 (1988). E. Ohtsuka, S. Matsuki, M. Ikchara, Y. Takahashi, and D. J. Matsubasu, J. Biol. Chem. 260, 2605 (1985). R. A. Lew, T. Tetaz, M. J. Glucksman, J. L. Roberts, and A. I. Smith, J. Biol. Chem. 269 (1994) (in press). M. Cascio, in "Methods in Neurosciences, Volume 25: Receptor Molecular Biology" (Stuart C. Sealfon, ed.) (in press). F. Fossiez, G. Lemay, N. Labont6, F. Parmentier-Lesage, G. Boileau, and P. Crine, Biochem. J. 284, 53 (1992). A. H. Davies, J. B. M. Jovett, and I. M. Jones, Bio/Technology 11, 933 (1993). J. M. Cregg, T. S. Vedvick, and W. C. Raschke, Bio/Technology 11, 905 (1993). G. Lemay, G. Waksman, B. P. Roques, P. Crine, and G. Boileau, J. Biol. Chem. 264, 15620 (1989). C. H. Schein and M. H. M. Noteborn, Bio/Technology 6, 291 (1988). T. L. Blundell, R. Lapatto, A. F. Wilderspin, A. M. Hemmings, P. M. Hobart, D. E. Danley, and P. J. Whittle, Trends Biochem. Sci. 15, 425 (1990). M. J. Antonaccio and D. W. Cushman, Fed. Proc., Fed. Am. Soc. Exp. Biol. 40, 2275 (1981). E. G. Erdos and R. A. Skidgel, FASEB J. 3, 145 (1989).

[18]

Proteolytic Processing of Amyloid Protein Precursor of Alzheimer's Disease D. H. Small, G. Reed, S. J. Fuller, A. Weidemann, K. Beyreuther, and C. L. Masters

Introduction The two major pathological features in the brain of patients with Alzheimer's disease are neurofibrillary tangles and amyloid plaques. The production of the amyloid plaques is thought to be directly related to the underlying pathogenic mechanism. One line of evidence for this assumption is that the frequency of amyloid plaques correlates approximately with the extent of cognitive impairment (1, 2). The major protein component of the amyloid plaques is a polypeptide known as the amyloid or/3A4 protein (3, 4). Amino acid sequencing of this polypeptide led to the cloning of its precursor, a much larger protein known as the Alzheimer's disease amyloid protein precursor or APP (5). Although the production of amyloid plaques is associated with Alzheimer's disease, APP is a constituent of many normal cell types (6). The identification of rare familial diseases involving point mutations in the APP gene has established the importance of APP in the pathogenesis of Alzheimer's disease (7-9). Although the production of the amyloid protein from APP is linked to the pathogenesis of Alzheimer's disease, the precise relationship between APP and the disease process is unclear. One hypothesis to explain this relationship is that the/3A4 amyloid protein is neurotoxic (10). However, other hypotheses also need to be considered. For example, the disruption of the normal function of APP, caused by inappropriate proteolytic processing, could also contribute to neurodegeneration.

Structure and Function of Amyloid Protein Precursor Multiple molecular weight forms of APP result from alternative mRNA splicing of the APP gene product (11). The major APP mRNA expressed in the brain encodes a protein containing 695 amino acids (5). Two other major transcripts encoding forms with 751 and 770 amino acid residues have been identified (12-14). APP TM is identical to APP 695, except for an extra 56-residue domain homologous to members of the Kunitz family of protease inhibitors. Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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The presence of this Kunitz protease inhibitor (KPI) domain confers the ability to inhibit a few serine proteases (15, 16). APP 77~contains an additional 19-residue domain homologous to the OX-2 antigen (17). The amyloid sequence itself is contained in a stretch of 43 amino acid residues that comprises a small portion of the ectodomain of APP and extends into the transmembrane domain (Fig. 1). Minor forms of APP involving other spliced products have also been identified. A P P 714 contains the OX-2, but lacks the KPI domain (18). A P P 365 and A P P 563 do not contain the transmembrane domain and are therefore thought to be secreted forms (19, 20). Finally, many cells express large amounts of a transcript lacking exon 15 ( A P P 733 o r L-APP) (21). The amyloid protein precursor is posttranslationally modified by N- and O-linked glycosylation, sulfation, and phosphorylation. There are two potential N-linked glycosylation sites close to the transmembrane domain (Fig. 1). Although the sites of O-linked glycosylation have not been identified, a cluster of threonine residues (found next to the acidic region) is a consensus sequence for O-glycosylation (22). The amyloid protein precursor can also be phosphorylated by one or more serine kinases (23); however, the function of this phosphorylation is unknown. As APP is phosphorylated in the ectodomain, this suggests that the phosphorylation of APP could regulate its interaction with extracellular molecules, such as the extracellular matrix (24). The function of APP is still unknown. In the central and peripheral nervous system, APP may be involved in the regulation of neurite outgrowth (24-28). In the embryonic chick brain, APP expression increases during the major phase of neurite outgrowth (24). Other studies (25-28) suggest that APP can directly stimulate neurite outgrowth from neurons or neuronal cell lines in culture. The involvement of APP in the development of the nervous system is also suggested by genetic studies. Drosophila lacking a gene homologous to the human APP gene possess a behavioral deficit that can be corrected with the human APP gene (29).

Proteolytic Processing of the Amyloid Protein Precursor Some of the proteases reported to cleave APP are shown in Table I (30-39). The identification of specific APP-processing enzymes is not an easy task as many different proteases may have the required specificities for cleavage. It seems likely that in vivo, specificity is defined not only by the amino acid sequence around the cleavage site, but also by cellular and subcellular compartmentation of the enzymes with APP. Several criteria should be fulfilled for the unequivocal identification of an APP-processing enzyme. These criteria are as follows.

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P R O T E O L Y T I C PROCESSING OF APP

~ch

s fate']

hepaHn-binding site

~-- Zn binding site ACIDIC THR-rich

KPI ~me~ 9

domain/ i

T~~

LLLLLIJ,,L

FIG. 1 Diagrammatic representation of the structure of APP 695, including the portion of the amyloid protein sequence containing the cleavage site for the APP secretase (Lys-16 in the amyloid sequence). Full-length APP possesses a large ectodomain containing the N terminus, a single transmembrane domain of 24 amino acid residues (residues 625-648 in the APP 695 sequence), and a short cytoplasmic tail of 24 amino acid residues containing the C terminus. Amyloid protein precursor contains two potential N-linked glycosylation sites and a number of domains, including cysteinerich, threonine-rich, and acidic (aspartate and glutamate-rich) domains. Heparinbinding and zinc-binding domains have been identified, along with a growth-promoting domain. APP TM and APP 77~contain an extra sequence with homology to the Kunitz family of proteinase inhibitors (KPI domain) inserted close to the threonine-rich region. APP 77~has an additional domain with homology to the OX-2 antigen. Failure to cleave APP at Lys-16 of the amyloid sequence by the APP secretase results in the preservation of the amyloid sequence. Amyloidogenic forms of APP may subsequently be degraded by amyloid-generating enzymes (AGEs), which cleave adjacent to the methionine residue at position - 1.

1. The protease should show the expected cleavage specificity. For example, the protease should be able to cleave synthetic peptides with sequences homologous to known cleavage sites in APP.

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TABLE I Putative Amyloid Protein Precursor Processing Enzymes Protease

Proposed cleavage site

Ref.

1. Calcium-activated serine protease 2. Multicatalytic protease (calcium regulated)

N-terminus of flA4 Glnl5-Lys 16of/3A4 N terminus of flA4 N terminus of flA4 C terminus of/3A4 Lys16-Leu17of flA4 Lysl6-Leu 17 of flA4 Lys16-Leu17 of flA4 Glu3-Phe4 of flA4 Multiple sites in APP ArgSl~ TM of APP

30 31 31 32 33 34 35 36 37 38 39

3. 4. 5. 6. 7. 8. 9. 10.

Chymotrypsin-like protease (clipsin) Prolyl endopeptidase Acetylcholinesterase-associated protease Cathepsin B Gelatinase A Multicatalytic protease (ingesin) Calpain I Thrombin

2. The spatiotemporal expression of the enzyme should match that of APP or its cleavage products. For example, increased expression of the protease should be associated with increased cleavage of APP, and the tissue distribution of the protease should to some extent reflect the distribution of its substrate. 3. Inhibitors of the protease should inhibit the processing of APP in situ. This would include the use of antisense oligonucleotide methodology. 4. The most stringent criterion for the identification of a processing enzyme is to show that in organisms (or cells in culture) engineered for a deletion in the processing enzyme, there is a reduction or failure to cleave APP. A major route of APP processing involves an enzyme called the "APP secretase," which cleaves APP between Lys-16 and Leu-17 in the flA4 amyloid sequence (40). The resulting C-terminally truncated APP possesses a molecular mass that is approximately 10 kDa lower than the transmembrane protein and is subsequently secreted from the cell (41). Since the secretase cleavage site was first identified, other potential secretase cleavage sites have been found (42, 43). Studies by Sisodia (44) using site-directed mutagenesis have shown that the APP secretase is probably not highly specific in the type of peptide bond it is able to cleave. Instead, the distance of the peptide bond from the plasma membrane is the most critical factor that defines the secretase specificity. Cole et al. (45) provided evidence for the processing of APP through the lysosomal system. Studies by Golde and co-workers (43) suggest that the processing of APP through the endosomal-lysosomal system could result in the production of a complex series of C-terminal derivatives. Some of these

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derivatives retain the amyloid sequence intact and may therefore be amyloidogenic; many cells may secrete these amyloidogenic products. In microglia and astrocytes, amyloidogenic cleavage may be a major pathway of processing (46). The presence of a consensus sequence (NPXY) for coated pit-mediated internalization (47) supports the proposition that APP may undergo endocytosis. The APP secretase is important for understanding the pathogenesis of Alzheimer's disease. Cleavage at Lys-16 destroys the amyloid sequence and thus prevents flA4 amyloid formation. In many cell types, the APP secretase may represent the major processing route. Furthermore, there is some evidence to suggest that a failure to cleave APP at the secretase step could result in amyloidogenic processing. Although candidate enzymes have been proposed (34, 35), the APP secretase has not yet been identified. As with the neuropeptide-processing endopeptidases, a molecular genetic approach may be the only way to clearly identify the secretase. Nonetheless, cell lines that are transfected with APP cDNA constructs are useful for studying the processing of APP through amyloidogenic and nonamyloidogenic pathways. Some general methods for studying the processing of APP are described in the following sections. These methods can be usefully adapted to studying a range of different aspects of APP processing and secretion. The monoclonal antibody (clone 22C11, which can be obtained from Boehringer Mannheim, Indianapolis, IN) used in these procedures recognizes a domain close to the N terminus (residues 66-81) (48). With the identification of other members of a now-expanding APP gene family (49, 50), it is possible that the 22C11 antibody may recognize other APP-like proteins. Thus, the specificity of the antibody must be demonstrated in each tissue under examination. Studying the processing of APP in cells transfected with an APP cDNA expression plasmid eliminates this problem.

T r a n s f e c t i o n of H e L a Cells with an A m y l o i d P r o t e i n P r e c u r s o r Expression Plasmid The method for transfecting HeLa cells with an expression plasmid employs the standard procedure of calcium phosphate coprecipitation. We have used an expression vector (pAPP-695) derived from pUC in which the A P P 695 sequence is inserted at the B a m H I site. The procedure is essentially as described by Weidemann et al. (41). 1. Approximately 24 hr prior to transfection, HeLa cells are split into 75flasks at a density of 1.5 x 10 6 cells/flask. Cells are cultured in 10 ml/

cm 2

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flask of Dulbecco's modified Eagle's medium (DMEM) containing 10% (v/v) fetal calf serum. At least 3 hr before transfection, the growth medium is replaced with fresh medium. 2. Plasmid DNA (24/zg) is precipitated with ethanol and air dried, and then dissolved in 438/zl of 10 mM Tris-HC1 buffer (pH 7.6). Then 62/zl of 2 M CaC12 is added under sterile conditions and the solution allowed to incubate for 10 min at room temperature. 3. During the incubation period, 500/xl of fresh 2 • HeBS (280 mM NaC1, 1.5 M Na2HPO4,50 mM HEPES, titrated to pH 7.13 with NaOH) is aliquoted into 5- or 10-ml round-bottom tubes. The DNA solution is added dropwise to the 2 • HeBS solution with stirring over a period of about 15 sec. The solution is allowed to incubate under sterile conditions at room temperature for 10-30 min, and then the DNA-CaC12 solution is added gently and evenly over the cells and the flask gently agitated. 4. The cells are incubated at 37~ in an atmosphere of 5% CO2 overnight (16-20 hr); the precipitated DNA is then removed, and the cells washed twice with Ca 2§ Mg2+-free phosphate-buffered saline (PBS). Finally the cells are incubated in 10 ml of DMEM containing 10% (v/v) fetal calf serum for 24-48 hr. During this period, the cells should express A P P 695 maximally. Approximately 5-10% of the cells should be transfected using this protocol. The expression and secretion of A P P 695 from transfected HeLa cells can be monitored both by immunoprecipitation and by Western blotting, using a commercially available monoclonal antibody (clone 22C11).

I m m u n o p r e c i p i t a t i o n of the A m y l o i d P r o t e i n P r e c u r s o r after Pulse-Chase Labeling The procedure for immunoprecipitation has been described previously (51). 1. Cells are labeled with [35S]methionine in methionine-free medium in the absence of serum for the required length of time (e.g., 20 min to 1 hr), using any one of several standard protocols. At the end of the pulse, the cells are washed once with methionine-containing medium, chased in methioninecontaining medium for 15 min to 2 hr, and then resuspended in 180 ~1 of Ca 2+ , Mg2+-free PBS containing 2 mM phenylmethylsulfonyl fluoride (PMSF). The cells are disrupted by adding 20/~1 of 10% (w/v) sodium dodecyl sulfate (SDS) and the proteins denatured in a boiling water bath for 5 min. At the end of this period, 200/zl of neutralization buffer [6% v/v) Nonidet P-40

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(NP-40), 200 mM Tris-HC1 buffer (pH 7.4), 300 mM NaC1, 10 mM EDTA, 4 mM NAN3] is added. The solution is sonicated (Branson sonifier on setting 1, four short bursts at 50% intermittency) and then centrifuged in a microfuge at 12,000 rpm for 10 min at 4~ and the pellet discarded. 2. The monoclonal antibody (22C 11 from Boehringer Mannheim) is added (10/zl of a 60-/zg/ml solution) and the tubes are incubated for 1 hr at room temperature with gentle shaking. 3. Protein A-Sepharose (PAS; Pharmacia-LKB, Piscataway, NJ) (7 mg/ incubation tube) is hydrated in STEN buffer [50 mM Tris-HC1 buffer (pH 7.5), 150 mM NaC1, 2 mM EDTA, 0.2% (v/v) Nonidet P-40] for 1 hr and then washed six times with 1 ml of STEN buffer. The PAS is then resuspended in 1 ml of STEN and 1.0 /zl of rabbit anti-mouse immunoglobulin G (IgG) (DAKO Corp., Carpinteria, CA) is added for every milligram of PAS and the mixture incubated for 60 min at room temperature with gentle shaking. The PAS is washed a further three times with STEN, followed by two washes in high-salt STEN (STEN buffer with 500 mM NaC1 instead of 150 mM NaC1) and one wash with STEN. 4. For each incubation tube, 7 mg of anti-mouse IgG-coupled PAS is resuspended in 50/zl of STEN and added to the samples, which are then incubated for 1 hr at room temperature with gentle shaking. After incubation, the gel is washed three times with STEN, twice with high-salt STEN, and once with 10 mM Tris-HCl buffer, pH 7.5. The immunoprecipitated APP can then be analyzed by SDS-polyacrylamide gel electrophoresis (PAGE) followed either by fluorography or analysis with a PhosphorImager (Fig. 2).

Western Blotting of Amyloid Protein Precursor If a nonquantitative assessment of APP levels in a tissue sample is required, Western blotting is an appropriate procedure. Western blotting provides information not only on the level of APP immunoreactivity, but also on the various molecular weight forms of APP in a tissue. The procedure is similar to the procedure of Weidemann et al. (41). 1. Fractions are normally analyzed on 10% (w/v) polyacrylamide gels in the presence of 0.1% (w/v) SDS. Electrophoresis is normally performed with 0.5-mm-thick minigels, using the Bio-Rad (Richmond, CA) Mini-PROTEAN system. After electrophoresis, the proteins are electrophoretically blotted onto polyvinylidene difluoride (PVDF) or nitrocellulose membrane. The PVDF membranes are presoaked in 100% methanol. We transfer at 300 mA for 16 hr, with cooling in a Bio-Rad Trans-Blot cell with plate electrodes.

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B

A

Mr

Mr 2007-

69-

46-

200-

r ~'

110 kDa

110 kDa

97 - ~ - 69--

~100

kDa

9

6-

FIG. 2 Immunoprecipitation of [35S]methionine-labeled APP from PC12 cells (A) and Western blot analysis of a cell homogenate and conditioned medium from HeLa cells (B). (A) PC12 cells were labeled with [35S]methionine (70/zCi/ml) for 1 hr and then the APP immunoprecipitated from a detergent extract of the cells as described in text. The immunoprecipitated [35S]APP was analyzed by SDS-PAGE on a 8.5% (w/v) polyacrylamide gel. (B) HeLa cells were transfected with pAPP-695, using the calcium phosphate coprecipitation method. Proteins were separated by SDS-PAGE, transferred onto nitrocellulose, and stained with a monoclonal antibody (clone 22C 11). Most of the cellular APP is in full-length form and migrates with an apparent molecular mass of l l0 kDa. The secreted form of APP695 in the conditioned medium is C-terminally truncated by the APP secretase and possesses an apparent molecular mass of 100 kDa.

The transfer buffer contains 25 mM Tris base, 190 mM glycine, and 20% (v/v) methanol. 2. After electroblotting, nonspecific binding sites are blocked by incubating for 2 hr at room temperature with a buffer containing 0.1 M Tris-HC1 buffer (pH 7.4), 150 mM NaCI, 0.25% (w/v) bovine serum albumin (BSA), 0.05% (v/v) Tween 20, 2 mM MgC12. The PVDF membrane is incubated with 22C 11 antibody (2/zg/ml) in blocking buffer for 2 hr at room temperature. The membrane is then washed three times with TBST [10 mM Tris-HC1 (pH 8.0), 150 mM NaC1, 0.05% (v/v) Tween 20] and incubated for 2 hr at room temperature with an alkaline phosphatase-conjugated anti-mouse IgG (normally 1" 10,000 dilution) in TBST. 3. After washing three times with TBST, the immunoreactive bands are visualized by staining with a chromogenic buffer for alkaline phosphatase, such as naphthol AS-MX/Fast Red [1 part naphthol AS-MX phosphate (0.4 mg/ml) in water mixed with 1 part Fast Red TR (6 mg/ml) in 0.2 M TrisHC1 (pH 8.0), containing 2 mM MgC12]. Full-length forms of APP are seen

[18] PROTEOLYTICPROCESSING OF APP

325

as multiple broad bands with apparent molecular weights of 100,000 to 130,000 (Fig. 2).

References

10. 11. 12. 13.

14.

15. 16. 17. 18.

M. Roth, B. E. Tomlinson, and G. Blessed, Nature (London) 209, 109 (1966). E. K. Perry, B. E. Tomlinson, G. Blessed, K. Bergmann, P. H. Gibson, and R. H. Perry, Br. Med. J. 2, 1457 (1978). G. G. Glenner and C. W. Wong, Biochem. Biophys. Res. Commun. 120, 885 (1984). C. L. Masters, G. Simms, N. A. Weinman, G. Multhaup, B. L. McDonald, and K. Beyreuther, Proc. Natl. Acad. Sci. U.S.A. 82, 4245 (1985). J. Kang, H. G. Lemaire, A. Unterbeck, J. M. Salbaum, C. L. Masters, K. H. Grzeschik, G. Multhaup, K. Beyreuther, and B. M011er-Hill, Nature (London) 325, 733 (1987). S. Sinha and I. Lieberburg, Neurodegeneration 1, 169 (1992). A. Goate, M. C. Chartier-Harlin, M. Mullan, J. Brown, F. Crawford, L. Fidani, L. Giuffra, A. Haynes, N. Irving, L. James, R. Mant, P. Newton, K. Rooke, P. Roques, C. Talbot, M. Pericak-Vance, A. Roses, R. Williamson, M. Rossor, M. Owen, and J. Hardy, Nature (London) 349, 704 (1991). M. C. Chartier-Harlin, F. Crawford, H. Houlden, A. Warren, D. Hughes, L. Fidani, A. Goate, M. Rosor, P. Roques, J. Hardy, and M. Mullan, Nature (London) 353, 844 (1991). M. Mullan, F. Crawford, K. Axelman, H. Houlden, L. Lilius, B. Winblad, and L. Lannfelt, Nat. Genet. 1, 345 (1992). B. A. Yankner, L. K. Duffy, and D. A. Kirschner, Science 250, 279 (1990). R. E. Tanzi, A. I. McClatchey, E. D. Lamperti, L. Villa-Komaroff, J. F. Gusella, and R. L. Neve, Nature (London) 331, 528 (1988). N. Kitaguchi, Y. Takahashi, Y. Tokushima, S. Shiojiri, and H. Ito, Nature (London) 331, 530 (1988). P. Ponte, P. Gonzalez-DeWhitt, J. Schilling, J. Miller, D. Hsu, B. Greenberg, K. Davis, W. Wallace, I. Lieberburg, F. Fuller, and B. Cordell, Nature (London) 331, 525 (1988). W. E. Van Nostrand, S. L. Wagner, M. Suzuki, B. H. Choi, J. S. Farrow, J. W. Geddes, C. W. Cotman, and D. D. Cunningham, Nature (London) 341, 546 (1989). R. P. Smith, D. A. Higuchi, and G. J. Broze, Jr., Science 248, 1126 (1990). H. Kido, A. Fukutomi, J. Schilling, Y. Wang, B. Cordell, and N. Katunama, Biochem. Biophys. Res. Commun. 167, 716 (1990). M. J. Clarke, J. Gagnon, A. F. Williams, and A. N. Barclay, EMBO J. 4, 113 (1985). T. E. Golde, S. Estus, M. Usiak, L. H. Younkin, and S. G. Younkin, Neuron 4, 253 (1990).

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III EXTRACELLULAR PROCESSING ENZYMES IN THE CNS 19. J. S. Jacobsen, H. A. Muenkel, A. J. Blume, and M. P. Vitek, Neurobiol. Aging 12, 575 (1991). 20. F. De Sauvage and J. N. Octave, Science 245, 651 (1989). 21. G. K6nig, U. M6nning, C. Czech, R. Prior, R. Banati, U. Schreiter-Gasser, J. Bauer, C. L. Masters, and K. Beyreuther, J. Biol. Chem. 267, 10804 (1991). 22. I. B. H. Wilson, Y. Gavel, and G. von Heijne, Biochem. J. 275, 529 (1991). 23. C. Haass, A. Y. Hung, M. G. Schlossmacher, T. Oltersdorf, D. B. Teplow, and D. J. Selkoe, Ann. N.Y. Acad. Sci. 695, 109 (1993). 24. D. H. Small, V. Nurcombe, R. Moir, S. Michaelson, D. Monard, K. Beyreuther, and C. L. Masters, J. Neuroscience 12, 4143 (1992). 25. E. M. Milward, R. Papadopoulos, S. J. Fuller, R. D. Moir, D. Small, K. Beyreuther, and C. L. Masters, Neuron 9, 129 (1992). 26. A. C. LeBlanc, D. M. Kovacs, H. Y. Chen, F. Villar6, M. Tykocinski, L. AutilioGambetti, and P. Gambetti, J. Neurosci. Res. 31, 635 (1992). 27. J. S. Whitson, Neurosci. Lett. 110, 319 (1990). 28. D. H. Small, V. Nurcombe, G. Reed, H. Clarris, R. Moir, K. Beyreuther, and C. L. Masters, J. Neurosci. 14, 2117 (1994). 29. L. Luo, T. Tully, and K. White, Neuron 9, 595 (1992). 30. C. R. Abraham, J. Driscoll, H. Potter, W. E. Van Nostrand, and P. Tempst, Biochem. Biophys. Res. Commun. 174, 790 (1991). 31. S. Kojima and M. Omori, FEBS Lett. 304, 57 (1992). 32. R. B. Nelson and R. Siman, J. Biol. Chem. 265, 3836 (1990). 33. S. Ishiura, T. Tsukahara, T. Tabira, T. Shimizu, K. Arahatz, and H. Sugita, FEBS Lett. 260, 131 (1990). 34. D. H. Small, R. D. Moir, S. J. Fuller, S. Michaelson, A. I. Bush, Q. X. Li, E. Milward, C. Hilbich, A. Weidemann, K. Beyreuther, and C. L. Masters, Biochemistry 30, 10795 (1991). 35. K. Tagawa, T. Kunishita, K. Maruyama, K. Yoshikawa, E. Kominami, T. Tsuchiya, K. Suzuki, T. Tabira, H. Sugita, and S. Ishiura, Biochem. Biophys. Res. Commun. 177, 377 (1991). 36. K. Miyazaki, M. Hasegawa, K. Funahashi, and M. Umeda, Nature (London) 362, 839 (1993). 37. S. Ishiura, T. Tsukahara, T. Tabira, and H. Sugita, FEBS Lett. 257, 388 (1989). 38. R. Siman and G. Christoph, Biochem. Biophys. Res. Commun. 165, 1299 (1989). 39. K. Igarishi, H. Murai, and J. Asaka, Biochem. Biophys. Res. Commun. 185, 1000 (1992). 40. F. S. Esch, P. S. Keim, E. C. Beattie, R. W. Blacher, A. R. Culwell, T. Oltersdorf, D. McClure, and P. J. Ward, Science 248, 1122 (1990). 41. A. Weidemann, G. K6nig, D. Bunke, P. Fischer, J. M. Salbaum, C. L. Masters, and K. Beyreuther, Cell (Cambridge, Mass.) 57, 115 (1989). 42. C. Haass, M. G. Schlossmacher, A. Y. Hung, C. Vigo-Pelfry, A. Mellon, B. L. Ostaszewski, I. Lieberburg, E. H. Koo, D. Schenk, D. B. Teplow, and D. J. Selkoe, Nature (London) 359, 322 (1992). 43. T. E. Golde, S. Estus, L. H. Younkin, D. J. Selkow, and S. G. Younkin, Science 255, 728 (1992). 44. S. S. Sisodia, Proc. Natl. Acad. Sci. U.S.A. 89, 6075 (1992).

[18] PROTEOLYTIC PROCESSING OF APP 45. 46. 47. 48.

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G. M. Cole, T. V. Huynh, and T. Saitoh, Neurochem. Res. 14, 933 (1989). C. Haass, A. Y. Hung, and D. J. Selkoe, J. Neurosci. 11, 3783 (1991). W. J. Chen, J. L. Goldstein, and M. S. Brown, J. Biol. Chem. 265, 3116 (1990). C. Hilbich, U. M6nning, C. Grund, C. L. Masters, and K. Beyreuther, J. Biol. Chem. 35, 26571 (1993). 49. W. Wasco, K. Bupp, M. Magendantz, J. F. Gusella, R. E. Tanzi, and F. Solomon, Proc. Natl. Acad. Sci. U.S.A. 89, 10758 (1992). 50. C. A. Sprecher, F. J. Grant, G. Grimm, P. J. O'Hara, F. Norris, K. Norris, and D. C. Foster, Biochemistry 32, 4481 (1993). 51. G. L. Caporaso, S. E. Gandy, J. D. Buxbaum, and P. Greengard, Proc. Natl. Acad. Sci. U.S.A. 89, 2252 (1992).

[19]

Strategies for Measurement of Angiotensin and Bradykinin Peptides and Their Metabolites in Central Nervous System and Other Tissues Duncan J. Campbell, Anne C. Lawrence, Athena Kladis, and Ann-Maree Duncan

Introduction Whether angiotensin and bradykinin are neuropeptides is a subject of continuing debate. The strength of the evidence for or against such a proposition is dependent on the methodological basis for such evidence. Rather than address this issue directly, in this chapter we describe some of the methodologies we have developed for the measurement of angiotensin and bradykinin peptides and their metabolites in the central nervous system (CNS) and other tissues. In the past, radioimmunoassay (RIA) of angiotensin and bradykinin peptides was based on the use of carboxy (C) terminal-directed antisera. This was due in large part to the ease with which a peptide may be coupled via its amino (N) terminus to carrier proteins for the purpose of immunization. However, for both angiotensin and bradykinin peptides important processing events take place toward the C terminus of the molecule (1-3)- (Figs. 1 and 2). For example, the decapeptide angiotensin I (Ang I) is cleaved between residues 8 and 9 by angiotensin-converting enzyme (ACE, kininase II, EC 3.4.15.1, peptidyl-dipeptidase A) to release angiotensin II (Ang II), and both Ang I and Ang II are cleaved between residues 7 and 8 by a number of endopeptidases to release angiotensin(1-7) [Ang(1-7)]. Both Ang II and Ang(1-7) are bioactive. An alternative pathway of conversion of Ang I to Ang II may involve the sequential cleavage of the two C-terminal residues of Ang I by carboxypeptidase activity (1). Moreover, the nonapeptide bradykinin(1-9) [BK(1-9)] is cleaved between residues 8 and 9 by carboxypeptidases N (kininase I) and M to release bradykinin(1-8) [BK(1-8)], and between residues 7 and 8 by ACE and other endopeptidases to release bradykinin(1-7) [BK(1-7)]. Both BK(1-9) and BK(1-8) are bioactive. When these differentially processed peptides are separated by high-performance liquid chromatography (HPLC), it is of assistance if the peptides of interest 328

Methods in Neurosciences, Volume 23 Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

[19] MEASUREMENTOF ANGIOTENSIN AND BRADYKININ

Mast cell chymase Neutrophil cathepsin G

Aminopeptidase A 1

2

Chymotrypsin

3

4

A s p-A~/~-V ~ a, ,-Ty ~ T rypsi n

6

329

AC E Humanheartchymase

Tonin

7

10

,e-H i s - ~ - P he-~~Le u

~Endopeptidase , %

24.15 ~

" E n d o p e p t i d a s e 24.11

Carboxypeptidases Prolylendopeptidase

FIG. 1 Diagrammatic representation of cleavage sites of angiotensin I by different enzymes. Both endopeptidases 24.11 and 24.15 cleave angiotensin I between residues 4 and 5, and between residues 7 and 8; in addition, endopeptidase 24.11 cleaves angiotensin I between residues 2 and 3. After removal of the amino-terminal aspartic acid by aminopeptidase A, the Arg2 residue can be cleaved by aminopeptidase N. For angiotensin II, the sites of cleavage by endopeptidases are the same as those shown for angiotensin I, except that endopeptidases 24.11 and 24.15 do not cleave between residues 7 and 8 of angiotensin II. ACE, Angiotensin-converting enzyme.

can be measured with the same RIA. To this end, we established N terminaldirected RIA for the measurement of angiotensin and bradykinin peptides and their C-terminal truncated metabolites. In previous attempts to raise N terminal-directed antisera to angiotensin peptides, although the peptides were coupled to the carrier protein via the C terminus, the antisera raised were predominantly directed to the C terminus (4, 5). However, Nussberger et al. (4) found that when Asn ~, VaP-Ang II was acetylated at the N terminus and coupled via the C terminus for immunization, they readily achieved N terminal-directed antisera. This result suggests that acetylation of the N terminus of a peptide renders the N terminus more immunogenic. We used this approach to raise N terminal-directed antisera against N-acetylated angiotensin and bradykinin peptide analogs

Endopeptidase24.15

KininaseII (ACE)

Arg(?-Pro-G ly~ he-Ser-Pro-P he-Arg AminopeptidaseP

~Endopeptidase 2 4 . 1 1 ~ ~ P ~ ~

\

l KininaseI (CarboxypeptidaseN) Prolylendopeptidase Carboxypeptidase M

FIG. 2 Diagrammatic representation of cleavage sites of bradykinin by different enzymes. ACE, Angiotensin-converting enzyme.

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

(3, 6), with the intention of acetylating peptides extracted from biological samples before RIA with these antisera.

Preparation of Antisera For the preparation of antisera directed against the N terminus of Ang II, angiotensin III (Ang III), and B K(1-9), the following peptides are synthesized: N-acetyl-AspArgValTyrIleHisProPheLys (N-Ac-Lysg-Ang II), N-acetyl-ArgValTyrIleHisProPheLys (N-Ac-Lys8-Ang III), and N-acetyl-ArgProProGlyPheSerProPheLys [N-Ac-Lys9-BK(1-9)]. Peptides are synthesized from tert-butoxycarbonyl-protected amino acids, using an Applied Biosystems (Foster City, CA) 430A automated peptide synthesizer. Acetylation of a-amino groups of Lysg-Ang II, Lys8-Ang III, and Lys9-BK(1-9) is performed on the protected resin before hydrogen fluoride treatment (6). Peptides are coupled to bovine thyroglobulin via the C-terminal lysine residue with glutaraldehyde (7), and antisera are raised in rabbits (8). Six rabbits are immunized with each peptide and the best antiserum against each peptide is subsequently used to establish an RIA.

Description of Radioimmunoassays All peptide concentrations are determined by amino acid analysis, using stocks of approximately 1 mg/ml in 20% (v/v) acetic acid in water, and stored at -30~ Working solutions [1 /~M in lysozyme (1 mg/ml), 10 mM acetic acid] are stored at -30~ and discarded after thawing once. All RIA components are diluted with casein phosphate buffer [casein (1 g/liter), 100 mM sodium phosphate, 10 mM disodium ethylenediaminetetraacetic acid (EDTA), sodium azide (1 g/liter), 154 mM sodium chloride, pH 7.0]. A pH optimum of 7.0 has been shown for each of the three assays described. Initially, the total RIA assay volume was 500 /A (6), but this has since been reduced to 250/~1 to increase sensitivity. Although initially prepared on ice, assays are now prepared at room temperature. Each assay tube contains 50/A of diluted antibody, 50/~1 of tracer (---2500 cpm), 50/~1 of standard or unknown peptide solution, and 100/~1 of buffer. Usually the assays are incubated at 4~ for 48 hr before separation of free from bound radioactivity. For the antibody A41 assay, addition of tracer is delayed t~or 48 hr, and the assay is incubated at 4~ for a further 24 hr before separation of free from bound radioactivity. Separation of free from bound radioactivity with albumin/dextran-coated charcoal is performed using a modification (9) of the method described by

[19] MEASUREMENT OF ANGIOTENSIN AND BRADYKININ

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Herbert et al. (10). Stock dextran-coated charcoal [Norit A charcoal (25 g/liter), dextran T10 (2.5 g/liter), 7.1 mM sodium barbitone, 7.1 mM sodium acetate, adjusted to pH ---7.4 with hydrochloric acid] is stirred with bovine serum albumin (BSA, 10 mg/ml) for 1-24 hr at 4~ and then diluted with 4 vol of 150 mM sodium chloride immediately before use. One milliliter of albumin/dextran-coated charcoal is added to each tube at 4~ and, after standing at 4~ for 10 min, the assay tubes are centrifuged at 5000 g for 10 min at 4~ the supernatants rapidly aspirated, and the charcoal pellets counted. Tracer peptides are iodinated with 125I using chloramine-T (11), and the monoiodinated peptides are purified by HPLC on a C18 column, using a gradient of acetonitrile in 0.1% (v/v) trifluoroacetic acid (TFA), and stored in aliquots at -30~ Tracer peptides can be stored for up to 2 months without deterioration in assay performance. Antibody A41 was raised against N-Ac-Lys9-Ang II. The antibody A41 assay uses N, O-diacetyl-Ang II (Ac-Ang II, acetylated as described below) as standard peptide and mono[~ZSI]iodo-Ac-Ang II as tracer. At a dilution of 1 : 270,000, binding of tracer is approximately 50%, and 50% displacement is obtained with ---8 fmol of Ac-Ang II/tube, with a detection limit of ---0.25 fmol/tube. The within-assay coefficient of variation is 6% and the betweenassay coefficient of variation is 19%. Antibody A41 was initially studied using N-Ac-Lys9-Ang II as standard and 125I-labeled N-Ac-Lys9-Ang II as tracer; however, displacement of 125I-labeled N-Ac-Lys9-Ang II by N-AcLys9-Ang II and Ac-Ang II was not superimposable, with incomplete displacement by Ac-Ang II, indicating that a proportion of the antibody population of A41 was specific for N-Ac-Lys9-Ang II. Consequently, N-Ac-Lys 9Ang II cannot be used as standard for the measurement of Ang II levels in biological samples; instead, Ac-Ang II must be used as standard. The use of 125I-labeled Ac-Ang II as tracer has the advantage that Ac-Ang II produces complete displacement of tracer. Antibody A52 was raised against N-Ac-Lysg-Ang III. The antibody A52 assay uses N-Ac-LysS-Ang III as standard peptide and 125I-labeled N-AcLysS-Ang III as tracer. At a dilution of 1 : 48,500, binding of tracer is approximately 50%; assays with antibody A52 have been performed using a total assay volume of only 500 ~1, and 50% displacement is obtained with --~16 fmol of N-Ac-LysS-Ang III/tube, with a detection limit of ---1.0 fmol/tube. The between-assay coefficient of variation is 12%. In contrast to the antibody A41 assay, displacement of 125I-labeled N-Ac-Lys8-Ang III by N-Ac-Lys 9Ang III is identical to that produced by N,O-diacetyl-Ang III (Ac-Ang III), and this assay can be used to measure Ang III in biological samples. Antibody B24 was raised against N-Ac-Lysg-BK(1-9). The antibody B24 assay uses N-Ac-Lys9-BK(1-9) as standard peptide. 125I-Labeled Tyr 8BK(1-9) is acetylated as described below before purification by HPLC, and

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mono[125I]iodo-Ac-Tyr8-BK(1-9) is used as tracer. At a dilution of 1 : 267,000, binding of tracer is approximately 50%; and 50% displacement is obtained with ---2 fmol of N-Ac-Lys9-BK(1-9)/tube, with a detection limit of---0.05 fmol/tube. The within-assay coefficient of variation is 14.5%. Displacement of nSI-labeled Ac-Tyr8-BK(1-9) by N-Ac-Lys9-BK(1-9) is identical to that produced by N,O-diacetyl-BK(1-9) [Ac-BK(1-9)], and this assay can be used to measure B K(1-9) in biological samples.

Characterization of Antisera A complete description of the specificities of the antisera is given elsewhere (3, 6). For all antisera, cross-reactivity studies revealed an absolute requirement for acetylation of the N terminus. For antisera A41 and B24 crossreactivities were 100% for peptides of eight or more residues, 75-80% for peptides of seven residues, and correspondingly less for shorter peptides. For antibody A52, cross-reactivity of Ac-Ang(2-7) was 87.5% of that for AcAng III, with a correspondingly lower cross-reactivity for shorter peptides. In practice, antibody A41 can be used for the measurement of Ac-Ang I, AcAng(1-9), Ac-Ang II, and Ac-Ang(1-7); antibody A52 can be used for the measurement of Ac-Ang(2-10), Ac-Ang(2-9), Ac-Ang III, and Ac-Ang(2-7); antibody B24 can be used for the measurement of Ac-B K(1-9), Ac-B K(1-8), and Ac-BK(1-7).

Acetylation of Peptides The method of acetylation is based on the procedure described by Dobson and Strange (12). Peptides or peptide extracts are taken to dryness in siliconized 13 x 100 mm borosilicate glass tubes, using a vacuum centrifuge (Savant Instruments, Hicksville, NY), then acetylated by sequential addition of 100 /A of water, 10 ~1 of triethylamine, and 5/A of acetic anhydride, with mixing by vortex after each addition. After centrifugation to remove particulate material, the sample is injected directly onto the chromatograph. Alternatively, the acetylated samples may be taken to dryness under vacuum and then dissolved in 120/A of 20% (v/v) acetic acid before centrifugation and injection onto the chromatograph (6). As described below, the acetylation procedure results in the acetylation of residues in addition to the a-amino group of each peptide. We have not identified these other acetylated residues, but they probably include Oacetylation of Try 4 of angiotensin and Ser 6 of bradykinin. In contrast to the N-acetyl group, these O-acetyl groups are labile and can be hydrolyzed by

[19] MEASUREMENT OF ANGIOTENSIN AND BRADYKININ

333

treatment with 10% (v/v) piperidine (3). Samples to be treated with piperidine are taken to dryness following acetylation, then dissolved in 100/xl of 10% piperidine in water and allowed to stand at room temperature for 60 min before evaporation to dryness again, dissolution in 120/zl of 20% acetic acid in water, centrifugation, and injection onto the chromatograph. S e p a r a t i o n of A c e t y l a t e d P e p t i d e s by H i g h - P e r f o r m a n c e Liquid Chromatography All samples are transferred to siliconized microfuge tubes and centrifuged in a microfuge at top speed (15,850 g) for 5 min at room temperature to remove particulate material before the supernatant is injected onto the chromatograph. All separations are performed on a 100 x 4.6 mm Brownlee RP-18 Spheri5 column preceded by a 15 x 3.2 mm RP-18 guard column (Applied Biosystems). The HPLC system consists of two pumps (model 6000A; MilliporeWaters, Milford, MA), an automated gradient controller (model 680; Millipore-Waters), and an injector (Rheodyne, Inc., Cotati, CA) with a 200-/zl sample loop. Solvent A is 0.1% TFA and 0.15 M NaC1 in water; solvent B is 0.1% TFA and 90% acetonitrile in water. Peptides are currently eluted by a linearly increasing gradient of 21-41% solvent B over 30 min, and this may need to be adjusted when the column is changed. The flow rate is 1 ml/min and 0.5-min fractions are collected into 12 x 75 mm borosilicate glass tubes containing 50/zl of protease-free bovine serum albumin (5 mg/ml; (Miles Diagnostics, Kankakee, IL) in water. The solvent blank prepared for assay tubes of the RIA standard curves is 0.5 ml of 31% solvent B in solvent A, added to 50 tzl of bovine serum albumin (5 mg/ml). Fractions and solvent blank tubes are evaporated to dryness under vacuum, and then dissolved in water immediately before RIA. When assayed with one RIA, fractions are dissolved in 120/zl of water and two 50-/zl aliquots taken for RIA of each fraction. When fractions are assayed with more than one RIA, the fractions are dissolved in a correspondingly greater volume of water before RIA. The elution positions of standard angiotensin peptides that were acetylated as described above are shown in Fig. 3A; those that were acetylated and then piperidine treated before HPLC are shown in Fig. 3B. An excellent separation of the different angiotensin peptides is obtained, with N-acetylated peptides (piperidine treated) eluting earlier than N,O-diacetylated peptides. A similar result was obtained for bradykinin peptides (Fig. 4A and B). L a b i l i t y of A c e t y l a t e d P e p t i d e s The first N terminal-directed RIAs we developed were for angiotensin peptides. During the development of these assays we did not suspect that acetyla-

334

III EXTRACELLULAR PROCESSING ENZYMES IN THE CNS 400

Ac Ang (1 7)

A

300

Ac-Ang II

200

Ac-Ang-(1-9)

~

.

100 0

5

400

10

15

20

25

30

35

40

45

50

55

50

55

B N AcAng (1 7 )

300

~

N AcAng II

N Ac Ang I .(1 9)

200

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25 30 35 40 Fraction number

45

FIG. 3 Elution positions on HPLC of standard angiotensin peptides that were either N,O-diacetylated as described in text (A), or N-acetylated (piperidine-treated) (B). HPLC fractions were assayed by RIA with antibody A41, and data have not been corrected for recoveries or cross-reactivity with the antibody. N,O-Diacetylated peptides are designated Ac-; N-acetylated peptides are designated N-Ac-. AcAng(1-7), N,O-diacetylangiotensin(1-7); Ac-Ang(1-9), N,O-diacetylangiotensin(l-9); Ac-Ang II, N, O-diacetylangiotensin II; Ac-Ang I, N,O-diacetylangiotensin I; N-Ac-Ang(1-7), N-acetylangiotensin(1-7); N-Ac-Ang(1-9), N-acetylangiotensin(l-9); N-Ac-Ang II, N-acetylangiotensin II; N-Ac-Ang I, N-acetylangiotensin I.

tion of residues other than the N terminus was occurring, in that the acetylated products appeared to be completely homogeneous, with an efficiency of acetylation of--~100% (6). However, during subsequent development of the N terminal-directed RIA for bradykinin peptides, it was apparent that the acetylated product was not homogeneous. In Fig. 4A it can be seen that small peaks of immunoreactivity elute in the position ofN-acetylated peptides

[19] MEASUREMENTOF ANGIOTENSIN AND BRADYKININ 35o-

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FIG. 4 Elution positions on HPLC of standard bradykinin peptides that were either N,O-diacetylated as described in text (A) or were N-acetylated (piperidine treated) (B). HPLC fractions were assayed by RIA with antibody B24, and data have not been corrected for recoveries or cross-reactivity with the antibody. N,O-Diacetylated peptides are designated Ac-; N-acetylated peptides are designated N-Ac-. AcBK(1-7), N, O-diacetylbradykinin(1-7); Ac-B K(1-8), N, O-diacetyl-bradykinin(1-8); Ac-BK(1-9), N,O-diacetylbradykinin(1-9); N-Ac-BK(1-7), N-acetylbradykinin(1-7); N-Ac-B K(1-8), N-acetylbradykinin(1-8); N-Ac-B K(1-9), N-acetylbradykinin(1-9).

(compare Fig. 4A and B). N-Acetyl-BK(1-9)represents <10% ofN, O-diacetyl-BK(1-9), but for N, O-diacetyl-BK(1-8) and N, O-diacetyl-BK(1-7) the N-acetylated peptides represent ---20% of the corresponding diacetylated peptides. Our experience with acetylation of bradykinin peptides led us to examine whether angiotensin peptides were also acetylated at residues other than the

336

III

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

N terminus. As shown in Fig. 3, this was in fact occurring, but the product of acetylation was completely homogeneous. One concern resulting from the data shown in Fig. 3A and B is that N-acetyl-Ang I elutes in the same position as N,O-diacetyl-Ang II. It was therefore necessary to exclude the possibility that the N, O-diacetyl-Ang II peak in acetylated biological samples contains a contribution from N-acetyl-Ang I. This possibility would be of particular concern for samples obtained following administration of an ACE inhibitor, where Ang II levels would be expected to decrease, in association with a large increase in Ang I levels. Evidence that the N,O-diacetyl-Ang II peak does not contain a contribution from N-acetyl-Ang I includes the following: (a) as stated above, when either Ang II or Ang I were acetylated, the product was completely homogeneous (6); (b) when human plasma samples or rat kidney homogenates were spiked with ---1000 fmol of Ang I before extraction, acetylation, and HPLC, there was no increase in the measured amount of Ang II in each sample (6, 13). Nevertheless, we observed that when acetylated samples were inadvertently allowed to stand at room temperature for several hours, partial deacetylation of the N,O-diacetyl-Ang I did occur. For this reason, and because of our experience with acetylated bradykinin peptides, we take particular care to prevent hydrolysis following acetylation. Samples are injected directly onto the chromatograph following acetylation or, when an automatic injector is used, the carousel of the injector is refrigerated to ---3~ For N,O-diacetylated bradykinin peptides (Fig. 4A), it will be noted that Nacetyl-BK(1-8) elutes close to the elution position of N, O-diacetyl-BK(1-9). This is of little concern in practice, however, because BK(1-8) levels in biological samples are much less than those of BK(1-9) (3). Our experience has been that piperidine treatment of acetylated extracts does not result in an appreciable increase in the measured peaks of immunoreactivity in HPLC fractions. For routine assay, samples are N,O-diacetylated as described above. When O-deacetylation may occur, this is corrected for because all estimates of endogenous peptide levels in biological samples are corrected for the recovery of standard peptides from tissue homogenates treated in an identical manner.

Extraction of Peptides from Plasma, Blood, and Tissues Measurement of peptides in biological samples requires the prevention of peptide generation and metabolism during processing of the sample. For the measurement of angiotensin peptides in plasma, trunk blood from decapitated rats or blood obtained by venepuncture from human subjects is collected directly into 0.1 vol of an inhibitor cocktail [50 mM 1,10-phenanthro-

[19] MEASUREMENT OF ANGIOTENSIN AND BRADYKININ

337

line, 125 mM EDTA, neomycin sulfate (2 g/liter), and 2% ethanol in water] containing an appropriate renin inhibitor at a concentration sufficient to inhibit renin activity completely (6, 13). The blood is then centrifuged and the plasma immediately extracted with Sep-Pak C~8 cartridges (MilliporeWaters). Each Sep-Pak cartridge is first moistened with 3 ml of methanol and then washed with 10 ml of 1% TFA in water. To minimize nonspecific adsorption, the cartridges are next coated with 1 ml of a 1% polypeptide solution (Polypep; Sigma Chemical Company, St. Louis, MO) and washed again with methanol-water-TFA (80:19:1, v/v/v) and with 1% TFA (14). The sample is then applied and the cartridges washed with 10 ml of 1% TFA- 1% sodium chloride (1 : 1, v/v), then with 2 ml of methanol-water-TFA (30:69: 1, v/v/v), before elution with 6 ml of methanol-water-TFA (80: 19: 1, v/v/v) into a siliconized 13 x 100 mm borosilicate glass tube. The eluate is then cooled to -80~ before evaporation to dryness under vacuum. Cooling of the sample is a precaution against sample loss due to boiling in the vacuum centrifuge. Plasma extracts can then be acetylated and subjected to HPLC. The measurement of circulating levels of bradykinin peptides requires different methodology because of the need to prevent activation of plasma prekallikrein. For the measurement of bradykinin peptides in rat blood, blood is collected from either conscious rats with previously implanted arterial cannulae or from anesthetized rats by needle puncture of the aorta or inferior vena cava. Two milliliters of blood is collected directly into a syringe containing 10 ml of 4 M guanidine thiocyanate (GTC), 1% TFA in water, and the mixture is then homogenized using a Polytron with a 1-cm aggregate (model PT 10-35; Kinematica, Lucerne, Switzerland) operating at maximum speed. The mixture is then centrifuged at 5000 g for 20 min at 20~ and the supernatant extracted by Sep-Pak as described above. Blood levels of angiotensin peptides can also be measured by this method. Tissues are rapidly removed from decapitated rats, weighed, and immediately homogenized in 10 ml of GTC-TFA, sonicated briefly, and then processed as described for blood. The weight of tissue homogenized is no more than 0.5-1.0 g. Higher amounts of tissues result in lower recoveries from the extraction procedure. The use of GTC-TFA for the homogenization of tissues is a modification of the method described by Zingg et al. (15). Extracts of blood and tissue are extracted with diethyl ether before acetylation and HPLC. Each dried extract is dissolved in 1 ml of 0.1 M hydrochloric acid, then extracted twice with 1 ml of diethyl ether, frozen, and taken to dryness again under vacuum. We have previously used 1.0 M hydrochloric acid for this procedure (3, 13, 16), but 0.1 M hydrochloric acid gives equivalent results.

338

III

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

The blank for each assay was assessed by extraction of 10 ml of GTC-TFA, and these blank extracts were processed as described above and then subjected to HPLC before RIA.

Validation of Methodology Details of the recoveries of peptides extracted by these methods are given elsewhere (3, 6, 13, 16). In general, peptide recoveries approximate 40-50%. A number of different approaches were used to validate the methodology described. 1. Radioimmunoassays of HPLC fractions of blank extracts gave results at or below the detection limit(3, 6). 2. Our identification of individual angiotensin and bradykinin peptides in biological samples was based on criteria additional to their recognition by the appropriate antisera. For N,O-diacetylated angiotensin and bradykinin peptides, the identified peptides were shown to elute in the same position as N,O-diacetylated standard peptides on HPLC. Moreover, in the case of bradykinin peptides, when samples were N-acetylated (piperidine treated), the identified peptides were shown to elute in the same position as N-acetylated standard peptides on HPLC. 3. C terminal-directed antisera raised against Ang II, Ang I, Ang(1-9), and BK(1-9) (3, 9, 13) were able to recognize these peptides when acetylated, although with a reduced efficiency. By using these C terminal-directed antisera we were able to confirm the identity of many of the peaks of immunoreactivity measured in HPLC fractions of biological samples by N terminaldirected RIA (3, 6, 13). 4. We studied the effect of time delay in removal of rat tissues on the measured tissue levels of angiotensin and bradykinin peptides. When a comparison was made between the left and right kidneys homogenized 30 and 90 sec after decapitation, respectively, no difference in angiotensin or bradykinin peptide levels was evident (3, 13). This does not indicate that renal angiotensin and bradykinin peptides are protected from metabolism; rather, the relatively constant level of angiotensin and bradykinin peptides during the delay before homogenization indicates that during this time peptide metabolism approximates the production rate of these peptides in the kidney. 5. To determine the stability of bradykinin peptides in GTC-TFA tissue homogenate, kidney homogenates were divided into two equal portions that were extracted either 1 or 2 hr after homogenization. No difference in the measured levels of bradykinin peptides was evident, indicating that peptides are stable in GTC-TFA tissue homogenates (3). By contrast, for GTC-TFA

[19] MEASUREMENT OF ANGIOTENSIN AND BRADYKININ

339

homogenates of blood we did observe a significant decrease in BK(1-9) levels, from 3.00 -+ 0.03 fmol/ml (mean _ 0.03 fmol/ml (mean _+ SEM, n = 6) for homogenates extracted after 1 hr, to 2.3 +__0.3 fmol/ml (p < 0.05) for homogenates extracted after 2 hr (3). 6. In another approach to assess the stability of angiotensin and bradykinin peptides in GTC-TFA tissue homogenates, kidney homogenates were divided into two equal portions, to one of which was added --~1000 fmol of either Ang II, Ang I, or BK(1-9). Subsequent analyses showed no evidence of metabolism of the added peptides, except for a small increase in B K(1-8) levels in the homogenate to which BK(1-9) was added, which represented <1% of the added amount of B K(1-9), again indicating that peptides are stable in GTC-TFA homogenates (3, 13). The same approach was used to show that metabolism of plasma angiotensin peptides did not occur during processing of plasma samples (6). 7. To examine whether homogenization of tissue in GTC-TFA causes an immediate arrest of bradykinin peptide degradation and generation and avoids the possible consequences of activation of prekallikrein, we also measured bradykinin peptide levels in kidneys that were snap-frozen. Following decapitation of each of six rats, the left kidney was immediately homogenized in GTC-TFA at room temperature, and the right kidney immediately clamped between two metal plates that had been cooled to the temperature of liquid nitrogen. The frozen kidney was then pulverized in a metal mortar and pestle that had been cooled to the temperature of liquid nitrogen. The powdered frozen kidney was added to GTC-TFA at 0~ and immediately homogenized by use of a Polytron. Similar bradykinin peptide levels were measured for kidneys that were either immediately homogenized in GTC-TFA at room temperature or rapidly cooled to the temperature of liquid nitrogen before pulverization and homogenization, thus providing further evidence for the efficiency with which bradykinin peptide degradation and generation were arrested by homogenization in GTC-TFA (3). 8. Another aspect of the process of validation was to show that the circulating and tissue levels of angiotensin and bradykinin peptides changed in the manner predicted following administration of an ACE inhibitor (3, 6, 13).

Potential Problems Many different methods for the extraction of peptides from tissue have been described. In the present studies, blood and tissues were homogenized in GTC-TFA because GTC is a potent chaotropic agent that rapidly denatures

340

III

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

proteins and arrests enzymatic activity, and 1% TFA, by lowering the pH of the sample, also inactivates many proteolytic enzymes. However, it is important to be aware of the possibility that the extraction method may modify some peptides. For example, oxidation of methionine residues may occur, and Smyth (17) has described a strategy to overcome this problem. Other possibilities are the deacetylation of O-acetylated residues (18) and the deamidation of asparagine and glutamine residues by exposure to acid (19). Fortunately these potential problems do not apply to angiotensin and bradykinin peptides. We chose to homogenize fresh tissue immediately because such an approach inactivates proteolytic enzymes and thus prevents peptide generation and metabolism during subsequent processing of the sample. Many authors freeze tissues before subsequent homogenization and extraction of peptides. We have avoided the freezing of tissue because of the potential for peptide generation and metabolism during freezing and thawing. An example of the problems that may occur is shown by our study of the effect of different methods of extraction of rat hypothalami on the measured levels of angiotensin peptides (16). Angiotensin II levels in rat hypothalami are low (--~18 fmol/ g wet weight), and Ang(1-7) levels are below the minimum detectable (<6 fmol/g). However, if hypothalami that had been frozen in liquid nitrogen were allowed to thaw before homogenization in GTC-TFA, the measured levels of Ang(1-7) were --~100 fmol/g. A similar result was obtained when frozen tissue was boiled in 0.1 M hydrochloric acid (16). It is important to use an HPLC protocol that produces optimal separation of the peptides of interest because it is not possible to predict whether other immunoreactive material may elute close to the peptides of interest. This problem is illustrated by our studies of angiotensin peptides in plasma of rats administered the ACE inhibitor perindopril. Shown in Fig. 5 are HPLC profiles for plasma angiotensin peptides of a control rat (Fig. 5A) and a rat administered perindopril (4.2 mg/kg/day) in the drinking water for 7 days (Fig. 5B). Note that ACE inhibition was associated with large increases in Ang(1-7) and Ang I levels, with a reduction in Ang II levels. Fractions were not collected for measurement of Ang(1-9) levels in these experiments. Angiotensin-converting enzyme inhibition was associated with the appearance of a peak of immunoreactivity in fraction 45. The identity of this peak is not known but it is probably related to the markedly increased Ang I levels. These profiles emphasize the need to optimize HPLC conditions to achieve maximal separation of cross-reacting peptides because such an approach ensures maximal separation of other "unanticipated" peaks of immunoreactivity from the peaks of interest. Becuase of the need to differentiate authentic peptide peaks from such "unanticipated" peaks, we always assay individual fractions rather than pool fractions for RIA.

341

[19] MEASUREMENT OF ANGIOTENSIN AND BRADYKININ 100

"A

80 tO

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60

Ac-Ang II

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40

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.

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FIG. 5 Characterization of angiotensin peptides in plasma of a control rat (A) and a rat administered perindopril (4.2 mg/kg/day) in the drinking water for 7 days (B). Plasma samples were extracted, N,O-diacetylated, run on HPLC, and angiotensin peptides measured by RIA with antibody A41. Fractions containing N, O-diacetylangiotensin(1-9) were not assayed in this experiment. Ac-Ang(1-7), N, O-diacetylangiotensin(1-7); Ac-Ang II, N, O-diacetylangiotensin II; Ac-Ang I, N, O-diacetylangiotensin I.

Advantages and Disadvantages of Methodology The advantages of the methodology described are as follows: (a) the combined use of HPLC and N terminal-directed antisera enables the simultaneous RIA of a number of peptides of interest. Moreover, determination of the ratio of the C terminal-truncated metabolites to the parent peptide gives important information concerning the activity of these pathways of peptide metabolism in vivo. For example, the Ang II/Ang I ratio in plasma or tissues

342

III EXTRACELLULARPROCESSING ENZYMES IN THE CNS is a sensitive indicator of Ang I conversion to Ang II at these sites in oioo; (b) an excellent separation of acetylated peptides is achieved by HPLC; (c) the RIAs are sensitive and robust. The N terminal-directed RIAs for the angiotensin peptides have sensitivities equivalent to the most sensitive described, and the N terminal-directed bradykinin RIA is severalfold more sensitive than the most sensitive described. Disadvantages of the methodology include the following: (a) the methodology is time consuming and tedious. To some extent this can be overcome by the automation of HPLC and of RIA; (b) precautions are necessary to prevent the deacetylation of labile O-acetyl groups. Alternatively, acetylated samples could be routinely treated with piperidine before HPLC; (c) acetylation may result in a lower peptide recovery than might be achieved if samples are not acetylated before HPLC; (d) the N terminal-directed RIA cannot be used to measure N terminal-extended peptides. Thus, while the antibody B24 assay is ideal for studies in the rat, where BK(1-9) is the predominant kinin peptide, this assay cannot be used to measure Lys~ which may be the predominant kinin peptide in humans (20).

Acknowledgments The studies reported here were supported by grants from the National Health and Medical Research Council of Australia, and by the National Heart Foundation of Australia.

References 1. D. J. Campbell, in "The Renin-Angiotensin System" (J. I. S. Robertson and M. G. Nicholls, eds.), p. 23.1. Gower Medical Publishing, London, 1993. 2. J. W. Ryan, Am. J. Physiol. 257, L53 (1989). 3. D. J. Campbell, A. Kladis, and A.-M. Duncan, Hypertension (Dallas) 21, 155 (1993). 4. J. Nussberger, G. R. Matsueda, R. Re, and E. Haber, J. Immunol. Methods 56, 85 (1983). 5. J. R. Stockigt, H. M. Shizgal, and W. F. Ganong, Int. Congr. Ser. Excerpta Med. 241, 290 (1972). 6. A. C. Lawrence, G. Evin, A. Kladis, and D. J. Campbell, J. Hypertens, 8, 715 (1990). 7. H. M. Geysen, S. J. Barteling, and R. H. Meloen, Proc. Natl. Acad. Sci. U.S.A. 82, 178 (1985). 8. J. Vaitukaitis, J. B. Robbins, E. Nieschlag, and G. T. Ross, J. Clin. Endocrinol. Metab. 33, 988 (1971).

[19] MEASUREMENTOF ANGIOTENSIN AND BRADYKININ

343

D. J. Campbell and A. Kladis, J. Hypertens. 8, 165 (1990). 10. V. Herbert, K.-S. Lau, C. W. Gottlieb, and S. J. Bleicher, J. Clin. Endocrinol. Metab. 25, 1375 (1965). 11. W. M. Hunter and F. C. Greenwood, Nature (London) 194, 495 (1962). 12. P. R. M. Dobson and P. G. Strange, in "Methods in Enzymology" (L. Birnbaumer and B. O'Malley, eds.), Vol. 109, p. 827. Academic Press, Orlando, FL, 1985. 13. D. J. Campbell, A. C. Lawrence, A. Towrie, A. Kladis, and A. J. Valentijn, Hypertension (Dallas) 18, 763 (1991). 14. K. Hermann, R. E. Lang, T. Unger, C. Bayer, and D. Ganten, J. Chromatogr. 312, 273 (1984). 15. H. H. Zingg, D. Lefebvre, and G. Almazan, J. Biol. Chem. 2,61, 12956 (1986). 16. A. C. Lawrence, I. J. Clarke, and D. J. Campbell, J. Neuroendocrinol. 4, 237 (1992). 17. D. G. Smyth, Anal. Biochem. 136, 127 (1984). 18. M. E. Goldman, M. Beaulieu, J. W. Kebabian, and R. L. Eskay, Endocrinology (Baltimore) 112, 435 (1983). 19. A. I. Smith and R. A. Lew, this volume [7]. 20. F. Alhenc-Gelas, J. Marchetti, J. Allegrini, P. Corvol, and J. Menard, Biochim. Biophys. Acta 677, 477 (1981). .

[20]

Distribution and Roles of Endopeptidase 24.11 Anthony J. Turner and Kay Barnes

Introduction A variety of peptidases located in the plasma membrane of many different cell types are able to effect the postsecretory processing or metabolism of synaptically released neuropeptides. These peptidases, typified by endopeptidase 24.11 (E-24.11) (neprilysin; EC 3.4.24.11) and angiotensin-converting enzyme (peptidyl dipeptidase A; EC 3.4.15.1), are integral membrane proteins oriented such that their active sites face the extracellular space, that is, they are ectoenzymes. Association of these peptidases with the plasma membrane usually involves a transmembrane peptide anchor either at the N or C terminus of the protein, although for some peptidases (e.g., aminopeptidase P) a glycolipid anchor is employed (1). For a relatively few brain neuropeptidases the structure, topology, and cellular localization have been precisely defined. Some of the strategies that can be employed to establish such information are the subject of this chapter and focus predominantly on E-24.11, although the methods are readily adaptable to other enzymes. Procedures for routine assay of the major neuropeptidases have been described elsewhere (2).

Endopeptidase 24.11 Endopeptidase 24.11 was first identified not in the nervous system but in the renal brush border, where it is a major integral membrane protein. It is a cell surface zinc metallopeptidase whose specificity is normally directed toward the hydrolysis of peptide bonds involving the amino group of hydrophobic residues (3). The enzyme is now known to be widely distributed (4) and, in the nervous system, serves to inactivate a range of neuropeptides including opioid and tachykinin peptides. The peptidase has also been located in the stratum oriens and stratum radiatum of the hippocampus, where it may have a key role in degrading somatostatin (K. Barnes, S. Doherty, and A. J. Turner, unpublished results, 1994). The range of peptides hydrolyzed by the enzyme in vitro is broad, ranging from the neutrophil chemotactir tripeptide fMet-Leu-Phe to larger neuropeptides such as vasoactive intestinal 344

Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

[20] ENDOPEPTIDASE 24.11 TABLE I

345

Properties of Pig Endopeptidase 24.11

Plasma membrane ectoenzyme Mr 87,000 Carbohydrate (13%) Metalloprotein (Zn2+) Inhibited by chelating agents (EDTA, 1,10-phenanthroline) Inhibited by phosphoramidon (Ki 2 nM), thiorphan Preferred specificity: Amino side of hydrophobic residues (e.g., Phe, Tyr, Leu, Trp) Main location in the nervous system: Choroid plexus, striatum, globus pallidus, substantia nigra, olfactory tubercle Physiologically relevant substrates: Enkephalins, tachykinins, atrial natriuretic peptides

polypeptide (28 amino acids), atrial natriuretic peptide (28 amino acids), and calcitonin gene-related peptide (37 amino acids). By using a combination of light and electron microscopic immunohistochemistry we have shown the enzyme to be heterogeneously distributed in the brain and localized to neuronal plasma membranes (5, 6). The enzyme may therefore have distinct physiological roles at different sites in the nervous system. Evaluating the precise localization of this, and related neuropeptidases, is therefore critical to understanding the precise architecture of neuropeptide synapses. Molecular cloning of E-24.11 has revealed its identity with the common acute lymphoblastic leukemia antigen (CALLA or CD 10), implying an important role for the enzyme in the immune system, and emphasizing the similarities of signaling mechanisms in, and interactions between, the nervous and immune systems (7). Some properties of E-24.11 are summarized in Table I.

E c t o e n z y m e S t a t u s of N e u r o p e p t i d a s e s Degradation of released neuropeptides could theoretically occur through the actions of ectoenzymes or soluble, secreted peptidases although the focus of this chapter is on the membrane peptidases. Specific experimental criteria need to be fulfilled to establish a putative neuropeptidase as an integral membrane protein and, more specifically, as an ectoenzyme. If anchored by a glycosyl-phosphatidylinositol (GPI) anchor then the protein may be releasable in a hydrophilic form by phosphatidylinositol-specific phospholipase C (PI-PLC) (8). GPI-anchored proteins are, by definition, ectoenzymes because the whole of the polypeptide is external to the cell surface and the only association with the plasma membrane is through the phospholipid tail. For transmembrane proteins, detergents are needed to release them in an intact form from the membrane. When solubilized by nonionic detergents such as

346

III

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

Triton X-100 or octylglucoside the hydrophobic anchor is protected from the aqueous phase by a detergent micelle and the protein exhibits amphipathic properties. The ability of the purified protein to reconstitute into liposomes is an indicator of amphipathic nature. The reconstitution procedure can be monitored by flotation analysis on centrifugation (9, 10). The fractions containing liposomes can be identified by use of a labeled phospholipid (e.g., [3H]dioleoylphosphatidylcholine) as a marker. Proteins that are associated only with the membrane in a peripheral manner, for example, cytosolic proteins associating with the membrane on homogenization, can often be removed by washing the membrane preparation with 0.5 M NaC1 and/or treatment with chelating agents. A useful technique to establish the integral nature of the association of a protein with the membrane is that of phase separation in the detergent Triton X-114. This technique, originally devised by Bordier (11) and now in wide use, relies on the ability of solutions of this particular detergent to separate into detergent-enriched and detergentdepleted phases at 30~ Amphipathic proteins, such as E-24.11, partition predominantly into the detergent-rich phase whereas hydrophilic proteins (peripheral or cytosolic proteins) partition into the detergent-depleted phase. A modification of this method (12) into a three-phase system can allow further distinction between transmembrane and GPI-anchored membrane proteins. The two-phase system described in the procedure below is adequate to establish the integral nature of a membrane protein. It can be applied either to membrane preparations or to purified proteins. Typical results for several membrane proteins after fractionation of a membrane preparation are shown in Table II, which compares some GPI-anchored proteins (alkaline phosphatase and membrane dipeptidase) and transmembrane proteins (E-24.11 and dipeptidyl peptidase IV) before and after treatment with PI-PLC.

Procedure: Temperature-Induced Phase Separation in Triton X-114 1. Dilute protein sample (maximum volume 50/zl and 0.5 mg of protein) to 0.2 ml with 10 mM Tris-HC1 buffer, pH 7.4, containing 0.15 M NaC1, 1% (v/v) precondensed Triton X-114 (11); mix and leave on ice for 5 min. 2. Layer the mixture onto a sucrose cushion consisting of 0.3 ml of a solution of 6% (w/v) sucrose buffered with 10 mM Tris-HCl, pH 7.4, containing 0.15 M NaC1, 0.06% (v/v) Triton X-114, in a microcentrifuge tube and incubate at 30~ for 3 min. Centrifuge in a swing-out rotor at 3000 g for 3 min at room temperature. 3. Transfer the upper aqueous phase (0.2 ml) to a fresh tube and add Triton X-114 to a final concentration of 0.5% (v/v). Mix and set on ice for 5 min.

[20]

347

E N D O P E P T I D A S E 24.11

TABLE II Triton X-114 Phase Separation of Pig Ectoenzymes a E n z y m e activity in detergent-rich p h a s e (% of total activity)

Source a. U n t r e a t e d m e m b r a n e s b. Octylglucoside-solubilized

E-24.11

DPP-IV

MDP

AP

84.9 72.8

90.3 81.9

94.0 92.6

94.6 80.0

ND

ND

4.3

1.9

supernatant c. P I - P L C - r e l e a s e d supernatant

a Microvillar membranes from pig kidney (60/xg of protein) (a), and the 31,000 g supernatant after treatment of microvillar membranes either with 60 mM octylglucoside for 1 hr at 4~ (b), or with Bacillus thuringiensis PI-PLC (1 unit/ml) for 1 hr at 37~ (c), were made up to 0.2 ml with 1% Triton X-114-150 mM NaCI-10 mM Tris-HCl buffer, pH 7.4, and subjected to phase separation at 30~ for 3 min as described by Bordier (11). The detergent-rich and detergent-depleted phases were separated through a sucrose cushion by centrifugation at 3000 g for 3 min at room temp and assayed for enzyme activities. Activities recovered are expressed as percentages of the total activity. AP, Alkaline phosphatase; DPP-IV, dipeptidyl peptidase IV; E-24.11, endopeptidase 24.11; MDP, membrane dipeptidase; ND, not detectable. AP and MDP are GPI-anchored enzymes whereas DPP IV and E-24.11 are transmembrane polypeptide anchored. [Adapted from N. M. Hooper and A. J. Turner, Biochem. J. 250, 865 (1988), with permission of Portland Press, London.]

4. Carefully overlay the sample from step 3 onto the original sucrose cushion; incubate at 30~ for 3 min and centrifuge at 3000 g for 3 min as before. 5. Repeat the wash procedure (step 3) by removing the upper aqueous phase (0.2 ml) to a fresh tube and add Triton X-114, this time to a final concentration of 2% (v/v). Mix and set on ice for 5 min. 6. Incubate the tube from step 5 at 30~ for 3 min, centrifuge at 3000 g for 3 min, and remove 0.15 ml of the supernatant to a fresh tube. This constitutes the detergent-depleted phase into which hydrophilic proteins partition. 7. Carefully remove the sucrose cushion from above the detergent-rich phase in the original tube and make up to 0.15 ml with 10 mM Tris-HC1 buffer, pH 7.4, containing 0.15 M NaC1. This constitutes the detergent-rich phase into which integral membrane proteins partition. 8. Assay each phase for required activities.

Subcellular Fractionation Subcellular fractionation of brain tissue can provide useful indications of the localization of specific peptidases and detection of a particular activity in intact synaptosomes that is not significantly increased on hypotonic or deter-

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E X T R A C E L L U L A R PROCESSING ENZYMES IN THE CNS

gent lysis is a good indicator of an ectoenzyme activity. Because lysed synaptosomes contain many nonspecific proteinases, it is highly desirable to have a specific method of monitoring the peptidase of interest, for example, by use of a specific inhibitor or antibody. Proteolysis with, for example, papain or trypsin under mild conditions may effect release of a membrane peptidase of interest in a hydrophilic form without significant disruption of the synaptosome population, again indicating an ectoenzyme. Some peptidases (e.g., E-24.1 l) are, however, resistant to release by proteinases. Integrity of synaptosomes can be conveniently assessed by measurement of the occluded lactate dehydrogenase activity (13). Traditional methods for preparing synaptosomes are time consuming and can lead to relatively low yields and damaged synaptosomes. A number of improvements in methodology have occurred and a particularly useful procedure involves a rapid fractionation method on a discontinuous Percoll gradient (14). An important advantage of Percoll over other fractionation media is its low viscosity, which allows more rapid sedimentation and the use of lower centrifugal forces; isotonicity of the medium can also be maintained. Preparation of the conventional crude mitochondrial or "P2" fraction is not required, avoiding a potentially disruptive resuspension step. The overall time for the preparation of synaptosomes by this method is less than 1 hr and leads to relatively homogeneous material that is specifically depleted of damaged synaptosomes, synaptic plasma membranes, as well as identifiable myelin and extra-synaptosomal mitochondria. The technique involves two centrifugation steps, first producing an S 1 fraction; further centrifugation of this fraction on a four-step discontinuous Percoll gradient produces five main fractions (interface fractions F1-F4 and a pellet, F5) and allows resolution of viable from nonviable synaptosomes. Purified viable synaptosomes are obtained in the fraction at interface 4. A detailed procedure from Dunkley et al. (14) for the fractionation follows.

Procedure" Preparation of Synaptosomes by Discontinuous Percoll Gradient Fractionation Seven discontinuous Percoll gradients are prepared by adding 2 ml each of 23%, 15%, 10% and 3% Percoll (v/v) in gradient buffer [0.32 M sucrose containing 1 mM ethylenediaminetetraacetic acid (EDTA) and 0.25 mM dithiothreitol (DTT), pH 7.4] to 10-ml rimmed polycarbonate tubes (EDTA may be omitted if a 1 mM concentration inhibits the enzyme of interest). The Percoll-sucrose solutions are adjusted to pH 7.4 with HCI or NaOH before use and the gradients layered with fine tubing inserted into 1-mm i.d. tubing connected to a peristaltic pump to deliver Percoll with minimum disruption

[20]

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E N D O P E P T I D A S E 24.11

Homogenize tissue (1 g of brain tissue in 9 ml of gradient buffer: 10 up-

and-down strokes at 700 rpm) 1000g for 10 min at 4~

Supernatant (Sl) (adjust to 14 ml with gradient buffer protein concentration approximately 5 mg/ml)

l

Layer 2 ml of S1 onto each of seven Percoll gradients

32,000 g for 5 min at 4oc

(excluding acceleration and deceleration time) Collect and pool synaptosomes at interface 4 (Fractions numbered from top to bottom)

Wash fractions twice with Krebs buffer, pH 7.4

15,000 g for 15 min at 4oc

Resuspend fractions gently in 1 ml of Krebs buffer, pH 7.4 (Restore metal ions by adding MnC12 to final concentration of 1 mM)

SCHEME I

Flow chart for preparation of synaptosomes.

of interfaces. For all centrifugation steps a Sorvall RC-5B plus SS34 rotor is used. Scheme I is a flow chart indicating the fractionation steps. An important feature of this technique is that it separates synaptic plasma membranes (located primarily in the less dense fractions F1 and F2) from intact synaptosomes (mainly fractions F3 and F4), as revealed by the distinct distributions of the postsynaptic muscarinic receptors ([3H]QNB-binding sites) from the presynaptic [3H]nicotine-binding sites (15). The ability of this technique to resolve pre- and postsynaptic elements can be applied to examine the perisynaptic localization of neuropeptidases. By using this procedure, for example, we have been able to demonstrate that E-24.11 is present on

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E X T R A C E L L U L A R PROCESSING ENZYMES IN THE CNS

both pre- and postsynaptic membranes in the pig substantia nigra whereas neuronal angiotensin-converting enzyme (ACE) is confined to dendritic membranes (16). This fractionation procedure can therefore provide useful biochemical support for alternative localization techniques such as immunocytochemistry (see below). Figure 1 compares the distribution of E-24.11 and angiotensin-converting enzyme among the Percoll gradient fractions after fractionation of pig striatal tissue.

Distribution of Membrane Peptidases in Central Nervous System Using Immunohistochemical Techniques Immunohistochemical methods offer the possibility of a precise localization of membrane peptidases in the nervous system. The principal difficulties when using this approach arise from the relatively small amounts of the cell surface enzymes in the nervous system, particularly the neuropil-associated fractions. Modification of the well-documented immunostaining methods that revolutionized the identification of peptide-containing neurons (17, 18) has proved a useful starting point for central nervous system (CNS) peptidase location (5, 19). In this section, attention is focused on criteria necessary for successful microscopic and ultrastructural location of membrane peptidases, with particular reference to E-24.11. Dual labeling of peptides and peptidases is also considered.

Antibody Specificity When analyzing the distribution of antigen in the nervous system, the specificity of the primary antibodies is of paramount importance. The presence of contaminating antibodies may be due either to unwanted immunoglobulins present in the serum before immunization or to impurities in the immunogen used to generate the antiserum. The specificity of the antibody may be improved, for example, by using a pure antigen and by affinity purifying the antisera. An alternative approach is to develop a monoclonal antibody but this can, in itself, be a lengthy procedure (20). The advantage of using monoclonal antibodies is that the nonspecific background is often much less than that produced by polyclonal antisera. Furthermore, once developed, the homogeneous clones secreting a single antibody may be stored, thus providing a readily available source of antibodies. One disadvantage of monoclonal antibodies is that in tissues where the antigen of interest is present in low

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FIG. 1 Fractionation of pig brain striatal tissue on a Percoll gradient. See text for details. S 1, Crude synaptosome fraction that was layered on the discontinuous gradient; F1-F4, fractions at interfaces; F5, the pellet. Bars indicate standard error of the mean. (A) Binding of [3H]nicotine (three experiments). (B) Binding of [3H]QNB (four experiments). Both ligands expressed as picomoles per milligram of protein. (C) E-24.11 activity expressed as picomoles per minute per milligram of protein (five experiments). (D) ACE activity expressed as picomoles per minute per milligram of protein (four experiments). Significant differences, by t test, were as follows: nicotine, p < 0.01 for F4 vs S1 or F1; QNB, p < 0.01 for F5 vs F1 or S1 and F4 vs F1;p < 0.05 for F4 vs S1; E-24.11, p < 0.01 for F4 vs F2; ACE, p < 0.05 for F1 vs F4 or F5. [Reproduced from Barnes et al. (16) with permission of Raven Press, Ltd., New York.]

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111 EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

amounts, the immunostaining signal may be weak. For example, the mouse monoclonal antibody to E-24.11, GK7C2, gives satisfactory staining results with sections of kidney and intestine, but not in brain (19), where the antigen is several orders of magnitude less abundant. Monoclonal antibodies (MAbs) may be useful, however, for isolation of highly pure antigen that can then be used to generate more specific rabbit polyclonal antibodies. For this procedure the monoclonal antibody is immobilized on Sepharose 4B, which is then used as an immunoaffinity resin for purification of the antigen. Elution of the antigen can usually be effected by change of pH or by use of chaotropic reagents. High purification factors can be achieved in such procedures. For example, the first purification of brain E-24.11 by this methodology resulted in a 5000-fold increase in specific activity over the solubilized membrane fraction (21). Immunoaffinity-purified antigen can then be used to generate monospecific polyclonal antiserum that can be purified on a column of Sepharose-4B to which the pure antigen has been attached. Affinity-purifiedrabbit polyclonal antibodies to the membrane peptidases E-24.11, aminopeptidase N, and aminopeptidase W have all been generated in this way by making use of appropriate monoclonal antibodies (22, 23). Specific inhibitors can also be useful for the affinity purification of peptidases. For example, lisinopril-Sepharose is highly effective for the purification of angiotensin-converting enzyme (24) and cilastatin-Sepharose has been used for one-step purification of membrane dipeptidase (25). Further characterization of any newly isolated antibody should include Western blotting, ideally against proteins from the tissue under study. When using the antibody for immunocytochemistry its specificity may be further checked by substituting the primary antibody for preimmune immunoglobulin or by preabsorption of the antibody with an excess of the antigen for 30 min at room temperature followed by centrifugation (1000 g for 10 rnin). In both cases staining should be abolished if the antibody is specific.

Fixatives For localization of membrane peptidases, maintenance of immunoreactivity and morphology has been achieved by using buffered fixatives based on aldehydes (18, 26). Routinely, for light microscopical studies we use 4% (w/v) paraformaldehyde applied to thawed cryosections for 20 min or introduced by perfusion. Paraformaldehyde is a monoaldehyde that has the advantage of penetrating tissues rapidly, but forming cross-links more slowly. For electron microscopy higher concentrations of paraformaldehyde (up to 8%) or the addition of glutaraldehyde (0.05-0.1%) to 4% paraformaldehyde is essential for good

[20] ENDOPEPTIDASE 24.11

353

ultrastructural preservation. Glutaraldehyde is a dialdehyde that rapidly cross-links proteins but penetrates the tissue more slowly than the monoaldehyde. High concentrations of glutaraldehyde (>0.1%) and long postfixation periods (> 1 hr) should be avoided as these have deleterious effects on the antigenicity of membrane peptidases. For example, it is not possible to reveal E-24.11 immunocytochemically if postfixation times of over 1 hr are used (27, 28). Various strategies can be adopted to minimize fixation damage. 1. Introduction of the dialdehyde perfusate (glutaraldehyde) can be followed by perfusion with the monoaldehyde (paraformaldehyde) or phosphate-buffered saline (PBS) (150 mM NaC1; 10 mM NaH2PO4 92H20, pH 7.4), or by 0.02% glycine, which reacts with the remaining aldehyde groups, thus preventing further cross-linking. 2. The sections may be quenched with 1% sodium borohydride to reduce aldehyde groups, thus suppressing nonspecific background (29). [Sodium borohydride (1%) is stable at high pH and may be stored at pH 12.0; immediately prior to use, the pH is lowered to pH 8.0.] 3. Glycine (0.02%, w/v) may be included in the incubation medium (store the glycine at 4~

Preparation of Tissues Cryostat Sections Immunocytochemical mapping of peptidase antigens throughout the brain reveals any heterogeneous expression and allows comparisons to be made with other peptidases and peptides. For example, the general similarity of the distribution of the two brain hydrolases, acetylcholinesterase and endopeptidase 24.11, was identified by such mapping (5, 19). The simplest and most effective way of obtaining good light microscopical immunostaining for the membrane peptidases is to freeze fresh brain tissue and cut cryostat sections. To prevent the brain slices or whole hemispheres from shattering, the tissue is immersed in 3% (w/v) low-viscosity carboxymethyl cellulose in water poured into a suitable mold (we use a cork base and tape) before freezing in isopentane cooled with liquid nitrogen, or in solid carbon dioxide. The prepared tissues may be stored in liquid nitrogen or in a -70 ~ freezer. Cryostat sections of fresh CNS tissue may be thawmounted directly onto untreated coverslips or slides. Vibratome Sections Vibratome sections (Campden Instruments, London, England) allow the correlation of light and electron microscopical studies of membrane pepti-

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

dases; our procedures are based on those described by Priestley and Cuello (17) for peptides. Sections (40-60/zm thick) are cut from blocks of perfused fixed tissue and immunostained as free-floating sections. After immunostaining, the sections may be processed for electron microscopy and flat embedded in Durkupan resin (FSA Laboratory Supplies, Loughborough, England) between a microscope slide and plastic coverslip. For viewing under the light microscope, the coverslip may be removed after curing the resin. Areas immunostaining for the peptidases are excised with a scalpel and remounted onto preformed cones of resin, using Superglue. Ultrathin sections are cut and collected on electron microscope grids. Penetration of Reagents One major difficulty when immunostaining Vibratome sections is that uneven reagent penetration may lead to failure to detect an antigen. The treatment of tissues with detergents (e.g., 0.1-1.0% Triton X-100 in phosphate-buffered saline) after fixation for 10-20 min, while suitable for light microscopy, causes excessive damage to ultrastructure. The most successful pretreatment for enhancing peptidase staining without excessive tissue deterioration involves the freeze/thawing of sections as described by Bolam et al. (30). Sections are immersed in a cryoprotectant (25% sucrose, 10% glycerol in 0.05 M sodium phosphate buffer, pH 7.4) in vials for approximately 10 min, placed on nylon mesh nets (Lockertex; Henry Simon, Ltd., Warrington, Cheshire, England) frozen in isopentane cooled by liquid Nz, and then plunged into liquid N2, prior to thawing in 0.1 M sodium phosphate buffer, pH 7.4.

Immunostaining Techniques Immunop e roxidas e By using polyclonal antibodies coupled with a biotin-streptavidin-peroxidase system to amplify the signal, it has been possible to map membrane peptidases in the CNS (5, 19). Figure 2 illustrates the distribution of E-24.11 in the caudate putamen and striatonigral pathway stained with immunoperoxidase. At the electron microscopical level the tendency of the product to diffuse from its oxidation site and to be absorbed onto adjoining membranes makes precise localization of the antigen to a specific membrane difficult. For example, there is general agreement that endopeptidase 24.11 is present on plasma membranes of axons and dendrites in the neuropil (Fig. 3a) (6, 27), but the reported staining of glial cell membranes (27) has not been unambiguously observed (6, 16).

[20]

355

E N D O P E P T I D A S E 24.11

#~

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FIG. 2 Parasagittal section of fetal pig brain stained for E-24.11. The 25-/zm section was cut from the medial surface of the block. E-24.11 staining is continuous throughout the striatonigral pathway, including the caudate putamen (CP), entopeduncular nucleus (EP), and substantia nigra (SN). Note also the intense staining of the ventral pallidum (VP) and olfactory tubercle (OT). IC, Internal capsule; T, thalamus. Bar: 2.5/zm.

Procedure: Peroxidase Staining This procedure adapted from Matsas et al. (19) is suitable for cryostat sections mounted on coverslips or slides or for free-floating Vibratome sections. Except where stated all incubations are at room temperature. This schedule is as used for a rabbit polyclonal antibody to E-24.11, but may be readily adapted for other antibodies, either polyclonal or monoclonal. 1. Either dissect flesh brain and prepare blocks for cryostat (see above) or perfuse the animal with appropriate fixative (see above), dissect brain, and postfix for up to 1 hr. 2. Cryostat blocks: freeze/thaw cut sections (8-15/zm) onto coverslips or slides. Fix for 20 rain with 4% paraformaldehyde. Vibratome blocks: Stick small blocks of tissue to chuck with Superglue. Cut sections (40-60/zm) and transfer to Tris-buffered saline (50 mM Tris; 150 mM NaCI, pH 7.4) (TBS) in scintillation vials, using a paint brush.

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

3. Wash the fixed sections in TBS: three changes in 15 min. 4. Block nonspecific protein binding sites with 0.2% (w/v) gelatin in TBS for 30 min. 5. Block endogenous peroxidase activity with 0.01% (w/v) phenylhydrazinium chloride in TBS at 37~ for 30 min. 6. Rinse in 0.2% gelatin-TBS. 7. Incubate in primary antibody (e.g., rabbit anti-pig E24.11, diluted 1:100 at room temperature for 2 hr or at 4~ overnight). All antibodies are diluted in 0.2% gelatin-TBS plus 1% normal serum (heat-inactivated serum of species in which secondary antibody was raised). 8. Wash in TBS: three changes in 10 min. 9. Incubate in donkey anti-rabbit biotinylated species-specific antibody (diluted 1:50 in 0.2% gelatin-TBS) for 1 hr. 10. Wash in TBS: three changes in 10 min. 11. Incubate in streptavidin-biotinylated horseradish peroxidase complex (diluted 1:400 in 0.2% gelatin-TBS) for 1 hr. 12. Wash in TBS" three changes in 10 min. 13. For light microscopy: Incubate the sections with 0.02% (w/v) 3-amino3-carbazole in 5% (v/v) N,N-dimethylformamide in 50 mM sodium acetate buffer, pH 5.0, and then add 0.01% H202 . For electron microscopy, diaminobenzidine (DAB) is substituted for 3-amino-9-ethylcarbazole. The DAB (Amersham, Arlington Heights, IL) is best stored in 1-ml aliquots in 0.1 M phosphate buffer, pH 7.2, at -20~ (2.5%, w/v). Dilute a 1-ml aliquot to 50 ml with 0.1 M phosphate buffer and add 15/A of H202 immediately before use. 14. Stain for 5-20 min (optimize for individual system) and stop the reaction with distilled water. 15. Mount coverslips for light microscopy in 95% glycerol in phosphatebuffered saline (PBS). For electron microscopy transfer sections to 0.1 M sodium phosphate buffer, pH 7.4.

lmmunogold Labeling Immunolabeling of ultrathin resin sections with gold-labeled probes, introduced by Roth and Binder (31), has been widely used in immunocytochemistry of the nervous system. Gold labels have a number of advantages over the peroxidase technique and are particularly useful when labeling membrane peptides. The discrete gold particles are small and locate the antigen site more accurately (compare Fig. 3a and b). Gold particles of differing sizes may be used to detect two or more antigens, making them potentially useful for locating peptidases and their putative substrates. We have employed a preembedding protocol (32) that utilizes 1-nm gold-labeled immunoglobulins followed by a silver enhancement to localize E-24.11 (28). Silver enhancement provides a means by which the 1-nm colloidal gold particles can be

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FIG. 3 Porcine brain sections immunostained for E-24.11 by using peroxidase and gold conjugates. The ultrathin section in (a) is of globus pallidus, showing immunoperoxidase staining (arrows), whereas in (b) longitudinally oriented axonal membranes from substantia nigra are decorated with immunogold that has been silver enhanced (arrows). B1, B2, Boutons; D, dendrite; A, axons. Bars: 0.5 #~m. enlarged by precipitation of metallic silver. The resulting high-contrast, dark brown to black signal is remarkably similar in appearance to the diaminobenzidine product for horseradish peroxidase at the light microscopic level, but may be seen as discrete particles at the electron microscopic level (Figs. 3b and 4a and b). A more detailed discussion of the silver intensification technique may be found in Chan et al. (32). The penetration of gold probes into Vibratome sections is limited, however, to the surface layers. The improved procedures for the production of ultrathin cryosections (33, 34) offer an alternative method for locating peptidases. Primary antibodies followed by gold-labeled protein A or immunoglobulins are applied to the frozen sections (see Fig. 5). E-24.11 (28), aminopeptidase N (35), and angiotensin-converting enzyme (K. Barnes and A. J. Turner, unpublished observations, 1994) have all been localized by this technique. Procedure: Immunogold Silver Enhancement This procedure is modified from Chan et al. (32), using the Amersham 1-nm SE kit. All reagents are made up in "Millipore water" and incubations are at room temperature except where stated.

358

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

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FIG. 4 Double labeling of the substantia nigra for E.24-11 and SP. Both sections (a) and (b) were immunostained for SP (peroxidase) and E-24.11 (immunogold). Colocalization of the two antigens on boutons is indicated by double arrows. Silver enhancement for 8 min. Bars: 0.3/~m.

Washing buffer: 0.8% (w/v) BSA and 0.1% (w/v) immunogold silver staining (IGSS) gelatin in PBS-azide, pH 7.4. Make up 100 ml for stock solution Blocking buffer: To the stock washing buffer add 5% (v/v) normal goat serum Incubation buffer: To the stock washing buffer add 1% (v/v) normal goat serum 1. Collect Vibratome sections in TBS in scintillation vials. 2. Place sections in 1% sodium borohydride in 0.1 M phosphate buffer, pH 8.0, for 30 min. 3. Rinse the sections thoroughly with 0.1 M phosphate buffer, pH 7.4, or until bubbles disappear. 4. Immerse in blocking buffer for 30 min. 5. Incubate in primary antibody diluted in "incubation buffer" overnight at 4~ 6. Wash in washing buffer: three changes in 15 min.

[20] ENDOPEPTIDASE 24.11

359

Fic. 5 Ultrathin cryosections of pig substantia nigra double-labeled for E-24.11 and SP using immunogold. (a) E-24.11, 15-nm gold; SP, 5-nm gold; (b) E-24.11, 5-nm gold; NC1, 15-nm gold. Straight arrows indicate membranes labeled for E-24.11 and curved arrows dense-core vesicles labeled for SP. B, Bouton. Bars; (a) 0.3 /xm; (b) 0.2/xm. 7. Incubate in secondary gold-labeled IgG diluted 1 : 50 in incubation buffer for 2 hr. 8. Wash in washing buffer: three changes in 15 min followed by one change into 0.1 M phosphate buffer, pH 7.4. 9. Fix the sections in 2% (v/v) glutaraldehyde in 0.1 M phosphate buffer, pH 7.4 (to prevent uncoupling of gold from IgG). 10. Rinse in 0.1 M phosphate buffer, followed by three brief washes in deionized water. 11. Follow the silver amplification procedure according to the instructions on the Intense SE kit. (Using ice-cold solutions an intensification time of 6-8 min is suitable for both light and electron microscopy.) 12. Rinse quickly in deionized water (three changes) and return sections to 0.1 M phosphate buffer, pH 7.4. 13. Process sections for electron microscopy (17).

Dual Localization of Antigens The colocalization of membrane peptidases and their possible target neuropeptides is important as it may offer some clues as to the physiological roles of the enzymes. One successful method combines an immunogold and

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E X T R A C E L L U L A R P R O C E S S I N G E N Z Y M E S IN THE CNS

immunoperoxidase technique on Vibratome sections, and the other uses two differing sizes of immunogold on ultracryosections (see Figs. 4 and 5). Because preparing and sectioning Vibratome sections is technically less demanding than routinely obtaining good cryosections from CNS tissue, only a detailed protocol of the immunogold and peroxidase protocol is given here.

Procedure: Dual Immunogold and Immunoperoxidase For this procedure, the primary antibodies must be raised in different species. In Fig. 4, for example, a monoclonal antibody, NC 1 to SP (Accurate Chemical and Scientific Corporation, Westbury, NY) (peroxidase), raised in rats, was used simultaneously with a rabbit polyclonal to E-24.11, RP123 (immunogold). 1. Follow steps 1-4 as in the immunogold procedure. 2. Block endogenous peroxidases as in step 5 of the immunoperoxidase procedure. 3. Follow steps 5 and 6 as in the immunogold procedure, but add both primary antibodies simultaneously. 4. Incubate with biotinylated antibody as in step 9 of the immunoperoxidase procedure. 5. Wash using washing buffer (without azide, which inhibits the peroxidase activity). 6. Incubate with streptavidin as in step 11 of the immunoperoxidase procedure. 7. Follow steps 12-14 of the immunoperoxidase procedure for development of color using DAB. 8. Rinse in washing buffer, then follow steps 6-13 of the immunogold procedure.

Conclusions The widely differing techniques of phase separation with Triton X-114, subcellular fractionation, and immunocytochemistry may all be used to help establish the topology, roles, and distribution of membrane peptidases. In particular, E-24.11 has been established as an integral membrane protein and ectoenzyme. The localization of E-24.11 on neuronal membranes, and its colocalization on the plasma membrane of substance P-rich boutons in pig substantia nigra, endorse its probable role as a neuropeptidase. Its cooccurrence with other peptides, for example, enkephalins in the globus pallidus, has been established at the light microscope level, but as yet remains to be confirmed by dual labeling at the ultrastructural level.

[20] ENDOPEPTIDASE 24.11

361

Acknowledgments We thank the Medical Research Council and the Wellcome Trust for support of our work.

References N. M. Hooper and A. J. Turner, FEBS Lett. 229, 340 (1988). 2. A. J. Turner, N. M. Hooper, and A. J. Kenny, in "Neuropeptides: A Methodology" (G. Fink and A. J. Harmar, eds.), p. 189. Wiley, Chichester, 1989. A. J. Turner, in "Neuropeptides and Their Peptidases" (A. J. Turner, ed.), p. 183. Ellis Horwood, Chichester, 1987. S. Howell, H. Murray, C. S. Scott, A. J. Turner, and A. J. Kenny, Biochem. J. 278, 417 (1991). K. Barnes, R. Matsas, N. M. Hooper, A. J. Turner, and A. J. Kenny, Neuroscience 27, 799 (1988). K. Barnes, A. J. Turner, and A. J. Kenny, Neurosci. Lett. 94, 64 (1988). 7. A. J. Turner, Adv. Neuroimmunol. 3, 163-170 (1993). 8. N. M. Hooper, M. G. Low, and A. J. Turner, Biochem. J. 244, 465 (1987). 9. A. W. Scotto and D. Zakin, Biochemistry 24, 4006 (1985). 10. N. M. Hooper and A. J. Turner, Biochem. J. 261, 811 (1989). 11. C. Bordier, J. Biol. Chem. 256, 1604 (1981). 12. N. M. Hooper and A. Bashir, Biochem. J. 280, 745 (1991). 13. M. K. Johnson and V. P. Whittaker, Biochem. J. 88, 404 (1963). 14. P. R. Dunkley, P. E. Jarvie, J. W. Heath, G. J. Kidd, and J. A. Rostas, Brain Res. 372, 115 (1986). 15. B. Thorne, S. Wonnacott, and P. R. Dunkley, J. Neurochem. 56, 479 (1991). 16. K. Barnes, A. J. Turner, and A. J. Kenny, J. Neurochem. 58, 2088 (1992). 17. J. V. Priestley and A. C. Cuello, IBRO Handb. Ser. 3, 273 (1983). 18. L. A. Sternberger, "Immunocytochemistry," 3rd ed. Wiley, New York, 1985. 19. R. Matsas, A. J. Kenny, and A. J. Turner, Neuroscience 18, 991 (1986). 20. A. C. Cuello, C. Milstein, and G. Galfr6, IBRO Handb. Ser. 3, 215 (1983). 21. J. M. Relton, N. S. Gee, R. Matsas, A. J. Turner, and A. J. Kenny, Biochem. J. 215, 519 (1983). 22. N. S. Gee, R. Matsas, and A. J. Kenny, Biochem. J. 214, 377 (1983). 23. M. A. Bowes and A. J. Kenny, Immunology 60, 247 (1987). 24. N. M. Hooper and A. J. Turner, Biochem. J. 241, 625 (1987). 25. G. M. Littlewood, N. M. Hooper, and A. J. Turner, Biochem. J. 257, 361 (1989). 26. P. Somogyi and H. Takagi, Neuroscience 7, 1779 (1982). 27. D. Marcel, H. Pollard, P. Verroust, J. C. Schwartz, and A. Beaudet, J. Neurosci. 10, 2804 (1990). 28. K. Barnes, A. J. Turner, and A. J. Kenny, Neuroscience 53, 1073 (1993). 29. T. Kosaka, I. Nagatsu, J.-Y. Wu, and K. Harma, Neuroscience 18, 975 (1986). 30. J. P. Bolam, C. M. Francis, and Z. Henderson, Neuroscience 41, 483 (1991). o

,

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III EXTRACELLULARPROCESSING ENZYMES IN THE CNS 31. J. Roth and M. Binder, J. Histochem. Cytochem. 26, 163 (1978). 32. J. Chan, C. Aoki, and V. M. Pickel, J. Neurosci. Methods 33, 113 (1990). 33. G. Griffiths, A. McDowell, R. Back, and J. Dubochet, J. Ultrastruct. Res. 89, 65 (1984). 34. K. T. Tokuyasu, Histochem. J. 21, 163 (1989). 35. K. Barnes, A. J. Kenny, and A. J. Turner, Eur. J. Neurosci. 6, 531 (1994).

[21]

Identification and Distribution of Endopeptidase 24.16 in Central Nervous System F. Checler, P. Dauch, H. Barelli, V. Dive, Y. Masuo, B. Vincent, and J. P. Vincent

Introduction Endopeptidase 24.16 was first detected during the course of the elucidation of the mechanisms by which the 13-residue peptide neurotensin was inactivated by purified rat brain synaptic membranes (1). This work established that the primary cleavages took place at the Arg8-Arg9, Prol~ 1~, and Tyr ~Ile ~2peptidyl bonds, leading to the biologically inactive N-terminal fragments neurotensin(1-8), neurotensin(1-10), and neurotensin(1-11) (1). By means of specific peptidase inhibitors commercially available or provided by other groups, we demonstrated that endopeptidases 24.15 and 24.11 were responsible for the formation of neurotensin(1-8) and neurotensin(1-11), respectively, and that endopeptidase 24.11 also partly contributed to neurotensin(I-10) production (2, 3). However, the fact that the major formation of neurotensin(1-10) totally resisted inhibition by peptidase inhibitors was the first clue to the presence of an unknown proteolytic activity. It is important to note that the starting material, that is purified rat brain synaptic membranes extensively washed by a hypotonic buffer (5 mM Tris-HC1, pH 7.5), allowed us to eliminate abundant soluble proteases able to generate neurotensin(1-10) such as proline endopeptidase and carboxypeptidase A, which would have hindered the detection of endopeptidase 24.16. This enzyme, which was first referred to as "neurotensin-degrading neutral metalloendopeptidase" with respect to its sensitivity to pH and metal chelators, was purified from rat brain synaptic membranes (4) and later from peripheral tissue sources such as rat ileum and kidney (5, 6). The physicochemical and biochemical characteristics of the peptidase confirmed its novelty. According to the 1992 IUBMB Enzyme Nomenclature (7), this enzyme is listed as neurolysin (EC 3.4.24.16). Several lines of evidence suggested that endopeptidase 24.16 could be of importance for the physiological termination of the neurotensinergic message. Thus, we showed that neurotensin analogs in which Tyr ~ was replaced by an aromatic D-amino acid totally resisted proteolysis by rat brain synaptic membranes and endopeptidase 24.16, in vitro as well as in vivo after intracerebroventricular administration in the rat (8). This was interesting with respect to the fact that such modified analogs, although poor agonists of neurotensin Methods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

receptors in in vitro binding experiments [100- to 1000-fold lower affinity when compared to neurotensin (9)], behaved as a 10-fold more potent hypothermiant agent than the parent peptide in the rat (10-12). These data, which suggested that the D-amino acid substitution had conferred resistance to degradation, confirmed the importance of the Prol~ 11 peptide bond as a target of physiological inactivating peptidases. The possibility that endopeptidase 24.16 could fulfill this role was further reinforced by the observation that neurotensin(1-10) and its C-terminal counterpart, neurotensin(ll-13), were the only two degradation products that were consistently detected when we studied neurotensin degradation by membrane preparations or cell lines from various neural or extraneural origins (Table I). To improve the investigation of the biochemical and physicochemical aspects of endopeptidase 24.16 and to assess directly the hypothesis of its implication in neurotensin degradation, it was necessary to develop specific tools directed toward the enzyme. This chapter describes the characteristics of a fluorimetric substrate, specific inhibitors, and a polyclonal antibody developed against rat brain endopeptidase 24.16. Furthermore, we describe advances concerning the distribution of the enzyme in the central nervous system (CNS).

H i g h - P e r f o r m a n c e L i q u i d C h r o m a t o g r a p h y of N e u r o t e n s i n a n d Its S y n t h e t i c Partial S e q u e n c e s High-performance liquid chromatography (HPLC) is carried out on a Waters Associates apparatus equipped with a model 481 detector, two model 510 pumps, a WISP injector, and an automated gradient controller. Elutions are carried out at room temperature onto a RP18 Lichrosorb column (Merck, Rahway, NJ), at a flow rate of 1 ml/min and absorbance is monitored at 230 nm with a detector setting of 0.1 full scale. Samples are chromatographed using two different buffer/solvent systems. System 1: 35-min nonlinear gradient (gradient 7) of 20 mM ammonium acetate (pH 6.4)-acetonitrile from 90:10 (v/v) to 65:35 (v/v). The retention times of neurotensin and a series of neurotensin synthetic fragments are given in Table II and a typical chromatogram is shown in Fig. 1. System 2: 42-min linear gradient of 0.1% (v/v) trifluoroacetic acid (TFA), 0.05% (v/v) triethylamine (TEA)-0.1% (v/v) TFA, 0.05% (v/v) TEA in acetonitrile from 90" 10 to 60" 40. The retention times of neurotensin and synthetic peptides are given in Table II.

TABLEI Neurotensin" Metabolites Derived from Primary Inactivating Cleavagesb Metabolite Tissue or cells Central Synaptic membranes Primary cultured neurons Glial cells Neuroblastoma cells N l E l 15 Peripheral Rat fundus plasma membranes Circular smooth muscle plasma membranes Dog ileum myenteric plexus membranes Dog ileum longitudinal smooth muscle plasma membranes Dog ileum deep muscular plexus membranes Dog ileum submucous plexus membranes HT29 cells

NT (1-8)

NT (1-10)

NT (11-13)

NT (1-11)

NT (9-13)

+ + + +

+ + + +

+ + + +

+

+ + + +

+ +

+ + +

-

-

+ + + +

+ + + +

+ + +

+ + +

+ + +

NT (1-7)

-

+ + + -

-

+

" Neurotensin sequence: pGlu-Leu-Tyr-Glu-Asn-Lys-Pro-Arg-Arg-Pro-Tyr-Ile-Leu. In various tissues or cells.

-

-

+ + +

+ + -

+ + + -

+ + -

NT (1-12)

-

+ + + + + -

366

III EXTRACELLULAR PROCESSING ENZYMES IN THE CNS TABLE II

Elution Times of Neurotensin Partial Sequences after HPLC Analysis Elution times (min)

Sequence NT NT NT NT NT NT NT NT

Z

c

System 2

12 15 19 23 20 26.5 27 31.5

14 16.4 18.8 22.4 25.6 27.2 27.5 32.4

(1-7) (1-8) (1-10) (1-11) (11-13) (9-13) (1-12)

r

o

System 1

is. ,T. i!

ZZ

ZZ

ZZ

Z

9

T I M E , MIN

FIG. 1 HPLC separation of synthetic neurotensin fragments. About 1-2 nmol of each peptide was separately injected onto an RP18 Lichrosorb column and eluted according to gradient system 1.

[21] ENDOPEPTIDASE 24.16 IN CNS

367

Tritiated neurotensin is the most useful radiolabeled substrate for neurotensin degradation studies. Tritiated neurotensin {[3,11-tyrosyl-3,5-3H(N)]neuro tensin} can be obtained from New England Nuclear (Boston, MA) at a specific radioactivity of about 50-100 Ci/mmol. This peptide, which is labeled on both Tyr-3 and Tyr-11 of the neurotensin sequence, allows the detection of most of the N- and C-terminal degradation products, even when low quantities render them difficult to detect by optical density. The retention times of the tritiated degradation products are identical to those of their unlabeled counterparts in both buffer/solvent systems. Furthermore, free tritiated tyrosine could be detected and eluted with a retention time of about 5.8 min in HPLC system 1.

Hydrolysis of Mcc-Pro-Leu-Gly-Pro-o-Lys-Dnp Mcc-Pro-Leu-Gly-Pro-o-Lys-Dnp (QFS; Novabiochem, Meudon, France) was first reported to behave as a substrate of Pz-peptidase, a thiol-metalloendopeptidase widely distributed within mammalian tissues (13). We have shown that this quenched fluorimetric substrate is cleaved at the Leu-Gly peptidyl bond by endopeptidase 24.16, leading to the formation of the fluorescent product Mcc-Pro-Leu, and could be used to monitor this activity routinely (14).

Fluorimetric Assay QFS (50/zM) is incubated at 37~ with the enzymatic source in a final volume of 100/zl of 20 mM Tris-HCl, pH 7.5. Incubations are stopped by the addition of 2.5 ml of 80 mM sodium formate, pH 3.7. Activity is monitored by a fluorimeter, setting 345 nm and 405 nm as excitation and emission wavelengths, respectively. Quantitation of the Mcc-Pro-Leu release is deduced from a standard curve established with known amounts of synthetic MccPro-Leu. Figure 2A illustrates the initial velocity measurements of QFS hydrolysis by purified endopeptidase 24.16 according to the substrate concentration. The corresponding Lineweaver-Burk plot indicates a Km value of about 25/zM. Kinetic analysis carried out at a saturating concentration of QFS (i.e., under maximal velocity conditions) indicates that the degradation rate can be considered as initial within the first 60 min (Fig. 2B). The specific activity of purified rat brain endopeptidase 24.16 toward QFS is about 215 nmol/hr of protein.

368

III EXTRACELLULAR PROCESSING ENZYMES IN THE CNS A "7

2.5

g o _J

2

~ n_

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X

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J=

J= 1"51

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_ ~

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o o

0 20 40 60 1/[OF02 ], mM "1

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100

50

150

[OF02],I~M

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250

B 2.5

*, a_

:i '~

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1

o E e~ 0.5

-

0

E 5 15 30

60

120

Time,min

180

FIG. 2 Kinetic parameters of QFS (QFO2) hydrolysis by purified rat brain endopeptidase 24.16. QFS was incubated at 37~ with purified endopeptidase 24.16. Hydrolysis was fluorimetrically recorded and quantified as described in text. (A) Initial velocity as a function of substrate concentration. Inset: Lineweaver-Burk plot of the data. (B) Mcc-Pro-Leu formation as a function of time at a 100/~M concentration of QFS.

HPLC Assay It is always important to confirm the site of cleavage of QFS by HPLC analysis. QFS (4 nmol, 20/~M) is incubated at 37~ in a final volume of 200 /A of 20 mM Tris-HC1, pH 7.5. Acidified samples are analyzed in the buffer/ solvent system 2 according to two distinct gradients.

369

[21] ENDOPEPTIDASE 24.16 IN CNS Mcc-Pro-Leu

f

r _.1

6

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rut) u.

o

6 f,.)

o r

QFS

)

J

_/ I

0

0

' ' '2'5

b

2

I

40

I

Time (rain)

FIG. 3 QFS hydrolysis by endopeptidase 24.16: HPLC analysis according to two solvent/buffer systems. QFS (4-5 nmol) was incubated with purified endopeptidase 24.16 and HPLC analysis was performed according to gradients 1 (left-hand side) and 2 (right-hand side).

Gradient l" 42-min linear gradient of 0.1% TFA, 0.05% TEA-0.1% TFA, 0.05% TEA in acetonitrile from 75"25 to 20"80. Gradient 2" Biphasic gradient of 0.1% TFA, 0.05% TEA-0.1% TFA, 0.05% TEA in acetonitrile from 90" 10 to 75"25 in 15 min, then from 75" 25 to 20"80 in 40 min. Figure 3 illustrates the HPLC of QFS hydrolysis by purified endopeptidase 24.16 after analysis with gradient 1 (left-hand side) and gradient 2 (righthand side).

I n i b i t o r s of E n d o p e p t i d a s e 24.16

Dipeptides Previously we showed that several dipeptides that mimic the Pro-X bond of neurotensin hydrolyzed by endopeptidase 24.16 could inhibit the peptidase (5, 15). Table III indicates that the rank of order of dipeptides potencies as well as the KI values appeared similar with both neurotensin and QFS substrates. Pro-Ile was the most potent blocker of endopeptidase 24.16 (Table

370

III

E X T R A C E L L U L A R PROCESSING E N Z Y M E S IN THE CNS TABLE I I I

E f f e c t o f D i p e p t i d e s on R a t E n d o p e p t i d a s e 24.16 a KI values on endopeptidase 24.16 (/zM) Dipeptide

Ileum b

Brain b

Brain c

Pro-Ile Pro-Leu Pro-Tyr Pro-X (X = Ala, Ser, Gly) X-Pro (X = Ser, Pro, Val, His, Met) Val-Tyr Trp-Tyr X-Tyr (X = Ala, Ser, Lys, Gly, His)

50 190 200 ~>1000 >>1000 380 460 -> 1000

46 130 100 -> 1000 ~>1000 200 290 -> 1000

89 296 469 -> 1000 ~>1000 734 947 -> 1000

a From central and peripheral origin. b K1 values were established with tritiated neurotensin as substrate. r KI values were established with QFS as substrate.

III). A 10 - 2 M concentration of Pro-Ile drastically reduced QFS and neurotensin hydrolyses (Fig. 4A and B, respectively). Complete dose-response curves revealed IC50 values of 160/xM for QFS (Fig. 4C) and 1 mM for neurotensin (Fig. 4D). According to the relation IC50 = KI(1 + S/Km), we established that the Kt of Pro-Ile toward endopeptidase 24.16 corresponded to 90 ~M whatever the substrate used (16). Interestingly, Pro-lie exerts a selective effect on endopeptidase 24.16 because a 5 mM concentration of the dipeptide did not affect a series of other metallo exo- and endopeptidases (Table IV) from murine source. All X-Pro and X-Tyr dipeptides appeared unable to affect endopeptidase 24.16 (Table III).

Phosphodiepryl 03 Endopeptidase 24.16 belongs to the zinc-containing metallopeptidase family (4, 6). Specific inhibitors developed for this class of peptidases contain a metal coordinating group such as carboalkyls, thiols, hydroxamates, and phospho groups. Concerning the latter, it has been shown that phosphonamide peptides could potently inhibit thermolysin, carboxypeptidase A (17, 18), and bacterial collagenases (19). Interestingly, the structure of one of these peptides, phosphodiepryl 03 [N-(phenylethylphosphonyl)-Gly-L-Pro-aminohexanoic acid], resembled the Gly-Pro-D-Lys-Dnp sequence of the C-terminal product generated after cleavage of QFS by endopeptidase 24.16 and therefore represented a putative inhibitor of this enzyme. Figure 5A

[21] ENDOPEPTIDASE 24.16 IN CNS

371

A

B

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Time, miD

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100

80 75

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60 so 40

0

0

5

4

3

2

"

4

I

3

I

2

-Log ([Olpeptide] / M)

FIG. 4 Effect of Pro-Ile on QFS and neurotensin hydrolysis by endopeptidase 24.16. QFS (50/xM) (A) and neurotensin (20/~M) (B) were incubated with endopeptidase 24.16 in 100/zl of 20 mM Tris-HC1, pH 7.5, in the absence (A and B, left-hand side) or in the presence (A and B, right-hand side) of 10 -2 M Pro-Ile and analyzed by HPLC. The dose-dependent effect of Pro-Ile on QFS (C) or neurotensin (D) hydrolysis was determined after fluorimetric or HPLC procedures, respectively. Values are expressed as the percentage of control obtained in the absence of inhibitor and are the mean _SEM of four to six determinations.

372

III

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS TABLE IV

KI Values of Pro-lie and Phosphodiepryl 03 on Endoand Exopeptidases KI values Peptidase

Pro-lie (/zM)

Phosphodiepryl 03 (nM)

Endopeptidase 24.16 Endopeptidase 24.15 Endopeptidase 24.11 Angiotensin-converting enzyme Carboxypeptidase A Leucine aminopeptidase Trypsin Pyroglutamyl-peptide hydrolase I

90 >5000 >5000 >5000 5000 >5000 > 5000 >5000

0.9 7.5 > 1000 > 1000 > 1000 > 1000 nd a nd

a

nd, Not determined.

shows that 10 -6 M phosphodiepry103 fully abolished neurotensin hydrolysis by purified endopeptidase 24.16. Complete dose-response curves of the inhibition of neurotensin (Fig. 5B) and QFS (Fig. 5C) hydrolysis by phosphodiepryl 03 allowed derivation of K i values of 0.9 and 0.3 nM, respectively (20). As shown in Table IV, phosphodiepryl 03 also inhibited endopeptidase 24.15 (although with a eightfold lower potency), but was unable to affect other zinc-containing enzymes such as endopeptidase 24.11, angiotensin-converting enzyme, carboxypeptidase A, and leucine aminopeptidase.

M e a s u r e m e n t of E n d o p e p t i d a s e 24.16 A c t i v i t y

In Vitro Studies The main pitfall concerning the assay of endopeptidase 24.16 in a complex mixture of enzymatic activities derives from the fact that the enzyme shares some common features with endopeptidase 24.15, although both activities are clearly distinguishable (21). Thus, both enzymes can hydrolyze neurotensin and QFS with similar affinities (Table V) whereas phosphodiepryl 03 exhibits a high potency but limited selectivity toward the two peptidases (Table V). On the other hand, the inhibitor Cpp-Ala-Ala-Tyr-pAB was found to be 50- to 100-fold more potent for endopeptidase 24.15 than for endopeptidase 24.16. According to these parameters, it is possible to estimate the

[21] ENDOPEPTIDASE 24.16 IN CNS

A

t

373

NT

~.1o T T ~i3

1

i o 0

100

/

~ -~

100

5O

0

50

10

i 9

l 8

i 7

6

0

11

I

10

g

8

7

-Log[Inhibitor],M FIG. 5 Effect ofphosphodiepry103 on neurotensin and QFS hydrolysis by endopeptidase 24.16. Neurotensin (2 nmol, 20/zM) was incubated with endopeptidase 24.16 in the absence (A, left-hand side) or in the presence (A, right-hand side) of 1 tzM phosphodiepry103. Complete dose-response curve (B) was carried out after preincubation with various concentrations of phosphodiepry103 and neurotensin degradation was monitored by HPLC. Values are expressed as the percentage of control obtained in the absence of inhibitor. (C) Dose-response curve of the inhibition of QFS hydrolysis by endopeptidase 24.16. Each value was derived from fluorimetric analysis and is expressed as the percentage of control observed without inhibitor.

extent of inhibition of endopeptidase 24.16 or endopeptidase 24.15 when using neurotensin (20 tzM) and QFS (50/zM) as substrates (Table VI) from the following equation" Vi

K m + [S] m

v

[S] + Kin(1 + [I]/K~)

374

III

EXTRACELLULAR PROCESSING ENZYMES IN THE CNS TABLE V

Affinities of S u b s t r a t e s and Inhibitors t o w a r d E n d o p e p t i d a s e s 24.15 and 24.16 Km or KI values (/zM)

Agent

Endopeptidase 24.16

Substrate Neurotensin QFS Inhibitor Pro-Ile Phosphodiepryl 03 Cpp-Ala-Ala-Tyr-pAB

Endopeptidase 24.15

2 25

2 8.6

90 0.9 • 10 -3 1

NI a 7.5 • 10 -3 0.016

a NI, Not inhibited.

where K m and KI correspond to the affinities of substrates and inhibitors given in Table V, [S] and [I] are the concentrations of substrate and inhibitor, and Vi and V are the initial velocity measurements in the presence (Vi) or absence (V) of inhibitor. Examples of such calculations are given in Table VI. It derives from the above statements that the best tool to block endopepti-

TABLE VI

Effect of Inhibitors on N T and Q F S H y d r o l y s i s by E n d o p e p t i d a s e s 24.15 and 24.16 Activity (% inhibition) NT (20/~M)

Inhibitor and concentration (M) Phosphodiepryl 03 10 -9 10 -8 10 -7 10 - 6

Pro-Ile 10 -4 10 -3 10 -2 Cpp-Ala-Ala-Tyr-pAB 10 -8 10 -7 10 -6

10 -5

24.16

QFS (50 ~M)

24.15

24.16

24.15

1 50 93

0 10 55

21 79 97

1 62 95

100

93

100

100

9 51 91

0 0 0

27 79 97

0 0 0

0 1 8 52

5 34 85 98

0 0 25 77

8 48 90 100

[21]

E N D O P E P T I D A S E 24.16 IN CNS

375

dase 24.16 in vitro remains the dipeptide Pro-Ile. Whatever the substrate employed, a 10 -2 M concentration of this dipeptide almost fully abolished endopeptidase 24.16 activity without affecting endopeptidase 24.15 (Tables V and VI). When using QFS, it is also advantageous to include 0.5/xM CppAla-Ala-Tyr-pAB in the incubation in order to prevent the degradation of QFS by endopeptidase 24.15 (see Table VI).

Distribution o f Endopeptidase 24.16 in Rat Central Nervous System The above-described experimental procedure allowed us to establish the precise cartography of endopeptidase 24.16 in 60 brain areas (Figs. 6 and 7). The peptidase appears heterogeneously distributed with a 16-fold difference in the specific activities observed between the poorest and the richest cerebral areas (22). Endopeptidase 24.16 appeared in high concentrations in the olfactory bulb and tubercule, cingulate cortex, medial striatum, and globus pallidus and displays a particularly low activity in the CAI, C A 2 , and CA3 parts of the hippocampus (Fig. 6). The activity in the thalamus appears more homogeneously distributed with a maximum in the gelatinosus nucleus and a minimum in the ventral posterior nucleus (Fig. 7). Altogether, it is noticeable that the distribution of the peptidase in the brain is in agreement with that reported for neurotensin receptors (23). This topographical colocalization of the two molecular entities further argues in favor of a key role of endopeptidase 24.16 in the interruption of the receptor-mediated neurotensinergic signal.

In Vivo Studies In in vivo experiments, the route of administration must influence the selection of the inhibitor. When bolus administration is absolutely required, the low affinity and poor solubility of Pro-Ile precludes its use. By contrast, the high potency and hydrophilicity of phosphodiepryl 03 make it an efficient pharmacological tool. The use of Cpp-Ala-Ala-Tyr-pAB should permit verification that the protective effect of phosphodiepryl 03 is indeed mediated via an inhibition of endopeptidase 24.16 rather than through the blockade of endopeptidase 24.15. Preliminary experiments in our laboratory indicate that bolus intracerebroventricular injection of phosphodiepry103 potentiates neurotensin-induced hypothermia in rat. When the inhibitor can be infused in the tissue, a lower concentration of Pro-Ile is required, which renders the dipeptide amenable to pharmacological

OLFACTORY BULB

E a d pleaI(m layer

HYPOTWUMUS Supachiamatic nudeus

I n l d granular layer CEREBRAL CORTEX

1

Supnopk nudeus

HABWUUR NUCLEUS

Pariventriarlarnudes AMYGDALOID COMPLEX Cenlral n u d m

Arcuate nudeus

W nudeus

Lateral hypolhalarnicarea Mamrnilary n&us

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HIPPOCAMPAL FORMATION CAI

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Dentate gym

Ponlhe reticular nucleus

WHITE MAllER TRACTS Corps calmurn

CEREBELLUM

P

hternd capsule Ventral hippo+@ canmusure

S iflcactivity (ndrG P r e L mw mp)

FIG.6 Regional distribution of endopeptidase 24.16 in the rat central nervous system. Endopeptidase 24.16 activity was examined by means of QFS, fluorimetrically recorded, and quantified as described. Specific activities represent the mean *SEM of 4 to 10 independent determinations carried out with 2 to 5 independent homogenates prepared from pooled or individual brain micropunches.

377

[21] ENDOPEPTIDASE 24.16 IN CNS THALAMUS

Paratenial nudeus Paraventricular nucleus MediodusaJ nudeus

Laterodorsal nudeus Ventrolateral nucleus Ventromedial nudeus

m

Ventral posteriornudeus Gelatinosus nudeus Rhomboid nudeus Reuniens nucleus Posterior thalamic nudear group Dorsolateral 9eniculate nudeus

Medial geniculate nudeus

0

'

I

10

'

I

20

Specific activity (nmols Mcc-Pro-Leu / h / rag)

FIG. 7 Distribution of endopeptidase 24.16 in thalamic nuclei. Endopeptidase 24.16 activity was fluorimetrically recorded by means of QFS and quantified as described. Specific activities represent the mean -SEM of six to eight independent determinations carried out with two to five independent homogenates prepared from pooled or individual brain micropunches.

studies. We have shown that the infusion of a 5 mM concentration of ProIle in the arteriovenous mesenteric arcade irrigating an intestinal segment in the anesthesized dog drastically potentiated the recovery of exogenously infused neurotensin (24, 25). This effect is mimicked by the infusion of a 2/zM concentration of phosphodiepryl 03, while the same concentration of CPP-AAY-pAB totally fails to modify the recovery of neurotensin (24, 25). Such a combined approach with several inhibitors unambiguously identifies endopeptidase 24.16 as the target of Pro-Ile and phosphodiepryl 03 in vivo in dog intestine and for the first time directly demonstrates the participation of endopeptidase 24.16 in the physiological inactivation of peripheral neurotensin.

378

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E X T R A C E L L U L A R PROCESSING E N Z Y M E S IN THE CNS

A n t i b o d i e s D i r e c t e d t o w a r d R a t B r a i n E n d o p e p t i d a s e 24.16

Preparative Purification of Endopeptidase 24.16 and Immunization Procedure Endopeptidase 24.16 is purified from 100 rat brains according to the procedure previously described (4). After the last purification step, the enzyme is submitted to preparative sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and the bands corresponding to the activity are sliced and electroeluted (4~ 10 hr, 200 V). An aliquot of the electroeluate is then submitted to SDS-PAGE analysis and stained with Coomassie blue in order to estimate the rate of protein recovery. Purified endopeptidase 24.16 (30 /zg) in Freund's complete adjuvant is injected directly into the lymph nodes in the legs of a rabbit. Three weeks later, the rabbit is boosted intramuscularly and subcutaneously with 60/zg of peptidase in Freund' s incomplete adjuvant and two additional boosters (50 ~g of peptidase) are administered at 3-week intervals. The final injection (20/zg) occurs after a further month. The rabbit is first bled 1 month after the first injection and blood is then collected every week. The beginning of the immunological response is detected by dot-blot analysis about 7 weeks after the initial injection.

Purification of IgG The presence of several plasma proteases in the preimmune serum precludes the possibility of examining directly the putative anticatalytic properties of the immune serum. Therefore, the purified IgG fractions from both immune and preimmune sera are obtained after treatment with caprylic acid, according to the procedure previously described (26).

Anticatalytic Properties Purified endopeptidase 24.16 (0.1 /zg) is preincubated for 18 hr at room temperature with various dilutions of immune or preimmune IgG fractions in a final volume of 100/zl of 100 mM Tris-HC1, pH 7.5, containing 0.02% (w/v) bovine serum albumin. Neurotensin (2 nmol) is added and incubations are carried out for 1 hr at 37~ and terminated by acidification (10/zl of 1.25 M HC1). Neurotensin hydrolysis is monitored by HPLC, using the buffer/solvent system 2 described above. Our experiments indicate that the IgG-purified fraction of our immune

[21] ENDOPEPTIDASE 24.16 IN CNS

379

serum displays anticatalytic properties against endopeptidase 24.16 isolated from rat brain, whereas the preimmune fraction does not (27).

I m m u n o p re c ip ita tio n Purified peptidase (0.1 /xg) is preincubated under the conditions described above with 20/zl of preimmune (8 mg/ml) or immune (10 mg/ml) IgG fractions. Protein A-Sepharose (100/xl, 100 mg/ml) is added and incubated for 2 hr at room temperature under vigorous stirring. Incubations are then centrifuged (10 min, 10,000 g, 4~ and the supernatants are tested for neurotensin degradation and analyzed by HPLC as described above. These experiments clearly establish that the interaction of the IgG with the purified peptidase leads to full clearance of the activity from the supernatant, whereas the preimmune fraction is unable to immunoprecipitate the enzyme (27).

S D S - P A G E and Western Blot Analysis Aliquots of tissue homogenates or purified enzymes are suspended in 100/zl of sodium phosphate buffer, pH 7.5, containing 2% (w/v) SDS and 5% (v/v) 2-mercaptoethanol. Samples are boiled and electrophoresed overnight (15 mA/gel) according to a classic procedure (28) in 8% (w/v) acrylamide gels. Proteins are then blotted onto nitrocellulose sheets according to the procedure described by Towbin et al. (29). Efficiency of blotting is checked by revealing the transferred proteins with 0.2% (v/v) Ponceau Red in 3% (v/v) trichloroacetic acid. Nitrocellulose is then destained with 140 mM NaC1-20 mM Tris-HC1, pH 7.5 (buffer A), containing 0.2% (w/v) Tween 20, and incubated for 4 hr at room temperature in buffer A containing 3% (w/v) bovine serum albumin. Nitrocellulose is then exposed overnight to a 1 : 1000 dilution of the preimmune or immune IgG fractions [in buffer A containing 1% (w/v) bovine serum albumin]. Sheets are rinsed (5 x 5 min) with buffer A containing 0.2% Tween 20, then exposed for 3 hr to a 1:800 dilution of goat anti-rabbit immunoglobulins coupled to peroxidase according to the manufacturer recommendations (Pharmacia, Piscataway, NJ). Nitrocellulose is finally rinsed as described above and the IgG-antigen complexes are revealed with 100 ml of methanol-buffer A (1:9, v/v) containing 30 mg of 4-chloro-l-naphthol and by the addition of 100/zl of H202. The above-described procedure demonstrates that the IgG-purified fraction of the antiserum recognizes the denatured form of endopeptidase 24.16. The IgG appears to be highly specific for the enzyme because a single protein

380

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

kD 200P'-

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FIG. 8 Western blot analysis of proteins from various rat organs. Homogenates of various rat organs were prepared, electrophoresed, and blotted onto nitrocellulose sheets as detailed. Transferred proteins were incubated with a 1 : 1000 dilution of the purified immune IgG fraction directed toward endopeptidase 24.16 and the immunopositive reaction was revealed with goat anti-rabbit IgGs coupled to peroxidase. No reaction was observed after incubation with the preimmune IgG. Lanes: a, lung; b, spleen; c, stomach; d, liver; e, heart; f, testis; g, colon; h, caecum; i, duodenum; j, skeletal muscle; k, kidney; 1, ileum; m, brain. band, the molecular weight of which closely corresponds to that of the purified peptidase, is immunolabeled in a homogenate of whole rat brain (27). Western blot analysis shows that these IgGs are totally species specific because the 75-kDa protein is recognized only in murine brain (27) and reveals the ubiquitous organ distribution of endopeptidase 24.16 in the rat (Fig. 8). Finally, the IgGs provide clearcut evidence that endopeptidase 24.15 is not immunologically related to endopeptidase 24.16 because IgG totally fails to interact with endopeptidase 24.15 (Fig. 9). Finally, the IgG fraction is used to establish that endopeptidase 24.16 is colocalized with specific neurotensin receptors in a discrete population (less than 10% of the cells) of mouse embryo neurons in primary cultures (30). Furthermore, this tool allowed study of the distribution and subcellular localization of the enzyme at the light and electron microscopic levels, which revealed the dual astrocytic and neuronal presence of endopeptidase 24.16 in rat substantia nigra and ventral tegmental areas, corresponding to brain zone enriched in neurotensin receptors (31).

Conclusion We have described here the development and use of a fluorimetric substrate, specific inhibitors, and a polyclonal antiserum directed toward endopeptidase 24.16. These new tools have permitted further investigation of the biochemi-

381

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Fic. 9 Western blot analysis of endopeptidases 24.15- and 24.16-enriched pools. Endopeptidase 24.16 obtained after the hydroxylapatite chromatography of the D~ pool [as described in (4)] (lane 1, a and b) and endopeptidase 24.15 pool H2 (21) (lane 2, a and b) were analyzed by SDS-PAGE (lanes la and 2a). Proteins were transferred and hybridized with the IgG-purified fraction of a polyclonal antiserum directed toward endopeptidase 24.16 (lanes lb and 2b) (27). The leftmost lane corresponds to the SDS-PAGE of synthetic bovine serum albumin (BSA). Antigen-IgG complexes were revealed as described (see text). cal properties of the enzyme. It is now possible to monitor the activity in a complex mixture of enzymes such as crude extracts, provided that some precautions are taken. Furthermore, the development of specific inhibitors has established, for the first time, that endopeptidase 24.16 indeed participates in the physiological metabolism of neurotensin. These inhibitors will be of further help in assessing whether the enzyme also contributes to the catabolic pathways of other neuropeptides. Finally, the development of a monospecific antiserum directed toward endopeptidase 24.16 has rendered possible the immunological screening of a rat brain Xgt 11 cDNA library and will allow the molecular cloning of the enzyme, which is in progress in our laboratory.

Acknowledgments We gratefully acknowledge V. Ch6ron for expert secretarial assistance and F. Aguila for excellent artwork. This work was supported by the Centre National de la Recherche Scientifique and the Institut National de la Sant6 et de la Recherche M6dicale.

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III EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

References F. Checler, J. P. Vincent, and P. Kitabgi, J. Neurochem. 41, 375 (1983). 2. F. Checler, P. C. Emson, J. P. Vincent, and P. Kitabgi, J. Neurochem. 43, 1295 (1984). F. Checler, J. P. Vincent, and P. Kitabgi, J. Neurochem. 45, 1509 (1985). 4. F. Checler, J. P. Vincent, and P. Kitabgi, J. Biol. Chem. 261, 11274 (1986). 5. H. Barelli, J. P. Vincent, and F. Checler, Eur. J. Biochem. 175, 481 (1988). 6. H. Barelli, J. P. Vincent, and F. Checler, Eur. J. Biochem. 211, 79 (1993). 7. E. C. Webb, Eur. J. Biochem., Suppl. 2, 519 (1989). 8. F. Checler, J. P. Vincent, and P. Kitabgi, J. Pharmacol. Exp. Ther. 227, 743 (1983). P. Kitabgi, F. Checler, J. Mazella, and J. P. Vincent, Rev. Basic Clin. Pharmacol. 5, 397 (1985). 10. F. B. Jolicoeur, A. Barbeau, F. Rioux, R. Quirion, and S. Saint-Pierre, Peptides (N.Y.) 2, 171 (1981). ll. P. T. Loosen, C. B. Nemeroff, G. Bissette, G. Burnett, A. J. Prange, Jr., and M. A. Lipton, Neuropharmacology 17, 109 (1978). 12. J. E. Rivier, L. H. Lazarus, M. H. Perrin, and M. R. Brown, J. Med. Chem. 20, 1409 (1977). 13. S. Aswaniumar and A. N. Radhakrishnan, Biochim. Biophys. Acta 304, 211 (1972). 14. P. Dauch, H. Barelli, J. P. Vincent, and F. Checler, Biochem. J. 280, 421 (1991). 15. H. Barelli, F. Girard, S. St. Pierre, P. Kitabgi, J. P. Vincent, and F. Checler, Neurochem. Int. 12, 351 (1988). 16. P. Dauch, J. P. Vincent, and F. Checler, Eur. J. Biochem. 202, 269 (1991). 17. P. A. Barlett and C. K. Marlowe, Biochemistry 26, 8553 (1987). 18. J. E. Hanson, A. P. Kaplan, and P. A. Barlett, Biochemistry 28, 6294 (1989). 19. V. Dive, A. Yiotakis, A. Nicolaou, and F. Toma, Eur. J. Biochem. 191,685 (1990). 20. H. Barelli, V. Dive, A. Yiotakis, J. P. Vincent, and F. Checler, Biochem. J. 287, 621 (1992). 21. H. Barelli, J. P. Vincent, and F. Checler, Neurochem. Life Sci. Adv. 10, l l5 (1991). 22. P. Dauch, Y. Masuo, J. P. Vincent, and F. Checler, J. Neurochem. 59, 1862 (1992). 23. E. Moyse, W. Rost~ne, M. Vial, K. Leonard, J. Mazella, P. Kitabgi, J. P. Vincent, and A. Beaudet, Neuroscience 22, 527 (1987). 24. F. Checler, H. Barelli, P. Dauch, B. Vincent, V. Dive, A. Beaudet, E. E. Daniel, J. E. T. Fox-Threlkeld, Y. Masuo, and J. P. Vincent, Biochem. Soc. Trans. 21, 692 (1993). 25. H. Barelli, J. E. T. Fox-Threlkeld, V. Dive, E. E. Daniel, J. P. Vincent, and F. Checler, Br. J. Pharmacol. 112, 127 (1994). 26. M. Steinbuch and R. Audran, Arch. Biochem. Biophys. 134, 279 (1969). 27. F. Checler, H. Barelli, and J. P. Vincent, Biochem. J. 257, 549 (1989). 28. U. K. Laemmli, Naure (London) 227, 680 (1970). 29. H. Towbin, T. Staehelin, and J. Gordon, Proc. Natl. Acad. Sci. U.S.A. 76, 4350 (1979). 30. J. Chabry, F. Checler, J. P. Vincent, and J. Mazella, J. Neurosci. 10, 3916 (1990). 31. J. Woulfe, F. Checler, and A. Beaudet, Eur. J. Neurosci. 4, 1309 (1992). o

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[221

Autoradiographic Techniques to Map Angiotensin-Converting Enzyme in Brain and Other Tissues Siew Yeen Chai and Frederick A. O. Mendelsohn

Background Angiotensin-converting enzyme (ACE), a peptidyl carboxypeptidase (EC 3.4.15.1, peptidyl dipeptidase A), is a membrane-bound ectoenzyme of 146 kDa that consists of a large extracellular domain with a transmembrane anchor and a small intracellular carboxyl terminus. It is responsible for the conversion of angiotensin I to the potent vasoconstrictor angiotensin II and the degradation of bradykinin. Although ACE is also capable of hydrolyzing a range of other peptides in vitro, including tachykinins, opioid peptides, neurotensin, bombesin, and luteinizing hormone-releasing hormone (1), it is not known if any of these other peptides are endogenous substrates of the enzyme in vivo. The cDNA sequence of human endothelial ACE consists of 4024 nucleotides that encode 1306 amino acids (2). The extracellular domain contains two regions of high sequence homology, both of which contain active site sequences. Although both sites are catalytically active, the C-terminal site appears to be responsible for most of the hydrolysis of angiotensin I (3) and binding of the ACE inhibitors (4). The N-terminal site appears to have some structural constraints that hinder the binding of some substrates and larger drug molecules (5). Testicular ACE differs from the somatic form of the enzyme in that it has only one active site and has a sequence identical to the C terminus of the endothelial enzyme (6). Angiotensin-converting enzyme is widely distributed and is found on the lumenal surfaces of all vascular beds. The enzyme was first described in the brain by Yang and Neff (7), who measured the enzyme activity in brain homogenates by a biochemical assay. The highest concentration of ACE was detected in the striatum, followed by the cerebellar cortex. Later biochemical studies that involved microdissection of brain structures demonstrated that ACE was heterogeneously distributed in discrete nuclei. However, earlier immunohistochemical mapping of brain ACE using different antibodies produced conflicting results. Most were unable to demonstrate the presence of the enzyme in any brain structures except for the choroid plexus, circumMethods in Neurosciences, Volume 23

Copyright 9 1995 by Academic Press, Inc. All rights of reproduction in any form reserved.

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

ventricular organs, and cerebral vasculature (8-10), except for one that detected ACE within neuronal elements in the hypothalamic paraventricular and supraoptic nuclei and in circumventricular organs (11). These discrepancies led us (12-15) and others (16) to develop an in vitro autoradiographic method to map the distribution of ACE in the brain using specific, highaffinity inhibitors of the enzyme. The advantages of this technique are, first, its sensitivity and, second, that we are able to determine the specificity of the radioligand and characterize its binding properties and are therefore confident that the radioligand is binding to the active site of ACE on the tissue sections.

Angiotensin-Converting Enzyme Inhibitors Availability of Specific High-Affinity Inhibitors The availability of specific high-affinity inhibitors provides excellent ligands with which to label ACE. We used a derivative of lisinopril, a substituted N-carboxymethyl dipeptide that is a potent, highly specific inhibitor of ACE. This compound, N-[(S)-l-carboxy-3-phenylpropyl]-L-lysyltyrosyl-e-proline, known as 351A, has a tyrosine ring on the lysine side chain to enable the incorporation of 1251 into the molecule without interfering with its binding properties to the catalytic site of ACE. Other ACE inhibitors that have been used as markers for the enzyme include captopril (16) and Ro 31-8472, a hydroxy derivative of cilazapril (5).

Radiolabeling of Inhibitors The compound 351A is iodinated by the chloramine-T method (17) and the reaction products separated on a Sephadex SP-C25 column (14). The specific activity of this compound, as determined by radioimmunoassay, is between 1600 and 1700/xCi//xg. In addition to 125I, tritium has also been used as the radioactive marker (16). The disadvantage of using tritium is the lower specific activity achieved during incorporation of the radioisotope.

Characterization of lnhibitors Using Membrane Fractions The binding properties of this radioligand to different tissues can be characterized either on membrane fractions or on tissue sections. The advantage of using membrane fractions is that a wide range of competing ligands can be

385

[22] AUTORADIOGRAPHY OF ACE

TABLE I Inhibitory Constants for Teprotide, Captopril, Enalaprilat, and Lisinopril a Ki values (nM) Competing ligand

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Striatum

Teprotide Captopril Enalaprilat Lisinopril

434 32.3 7.1 2.4

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used quickly to determine the specificity of the radioligand. Binding studies of the radioligand determined on tissue sections allow binding properties to be defined in discrete anatomical areas, but are much more time consuming and have greater errors than membrane binding systems. Initially, the binding properties of 125I-labeled 351A were characterized on rat lung (12) and brain striatal (14) membranes. A 1000- to 20,000-g membranerich fraction, prepared from rat lung or striatum, was incubated with 20 nCi of ~25I-labeled 351A in phosphate-buffered saline (PBS) containing 0.2% (w/v) bovine serum albumin (BSA). Nonspecific binding was determined in the presence of 1 /zM unlabeled lisinopril. Free and bound ligand were separated by filtration through glass fiber filters, the radioactivity was measured in a 3' counter, and binding isotherms were analyzed by a nonlinear, iterative, model-fitting program (18). 125I-Labeled 351A bound to lung membranes with a tl/2 o f approximately 2 min at 25~ reached a plateau by 30 min, and remained stable for more than 3 hr. The dissociation of the enzyme inhibitor complex in the presence of 1/zM lisinopril was slow, with a tl/2 of 65 min at 25~ (12). Specific binding of the radioligand to lung or striatal membranes was abolished by the addition of 1 mM ethylenediaminetetraacetate (EDTA), which chelates the essential divalent cation, Z n 2+, f r o m the catalytic site of the enzyme and thereby blocks its enzymatic activity (12, 14). A range of chemically unrelated ACE inhibitors, lisinopril, enalaprilat, captopril, and teprotide competed for the binding of 125I-labeled 351A to lung or striatal membranes (Table I) with affinities that paralleled their anticatalytic potencies. These competition studies suggest that 125I-labeled 351A specifically binds to the active site of ACE (12, 14).

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EXTRACELLULAR PROCESSING ENZYMES IN THE CNS

Endopeptidase 24.11 Angiotensin-converting enzyme displays many characteristics similar to endopeptidase (EC 3.4.24.11, neprilysin): its dependence on zinc for its catalytic activity, its broad spectrum of peptide substrates, and its tissue distribution. Therefore, specific inhibitors of endopeptidase 24.11, phosphoramidon and the hydroxamic acid derivative of Ala-Gly amide, which have much lower affinities for ACE, were included in the competition studies. These inhibitors displayed much lower affinities in competing for the binding of ~25I-labeled 351A with 50% inhibitory concentration (IC50) values of 4 and 50/zM, respectively (14). Moreover, in experiments in which the specific endopeptidase 24.11 inhibitor [3H]-N-[(2RS)-3-hydroxyaminocarbonyl-2-benzyl-l-oxopropyl]glycine([3H]HACBO-Gly) was used, captopril was ineffective in competing for the binding of this radioligand (19).

In Vitro A u t o r a d i o g r a p h y Preparation of Tissue Tissues used in in vitro autoradiography are frozen without fixation. However, for the binding of 125I-labeled 351A to rat tissues, it was found that perfusion with 1% paraformaldehyde at 4~ did not alter the binding patterns or properties except for a slight reduction in total radioligand binding. Preservation of tissue morphology was much improved. For larger tissues, particularly those obtained from human, the advantage of a light fixation is diminished owing to morphological damage incurred during postmortem delay. Rats are anesthetized with sodium methohexitane and perfused intracardially with isotonic saline followed by 1% paraformaldehyde. The tissues are then rapidly removed, snap-frozen in isopentane maintained at -40~ and stored at -70~ until use. For human tissues, the postmortem specimens are blocked, frozen at -40~ in dry ice, and stored. Ten- to 20-/zm sections are cut in a cryostat maintained at -15 to -20~ thaw-mounted onto gelatin-coated slides, and dried in a desiccator for 2 hr at 4~

Basic Technique In vitro autoradiographic localization of ACE involves an initial 15-min preincubation in a phosphate-buffered saline solution containing 0.2% bovine

[22] AUTORADIOGRAPHY OF ACE

387

serum albumin to remove endogenous substrates from the active site of the enzyme. This is then followed by a 60-min incubation in the phosphatebuffered saline containing 0.2% bovine serum albumin and 0.3/zCi/ml of the radioligand ~25I-labeled 351A at 20~ After the incubation, the sections are passed through four successive 1-min washes in ice-cold phosphate-buffered saline and dried under a stream of cold air. The sections are then loaded into X-ray cassettes and exposed to AgfaScopix CR-3B (Agfa-Gevaert, Matsel, Belgium) X-ray film for 3 to 7 days at room temperature. In each cassette, a set of radioactivity or enzyme standards is included. The preparation of these standards and their use in the quantitation of the autoradiographs are described below. The X-ray films are processed in a Kodak (Rochester, NY) RP X-Omat automatic developer and the optical densities on the films quantitated by either an Eye-Corn model 850 image analysis system (Spatial Data Systems, Springfield, VA) or an MCID image analysis system (Imaging Research, Inc., Ontario, Canada).

Nonspecific Binding Nonspecific binding to tissue sections was determined in parallel incubations containing 1 mM EDTA or 1/zM lisinopril. When using ~25I-labeled 351A at a concentration of 250 pM, the nonspecific binding was less than 1% of total binding, which did not produce a detectable image on the X-ray film.

Quantitation of Enzyme Activity Using Radioactivity Standards Simultaneous use of radioactivity standards during the exposure of tissue sections permits accurate conversion of optical density to units of radioactivity. The radioactivity standards are prepared from 20-/zm sections of a frozen tissue core 5 mm in diameter, prepared from the tissue of interest. Six to eight 20-/zm tissue disks are thaw-mounted onto a gelatinized slide and air dried. A range of known 125I radioactivity is then applied to the tissue disks in a volume of 5/xl and dried. Mean protein content of the tissue is then measured from 20-/xm sections that have been digested in 1.0 M NaOH. The area of these sections can be measured from images obtained on the X-ray film, using an image analyzer and the value of mean protein content per unit area calculated. The radioactivity standards are corrected for decay.

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The radioactivity standards enable calibration of optical density on the autoradiographs in terms of ~25I disintegrations per minute (dpm)/mm 2, or femtomoles of radioligand bound per mean milligram of protein on the tissue section.

Quantitation of Enzyme Activity Using Enzyme Standards In addition to calibrating the optical densities in terms of amount of radioactivity present in the tissue, the system may be calibrated in terms of ACE activity using enzyme standards. These are prepared by serial dilutions of tissue membranes suspended in phosphate-buffered saline containing 1% gelatin. Five 1-/A aliquots are then applied to gelatin-coated slides, dried at 4~ in a desiccator, and then carried through the same incubation process as for the tissue sections. These standards are exposed to X-ray film simultaneously with the tissue sections. Samples of the membrane suspension are assayed for ACE by a fluorimetric method (20) and for protein content by a modified Lowry procedure (21). The correlation between ACE activity and radioactivity is linear.

Sensitivity of Radioligand In addition to being highly specific, the technique of in vitro autoradiographic localization of ACE in tissue sections is remarkably sensitive. A conservative estimation of the sensitivity of this technique to quantitate the binding of ~25I-labeled 351A to ACE is based on the lowest standard that is easily detectable after a 7-day exposure of 10 dpm/mm 2. When the specific activity of 1600 Ci/mmol is used, this in vitro autoradiographic technique is able to detect approximately 15 molecules//~m 2.

Anatomical Localization of Angiotensin-Converting Enzyme After the sections have been subjected to the autoradiographic procedure, they were then either stained with thionin for brain or with haematoxylin and eosin for other peripheral tissues in order to aid anatomical localization of the radioligand-binding sites. The directly adjacent sections were sometimes stained for histology instead of the actual sections because of better preservation of tissue morphology. Figure 1A shows an autoradiographic image of FIG. 1 Autoradiographic image of ACE in the human substantia nigra (A) and the adjacent section, which has been stained with thionin (B). The arrows indicate the compact cells of the substantia nigra. R, red nucleus; cp, cerebral peduncle; SNR, substantia nigra pars reticulata.

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125I-labeled 351A binding to human substantia nigra and Fig. 1B shows the adjacent section, which had been stained with thionin. As can be seen, nSI-labeled 351A binding is confined to the reticular part of the substantia nigra (15). In some instances, the adjacent sections were stained for another enzyme, for example, acetylcholinesterase. Its distribution in the brain is well mapped and extensively studied. Figure 2A shows an autoradiographic image of ~25Ilabeled 351A binding to human caudate nucleus and Fig. 2B shows the adjacent section, which had been stained for acetylcholinesterase. The patches of higher ~25I-labeled 351A binding corresponded to the regions of lower acetylcholinesterase activity, confirming that ACE is more concentrated in the acetylcholinesterase-poor striosomes (15).

Localization of Angiotensin-Converting Enzyme in Brain The distribution of ACE in the rat brain, as detected by in vitro autoradiography using nSI-labeled 351A, could be broadly divided into five categories (14). First, it was found on the endothelial surface of moderate-sized cerebral vessels, as in the vasculature of all organs. Second, a high density of ACE was detected in the choroid plexus (Fig. 3), where it was shown by immunohistochemical studies to occur on the brush border of the epithelial cells. Third, high concentrations of ACE were detected in all forebrain circumventricular organs, where the enzyme could convert circulating angiotensin I to angiotensin II to act on the local high densities of angiotensin II receptors present in these structures. Fourth, ACE was found in sites that correspond to the distribution of angiotensin II immunoreactivity and angiotensin II receptors, such as the hypothalamic neurosecretory nuclei and the dorsal vagal complex. At these sites, ACE may participate in the local formation of angiotensin II. Fifth, ACE was also detected in brain sites that were not thought to be rich in angiotensin II or its receptors, for example, the basal ganglia, hippocampal formation, cerebellar cortex (Fig. 3), and inferior olivary nucleus. Angiotensin-converting enzyme at these sites could be involved in processing neuropeptides other than angiotensin. Although we were unable to demonstrate angiotensin II receptors in the basal ganglia of the rat, in the

FIG. 2 Autoradiographic image of ACE distribution in the human caudate nucleus (A) and the adjacent section, which has been stained for acetylcholinesterase (B). The arrows indicate the striosomal patches, which contain high concentrations of ACE and low levels of acetylcholinesterase. CN, Caudate nucleus; ic, internal capsule.

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human brain these structures contained moderate to high densities of the receptors (22). A similar overall pattern of distribution of ACE was observed in other mammalian species (Fig. 4). The distribution of ACE in the monkey Macaca fascicularis (23) and human striatum (15) was heterogeneous. Angiotensinconverting enzyme was enriched in the striosomes, which are defined by patches of low acetylcholinesterase activity (Fig. 2). In contrast to the rat, in rabbit, monkey (23), and human brains (15) a moderate to high density of ACE was found throughout the cerebral cortex (Fig. 4). Mapping of brain ACE with [3H]captopril (16) gave a pattern that was broadly consistent with results obtained with 125I-labeled 351A. High concentrations of [3H]captopril binding were found in the choroid plexus, basal ganglia, and hypothalamic neurosecretory nuclei. However, 125I-labeled 351A detected the presence of ACE in many other sites in the amygdaloid complex, hippocampus, thalamus, cerebellum, and brainstem (14). This is because the

[22] AUTORADIOGRAPHY OF ACE

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iodinated radioligand provided higher resolution autoradiographs that enabled more detailed anatomical localization of brain ACE. Moreover, it required shorter exposure times for its autoradiographs because of its higher specific activity. Applications

Accurate Quantitation o f Small Brain Nuclei The in vitro autoradiographic mapping of ACE in rat (12-14) and human (15, 24) brains revealed high concentrations of the enyzme in basal ganglia structures, including the caudate, putamen, internal and external globus pallidus, entopeduncular nucleus, and substantia nigra pars reticulata. These structures appear to be connected by a continuous pathway. Indeed, in the human basal ganglia, ACE is present in fibers in the internal capsule and cerebral peduncles (Fig. 5, color plate). Selective excitotoxin lesion of the rat striatum or 6-hydroxydopamine lesion of the substantia nigra pars compacta was carried out to investigate if ACE was associated with descending striatonigral or ascending nigrostriatal projections. In rats that had N-methyl-o-aspartic acid injected into the right striatum, ACE was decreased in the caudate putamen, globus pallidus, entopeduncular nucleus, and substantia nigra pars reticulata (Fig. 6, color plate). By contrast, 6-hydroxydopamine lesion of the substantia nigra, to lesion the ascending nigrostriatal dopaminergic system selectively, did not affect ACE levels in these structures (25). The high levels of ACE in other nuclei not associated with the basal ganglia were also not affected by either of the neurotoxins. These lesion studies confirmed that ACE is associated with neurons within the striatopallidal, striatonigral, and pallidonigral systems. Similarly, ACE was decreased in these basal ganglia structures in Huntington's disease but not in Parkinson's disease, confirming this assignment in the human brain (26). In vitro autoradiography enabled the detection of ACE in very small brain nuclei and even within neuronal fibers. Moreover, accurate quantitation of enzyme levels after chemical or other physical intervention can be carried out within a small brain nucleus because the autoradiograph can be overlaid on top of a stained section to determine the boundary of the nuclei.

Drug Penetration Studies Angiotensin-converting enzyme inhibitors have been successfully used in the treatment of hypertension and heart failure. However, the sites of action of these drugs are not clearly understood. Although ACE inhibitors

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[22] AUTORADIOGRAPHY OF ACE

395

were initially thought to mediate their effect via blockade of formation of circulating angiotensin II, their actions are more extensive than previously suspected. The hypotensive effect of these drugs outlasts inhibition of plasma ACE and they are effective in non-renin-dependent hypertension. Many investigators now believe that ACE inhibitors mediate their longterm hypotensive effects via inhibition of tissue ACE. The technique of in vitro autoradiography has also been used successfully to assess the sites and degree of inhibition of ACE inhibitors administered in vivo. This is possible because many of these drugs exhibit high affinity and tight binding to the active site of ACE, with relatively slow dissociation rates (27-29). The time course of tissue ACE inhibition was assessed by gavage-feeding rats with a particular dose of the drug followed by tissue collection at various time intervals (1-48 hr). The tissues were then sectioned and subjected to in vitro autoradiography as described above, except that the preincubation step was omitted. Trunk blood was collected for measurement of plasma ACE and drug levels. Similarly, dose-response studies were also carried out in which rats were administered different doses of a drug and tissue ACE inhibition measured by in vitro autoradiography. Acute administration of a single dose of an ACE inhibitor (lisinopril, perindopril, or benazepril) all produced varying degrees of tissue ACE inhibition (27-30). The difference in tissue ACE inhibition is probably due to tissue bioavailability and lipophilicity of the drug. Angiotensinconverting enzyme, which is located in brain structures other than the circumventricular organs, was not inhibited by an acute oral dose of most ACE inhibitors studied (Fig. 7, color plate) and the testicular enzyme was not blocked by any of the inhibitors administered. Angiotensinconverting enzyme at these sites is protected by the blood-testis or blood-brain barrier because it has been shown in membrane-binding studies that the enzyme in both these tissues is equally susceptible in vitro to all ACE inhibitors tested. The duration of tissue ACE inhibition was more prolonged than the suppression of plasma ACE and appeared to correlate more closely with inhibition of the pressor effect to exogenous angiotensin I (27). This

FIG. 4 Autoradiographic image of ACE distribution in a coronal section through the macaque monkey (A) and human (B) diencephalon, showing dense levels of ACE in the caudate nucleus (CN), putamen (Pu), nucleus acumbens (Acb), and external globus pallidus (GPe). Moderate densities of ACE are also present throughout the cerebral cortices, cc, corpus callosum; MS, medial septum; DB, diagonal band of Broca; ic, internal capsule; CI, claustrum.

396

III EXTRACELLULARPROCESSING ENZYMES IN THE CNS observation supports the hypothesis that the more long-term antihypertensive effect of an ACE inhibitor is due to tissue ACE inhibition and not just to the suppression of circulating angiotensin II levels.

Chronic Angiotensin-Converting Enzyme Inhibition Studies The chronic administration of an ACE inhibitor has been shown to cause a marked increase in plasma ACE level. The technique of in vitro autoradiography has been adapted to investigate this phenomenon. Angiotensin-converting enzyme belongs to a group of zinc metallopeptidases that are dependent on Zn 2§ for their catalytic activity. This property of ACE has enabled the design of experiments to evaluate tissue ACE inhibition and induction simultaneously. Tissue ACE inhibition was assessed as described in the previous section. In addition, tissue ACE induction was measured as follows" the enzyme was reversibly inactivated with ethylenediaminetetraacetic acid (EDTA), which chelated the Zn 2§ from the active site, resulting in dissociation of the bound inhibitor from the enzyme. Angiotensin-converting enzyme was then reactivated by the removal of EDTA and the active site replenished by the addition of Zn 2§ in the incubation together with the radioligand to measure total ACE in the tissue (31). The chronic administration of the ACE inhibitor lisinopril significantly increased ACE levels in plasma and lung but not in other tissues studied. This finding agreed with an earlier study that demonstrated increased ACE activity in the lung after chronic captopril treatment but not in the testis, kidney, or small intestine (32). The precise mechanism of plasma and tissue ACE induction, and its differential effect in the various tissues, are not known. FIG. 5 Computer-generated pseudocolor image of ACE distribution in a coronal section through the human diencephalon, showing high concentrations of ACE in the caudate nucleus, putamen, and globus pallidus, and in fibers projecting to the substantia nigra (arrows). Moderate densities of ACE are also present in the hippocampus (Hi) and insular cortex (ICx). The color scale is as follows: red represents high densities of ACE, yellow and green moderate, and blue low to undetectable levels of ACE. CN, Caudate nucleus; Pu, putamen; GPe, external globus pallidus; GPi, internal globus pallidus. FIG. 7 Computer-generated pseudocolor images of ACE distribution in coronal sections through the diencephalon of a control rat (A) and a rat 4 hr after an oral dose of lisinopril (10 mg/kg), showing blockade of ACE in the organum vasculosum of the lamina terminalis (OVLT) but not in the caudate putamen (CPu) or the choroid plexus (ClaP); ac, anterior commisure.

[22] AUTORADIOGRAPHYOF ACE

397

Conclusion The in vitro autoradiographic localization of ACE in the rat brain revealed the presence of the enzyme in many previously unreported sites and extended the mapping in sites where the enzyme was known to occur. Moreover, this technique provided the first detail mapping of ACE in the human brain. The extension of this technique to assess in vitro the degree of tissue ACE inhibition after ex vivo administration of ACE inhibitors provided more accurate and new information on the differential effect of these drugs in the various tissues. In the brain, the degree of blockade of ACE in structures within the blood-brain barrier appears to depend on the lipophilicity of the drugs. In addition, using in vitro autoradiography, we were able to assess accurately the degree of tissue ACE induction.

Acknowledgment These studies were supported by grants from the National Health and Medical Research Council and the National Heart Foundation of Australia and the Austin Hospital Medical Research Foundation. Siew Yeen Chai is supported by a National Health and Medical Research Council Australian Postdoctoral Fellowship. We gratefully acknowledge the generosity of Dr. C. Sweet of the Merck Institute for Therapeutic Research for the gift of 351A.

References 1. R. A. Skidgel, R. Defendini, and E. G. Erdos, in "Neuropeptides and Their Peptidases" (A. J. Turner, ed.), p. 165. Ellis Horwood, Chichester, 1987. 2. F. Soubrier, F. Alhenc-Gelas, C. Hubert, J. Allegrini, M. John, G. Tregear, and P. Corvol, Proc. Natl. Acad. Sci. U.S.A. 85, 9386 (1988). 3. L. Wei, F. Soubrier, P. Corvol, and E. Clauser, J. Biol. Chem. 266, 9002 (1991). 4. R. B. Perich, B. Jackson, F. Rogerson, and F. A. O. Mendelsohn, Mol. Pharmacol. 4211, 280 (1992). 5. R. B. Perich, B. Jackson, M. R. Attwood, K. Prior, and C. I. Johnston, Pharm. Pharmacol. Lett. 1, 41 (1991). 6. R. S. Kumar, J. Kusari, S. N. Roy, R. L. Softer, and G. C. Sen, J. Biol. Chem. 264, 16754 (1989). 7. H. Y. T. Yang and N. H. Neff, J. Neurochem. 19, 2443 (1972). 8. H. J. Wigger and S. A. Stalcup, Lab. Invest. 38(5), 581 (1978). 9. E. Rix, D. Ganten, G. Stock, and R. Taugner, Exp. Brain Res. S4, 126 (1982). 10. M. S. Brownfield, I. A. Reid, D. Ganten, and W. F. Ganong, Neuroscience 7(7), 1759 (1982).

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III EXTRACELLULAR PROCESSING ENZYMES IN THE CNS 11. R. Defendini, E. A. Zimmerman, J. A. Weare, F. Alhenc-Gelas, and E. G. Erdos, Neuroendocrinology 37, 32 (1983). 12. F. A. O. Mendelsohn, Clin. Exp. Pharmacol. Physiol. 11, 431 (1984). 13. F. A. O. Mendelsohn, S. Y. Chai, and M. Dunbar, J. Hypertens. 2(s3), 21 (1984). 14. S. Y. Chai, F. A. O. Mendelsohn, and G. Paxinos, Neuroscience 20, 615 (1987). 15. S. Y. Chai, J. S. McKenzie, M. J. McKinley, and F. A. O. Mendelsohn, J. Comp. Neurol. 291, 179 (1990). 16. S. M. Strittmatter, M. M. S. Lo, J. A. Javitch, and S. H. Snyder, Proc. Natl. Acad. Sci. U.S.A. 81, 1599 (1984). 17. W. M. Hunter and F. C. Greenwood, Nature (London) 194, 495 (1962). 18. P. J. Munson and D. Rodbard, Anal. Biochem. 107, 220 (1980). 19. G. Waksman, E. Hamel, M. C. Fournie-Zalusky, and B. P. Roques, Proc. Natl. Acad. Sci. U.S.A. 83, 1523 (1986). 20. J. Friedland and E. Silverstein, Am. J. Clin. Pathol. 66, 416 (1976). 21. E. F. Hartree, Anal. Biochem. 48, 422 (1972). 22. A. M. Allen, G. Paxinos, M. J. McKinley, S. Y. Chai, and F. A. O. Mendelsohn, J. Comp. Neurol. 312, 291 (1991). 23. S.Y. Chai, M. J. McKinley, G. Paxinos, and F. A. O. Mendelsohn, Neuroscience 42, 483 (1991). 24. A. M. Allen, S. Y. Chai, J. Clevers, M. J. McKinley, G. Paxinos, and F. A. O. Mendelsohn, J. Comp. Neurol. 269, 249 (1988). 25. S. Y. Chai, M. J. Christie, P. M. Beart, and F. A. O. Mendelsohn, Neurochem. Int. 10, 101 (1987). 26. A. M. Allen, D. P. MacGregor, S. Y. Chai, G. A. Donnan, S. Kaczmarczyk, K. Richardson, R. Kalnins, J. Ireton, and F. A. O. Mendelsohn, Ann. Neurol. 32, 339 (1992). 27. K. Sakaguchi, S. Y. Chai, B. Jackson, C. I. Johnson, and F. A. O. Mendelsohn, Neuroendocrinology 48, 223 (1988). 28. K. Sakaguchi, S. Y. Chai, B. Jackson, C. I. Johnston, and F. A. O. Mendelsohn, Hypertension (Dallas) 11, 230 (1988). 29. K. Sakaguchi, B. Jackson, S. Y. Chai, F. A. O. Mendelsohn, and C. I. Johnston, J. Cardiovasc. Pharmacol. 12, 710 (1988). 30. S. Y. Chai, R. S. Perich, B. Jackson, F. A. O. Mendelsohn, and C. I. Johnston, Clin. Exp. Pharmacol. Physiol. 19(s19), 7 (1992). 31. M. Kohzuki, C. I. Johnston, S. Y. Chai, B. Jackson, R. Perich, D. Paxton, and F. A. O. Mendelsohn, J. Hyperten. 9, 579 (1991). 32. F. Fyhrquist, T. Forslund, I. Tikkanen, and C. Gronhagen-Riska, Eur. J. Pharmacol. 67, 473 (1980).

Index

ACE, s e e Angiotensin-converting enzyme a-N-Acetylendorphin HPLC, 131-132 processing, 131-133 ACTH, s e e Adrenocorticotropin Adrenocorticotropin, s e e a l s o Proopiomelanocortin HPLC, 134-135,212-213 processing, 83, 85, 89-91 radioimmunoassay, 135, 212-213 two-site immunometric assay, 150, 152-153 Affinity chromatography carboxypeptidase E, 247-248 proprotein convertase 1, 105 Alzheimer's disease /3-amyloid protein role, 281, 317 endopeptidase 24.15 role, 290 secretase role, 320-321 Amidation, s e e Peptidyl-glycine a-amidating monooxygenase Amyloid precursor protein biological function, 318 glycosylation, 318 homology with protease inhibitors, 317-318 immunoprecipitation, 322-323 phosphorylation, 318 point mutations, 317 processing site, 320-321 proteolytic processing, 281,290-291,294, 317318, 320 secretase, 320-321 size, 317 structure, 317-319 transfection of HeLa cells, 321-322 Western blotting, 323-325 fl-Amyloid protein neurotoxicity, 317 role in Alzheimer's disease, 281

Angiotensin antisera characterization, 332 preparation, 330 cleavage sites, 328-329 epitopes, 329-330 peptides acetylation, 332-333,338 extraction from biological samples, 336-340 lability, 333-336, 338-339 separation by HPLC, 333,338, 340 radioimmunoassay, 328-332, 338, 342 Angiotensin-converting enzyme autoradiography anatomical localization of enzyme, 389, 391 chronic inhibition studies, 396 drug penetration assessment, 393,395-397 film processing, 386-387 nonspecific binding, 387 quantitation of enzyme activity, 387, 389 quantitation of small brain nuclei, 393 radioligand sensitivity, 389 tissue preparation, 386 biological function, 383 inhibitors availabilitry, 273,384 characterization, 384-385 radiolabeling, 384 localization in brain, 391-393,397 membrane association, 344, 383 neurotensin processing, 273 size, 383 tissue distribution, 383-384 Antibody, s e e a l s o Two-site immunometric assay characterization by radioimmunoassay, 197-198, 210-211 generation, 196-197

399

400

INDEX

Antibody ( c o n ' t ) peptide haptens, 196-197 proprotein processing assays, 176-177 proinsulin, 177-179 proopiomelanocortin, 185, 187, 190-192 prosomatostatin, 179-185 protein requirements for production, 308 purification, 197 Antisense RNA assay systems, 112-113, 115-116 blocking of protein expression, 109-110 cell transfection, 117-119 cost of methods, 113 mechanism of action, 111-112 mismatch toleration, 113-114 Northern analysis, 115-116 plasmid preparation, 117 probe selection concentration, 113 sequence, 110, 112-115 size, 113, 117 promoter selection, 115 requirements for protein targeting, 112-113 selection of protein targets, 112 subcloning of stable cell lines, 119 APP, s e e Amyloid precursor protein Autoradiography, s e e Angiotensin-converting enzyme; Liquid emulsion autoradiography Baculovirus, protein expression system, 312 Bradykinin antisera characterization, 332 preparation, 330 cleavage sites, 328-329 peptides acetylation, 332-333,338 extraction from biological samples, 336-340 lability, 333-336, 338-339 separation by HPLC, 333,338, 340 radioimmunoassay, 328-332, 338, 342 Captopril, angiotensin-converting enzyme inhibition, 273,392 Carboxypeptidase E affinity chromatography, 247-248 assays fluorescence, 237-239, 241-244 Northern blot, 237-238,248 radioactive, 237, 244-246

sensitivity, 245 Western blot, 237-238, 248 expression in embryogenesis, 52-53 immunoprecipitation, 247 inhibitors, 238,244, 246 pH optimum, 238-239 precursor, 248 species distribution, 237 substrate specificity, 238-239, 245 Carboxypeptidase H, s e e Carboxypeptidase E Cathepsin D, processing of endothelin, 254 Chinese hamster ovary cell, protein overexpression system, 94-95,312 amplification, 98-99 collection of conditioned medium, 99-101 screening, 97-98 transfection, 95-96 CLIP, HPLC, 134-135 Cloning, s e e Endopeptidase 24.15, cloning Corticotropin-releasing factor, regulation of proopiomelanocortin, 61 COS cell, protein expression system, 312 DABTC, s e e Dimethylaminoazobenzine thiocarbamate Dansyl-Phe-Ala-Arg carboxypeptidase substrate, 238-239 protease assay, 241-244 synthesis, 239-241 Digoxigenin immunological detection, 31, 51-52 probe labeling, 20-21, 51 Dihydrofolate reductase methotrexate binding, 94-95 overexpression system, 94-95 Dimethylaminoazobenzine thiocarbamate, peptide derivatization, 171-172, 176 DNA, s e e Oligonucleotides ECE, s e e Endothelin-converting enzyme Embryo in s i t u hybridization, 46-47, 49-51 staging in rat, 49-50 Endopeptidase, insulin secretory granule type II, s e e Insulin secretory granule type II endopeptidase Endopeptidase 24.11 immunostaining antibody specificity, 350, 352

INDEX dual localization of antigens, 359-360 labeling immunogold, 356-359 peroxidase, 354-356 tissue preparation cryostat sections, 353 fixatives, 352-353 reagent penetration, 354 vibratome sections, 353-354 inhibitors, 386 mechanism, 344 membrane association, 344-346, 360 reconstitution, 346-347 subcellular fractionation, 347-348, 360 substrates, 345, 363 tissue distribution, 344-345 Endopeptidase 24.15 cloning amino acid sequence from pure enzyme, 301 antibody screening, 301-302 by homologous nucleic acid sequences, 302303 library preparation, 300-301 library screening oligonucleotide screening, 304-305 polymerase chain reaction, 305-307 sequence identification, 300-304 strategies, 299-300, 314 tissue selection, 300 related peptidases, 297-299 role in Alzheimer's disease, 290 secondary structural motifs, 303-304 solubility, 297 species distribution, 298 substrate specificity, 297, 363,372-375 tissue distribution, 297 Endopeptidase 24.16 antibody purification, 378 assays fluorimetric, 367 HPLC, 368-369 cleavage site specificity, 363-364 cloning, 381 distribution in central nervous system, 375378 immunoprecipitation, 379 inhibitors dipeptides, 370 phosphodiepryl 03,370, 372 immunoglobin G, 378-379

401 neurotensin processing, 370, 372-374 polyacrylamide gel electrophoresis, 379-380 purification, 378 substrate specificity, 363,365, 367, 372-375 Western blotting, 379-380 /3-Endorphin, s e e a l s o Proopiomelanocortin HPLC, 137-138, 212 processing, 85, 89-91 radioimmunoassay, 212 Endothelin biological function, 251 cloning, 252 homology with sarafotoxins, 252, 258 immunoassay, 259-260 inhibition of release, 256 processing, 252, 256-257 sequence, 257 Endothelin-converting enzyme aspartyl protease inhibition, 253 pH optimum, 253-254 size, 254 assay bioassay, 261-262 fluorescence energy transfer, 260 HPLC, 258-259 immunoassay, 259-260 scintillation proximity assay, 260-261 sensitivity, 260 cleavage site specificity, 253,256-257 expression in transfected cells, 262 intracellular localization, 252 kinetic parameters, 258 metalloprotease cloning, 256 glycosylation, 255 inhibitors, 255-256 pH optimum, 254-255 purification, 255-256 size, 255 pH dependence, 257 substrate recognition, 257-258 Enkephalin convertase, s e e Carboxypeptidase E ET, s e e Endothelin Expression vector affinity tag, 308-309 phenotype selection, 308 promoter, 308 ribosome-binding site, 308 transcription terminator, 309

402

INDEX

Fluorescence energy transfer, endothelin-converting enzyme assay, 260 Fluorogenic substrates, s e e a l s o Dansyl-Phe-AlaArg amino acid composition, 164-165 peptidase assay, 165-167, 238-239, 284-285, 287-288 solubility, 239 synthesis, 239-241 Formaldehyde, preparation of fixing solution, 25 Furin cleavage site specificity, 155-156 discovery, 3 expression in embryogenesis, 52-53 riboprobe synthesis, 22 sequence, 4 in situ hybridization, 37, 41-42 substrate specificity, 13-14, 16, 52 Gastrin, processing effects of hypersecretion, 273-274 parasecretory, 272 GEMSA, s e e Guanidinioethylmercaptosuccinic acid Glutathione S-transferase proprotein convertase fusion protein, 11-12 protein expression system, 311 Gonadotropin-releasing hormone, HPLC, 131-132 Granulosa, cell culture, 162-163 Guanidinioethylmercaptosuccinic acid, carboxypeptidase inhibition, 238, 244, 246, 289 High-performance liquid chromatography ion-exchange chromatography separation of peptidases, 291-292 reversed-phase chromatography analytical column types, 128-129 column size selection, 129-130 criteria of peptide purity, 128 flow rates, 129-130 ion pair strategies, 133-135 sample preparation prechromatography cleanup, 127-128 tissue extraction, 126-127, 201-203 separation of peptides, 125-126, 203-206, 209210, 287-288 solvent systems, 130-131,203,287 size-exclusion chromatography calibration curve, 137

separation of peptidases, 292-293 solvent systems, 135, 137 Histidine tag expression vectors, 309-310 protein purification, 313 HIV, s e e Human immunodeficiency virus HPLC, s e e High-performance liquid chromatography Human immunodeficiency virus, protease inhibitor development, 315 Hybridization, in s itu , s e e I n s itu hybridization Immunogold, labeling of ultrathin frozen sections, 191 Immunometric assay, two-site, s e e Two-site immunometric assay Immunostaining antibody specificity, 350, 352 dual localization of antigens, 359-360 labeling immunogold, 356-359 peroxidase, 354-356 proinsulin, 177-179 proopiomelanocortin, 185, 187, 190-192 prosomatostatin, 179-185 tissue preparation cryostat sections, 353 fixatives, 352-353 reagent penetration, 354 vibratome sections, 353-354 Inosine, nucleotide base pairing, 307 Ion-exchange chromatography, separation of peptidases, 291-292 I n situ hybridization, s e e a l s o Oligonucleotides; Riboprobes detection of hybridization signal liquid emulsion autoradiography, 29-31, 49 nonradioactive detection, 31, 51-52 X-ray film, 29 double-labeling techniques, 33, 51-52 embryo tissue, 46-47, 49-51 hybridization conditions, 27-28 mixing solution preparation, 27, 48-49 prehybridization treatment, 48 acetylation, 26 dehydration, 26 delipidation, 26 denaturation, 26 proteinase K treatment, 26

INDEX hybridization (con't) probe selection oligonucleotides, 18-19, 28 riboprobes, 17-18, 28, 50 processing enzyme localization, 35, 37, 41-42 quantitative densitometry computer analysis, 34 sampling, 35 standard curve construction, 34-35 sensitivity, 45 specificity controls negative, 32-33 positive, 32 subbing of slides gelatin coating, 24, 47 lysine coating, 24, 47 tissue fixation, 25 preparation, 24, 47 storage, 25 Insulin secretory granule type II endopeptidase, cleavage specificity, 90 In situ

Kexin cleavage site specificity, 155-156 discovery, 3 expression in recombinant vaccinia virus, 88 fluorescence assay, 165-166 kinetic parameters, 166-167 prohormone processing, 88 sequence, 4 substrate specificity, 165-166 fl-Lipotropin, s e e a l s o Proopiomelanocortin HPLC, 137-138 processing, 85, 89-91, 137 two-site immunometric assay, 150 Liquid emulsion autoradiography counterstaining, 30 developing, 30 dipping of slides, 30 emulsion preparation, 29-30 mounting, 30 photography, 30-31 fl-LPH, s e e fl-Lipotropin MalE system, protein expression system, 311-12 a-Melanocyte-stimulating hormone, s e e a l s o Proopiomelanocortin HPLC, 213-215 processing, 90 radioimmunoassay, 213-215

403 fl-Melanocyte-stimulating hormone, s e e a l s o Proopiomelanocortin HPLC, 212 radioimmunoassay, 212 Methotrexate application in overexpression systems, 94-95, 98 inhibition of dihydrofolate reductase, 94-95 Microdialysis, in v i v o flow rates, 275 osmotic pressure, 276 principles, 275 probes, 275-276 sampling at peptide secretion site, 274-275 MSH, s e e Melanocyte-stimulating hormone Multicatalytic endopeptidase, substrate specificity, 291 NacEP, s e e a-N-Acetylendorphin Neprilysin, s e e Endopeptidase 24.11 Neurolysin, s e e Endopeptidase 24.16 Neurophysin cleavage site, 158 fractionation, 159 isolation, 160-163 processing, 158-159, 163-164 radioimmunoassay, 163 synthetic peptide synthesis, 159-160 Neurotensin HPLC, 364, 367 parasecretory processing, 272-273 proteolytic processing, 363,381 Oligonucleotides labeling, 23-24 screening of cDNA libraries, 304-305 in situ hybridization, 28 synthesis, 18-19 Oxytocin, s e e a l s o Prooxytocin-neurophysin antibodies, 195 isolation, 163 peptides extraction from tissue, 198-199, 201-203 HPLC, 203,205-206 high-voltage electrophoresis, 199-200 radioimmunoassay, 203,206 processing, 158-159, 163-164 radioimmunoassay, 163 PACE4 reverse transcriptase-polymerase chain reaction, 8-9

404

INDEX

PACE4 ( c o n ' t ) sequence, 6-7 in s i t u hybridization, 37, 41-42 PAM, s e e Peptidyl-glycine a-amidating monooxygenase PC1, s e e Proprotein convertase 1 PC2, s e e Proprotein convertase 2 PC3, s e e Proprotein convertase 1 PC4, s e e Proprotein convertase 4 PC5, s e e Proprotein convertase 5 PCR, s e e Polymerase chain reaction Peptidases, central nervous system activators, 289 affinity chromatography, 283 biological function, 281,296 cloning, s e e Endopeptidase 24.15 fluorimetric assay, 284-285, 287-288 HPLC analysis of peptide degradation, 287-288 identification from cDNA libraries, 283 inhibitors, 289 mechanistic classes, 288-289, 296 pH optimum, 288-289, 292-293 polyacrylamide gel electrophoresis, 282-283 purification, 293-294 role in disease, 281 solubility, 296-297 substrate specificity, 281-282 synthetic peptide substrates size, 284 synthesis, 284-285 terminal blocking, 284 tissue collection brain fractionation, 286 peptidase stability, 285-286 preparation of synaptosomes, 286-287 Peptide processing, in v i v o animal models, 266-267 animal preparation, 267, 270 calculations half life, 269-270 metabolic clearance rate, 269 organ extraction, 270-271 production rate, 269 volume of distribution, 270 experimental design, 268-269 parasecretory processing, 272-273 pathways, 266 peptide infusion, 268 plasma stability, 271-272 sampling at secretion site, 274-276

Peptidyl-glycine a-amidating monooxygenase antisense RNA methods, 114, 116 assay applications, 230, 235 incubation time, 222 optimization, 225-227 principles, 219-223 product separation, 222-223 substrates, 220, 222 tissue preparation, 221-222 brain enzyme assay, 231 blood collection, 231 distribution of activity, 231-232, 235 hypothalamic enzyme kinetic parameters, 232233 plasma levels, 233 tissue homogenization, 230-231 cofactors, 222, 226-227 heart enzyme assay, 224-225 effect of chronic corticosteroid activity, 228230 subcellular distribution, 227-228 tissue collection, 223-224 tissue homogenization, 224 peptide substrates, 219 Phosphodiepryl 03, peptidase inhibition, 370, 372374 P i c h i a p a s t o r i s , protein expression system, 312 Polymerase chain reaction codon degeneracy in primer design, 305-306 primer selection, 9 size, 306 product purification, 306-307 proprotein convertases, 4-5, 8-9 screening cDNA libraries, 305-307 vaccinia virus recombinants, 81-82 POMC, s e e Proopiomelanocortin Preproopiomelanocortin, s e e Proopiomelanocortin Prodynorphin, overexpression system, 107 Proenkephalin glycosylation, 102-103 purification of overexpressed protein collection from medium, 100-101 reversed-phase chromatography, 102-103 solubility, 102 Progesterone, radioimmunoassay, 163

INDEX Proinsulin cleavage sites, 177-178 immunostain processing assay, 177-179 structure, 177-178 Proopiomelanocortin coexpression with processing enzymes, 88-89 epitopes, 152 expression in embryogenesis, 53, 55, 57 processing, 16, 83-86, 133, 152, 208-209 AtT-20 cells, 190-192 cleavage sites, 187, 190, 208 developmental changes, 209, 212-217 dopamine role in processing, 133 immunostain assay, 185, 187, 190-192 reconstitution of pathways, 87-90 subcellular localization, 191-192 tissue specificity, 83-86, 208 vaccinia virus recombinant protein, 83-86, 9091 regulation by corticotropin-releasing factor, 61 two-site immunometric assay, 150, 152-153 Prooxytocin-neurophysin cleavage sites, 158, 167-169, 196 fractionation, 159 processing assay, 169, 171-176 cleavage, 158-159, 163-164 purification, 174 radioimmunoassay, 162 synthetic peptide chromophore derivative, 171-172 synthesis, 159-160, 167-168 Proprotein convertase 1 activity assay, 103-104 aggregation in overexpression systems, 99-100 antisense RNA methods, 114 catalytic residues, 5, 7-8 coexpression with substrates, 12-14, 89 discovery, 4-5 expression in embryogenesis, 53, 55, 57, 216 expression in recombinant vaccinia virus, 88 fusion protein generation, l 1-12 purification of overexpressed protein affinity chromatography, 105 anion-exchange chromatography, 105-106 collection from medium, 100- l01 fast protein liquid chromatography, 103 hydrophobic interaction chromatography, 105 yield, 107

405 reverse transcriptase-polymerase chain reaction, 4 ribonuclease protection assay, 62-64 riboprobe synthesis, 22 sequence, 6-7 in situ hybridization, 37, 41-42 substrate specificity, 13-14, 16 Xenopus laeois gene cloning, 10 sequence, 11 Proprotein convertase 2 catalytic residues, 5, 7-8 cloning, Aplysia californica gene, 10 coexpression with substrates, 12-14, 89 discovery, 4-5 expression in embryogenesis, 216 expression in recombinant vaccinia virus, 88 fusion protein generation, 1l - 12 overexpression system, 107 reverse transcriptase-polymerase chain reaction, 4 ribonuclease protection assay, 62-64 riboprobe synthesis, 22 sequence, 6-7 in situ hybridization, 37, 41-42 substrate specificity, 13-14, 16 Proprotein convertase 4 catalytic residues, 5, 7-8 gene splicing, 14 reverse transcriptase-polymerase chain reaction, 8 sequence, 6-7 Proprotein convertase 5 catalytic residues, 5, 7-8 gene splicing, 14 reverse transcriptase-polymerase chain reaction, 8 riboprobe synthesis, 23 sequence, 6-7 in situ hybridization, 37, 41-42 Prosomatostatin cleavage sites, 179-180 immunostain processing assay, 179-185 processing L. piscatorius pancreatic islets, 18 l, 184-185 rat brain cortical cells, 179-180, 184 purification, 173 rat brain protein blockage of intracellular transport, 184 peptide identification, 183

406

INDEX

Prosomatostatin ( c o n "t) subcellular distribution, 182-183 subcellular fractionation, 182 Radioimmunoassay adrenocorticotropin, 135,212-213 angiotensin, 328-332, 338, 342 application with HPLC, 135, 138-139, 203-206, 212-217 bradykinin, 328-332, 338, 342 characterization of antibodies, 197-198, 210-211 fl-endorphin, 212 melanocyte-stimulating hormone, 212-215 progesterone, 163 proopiomelanocortin, 210-211 prooxytocin-neurophysin, 162-163,203,206 vasopressin, 203,206 RIA, s e e Radioimmunoassay Ribonuclease protection assay data analysis, 60 evaluation of gene expression, 57 gel electrophoresis, 59 hybridization mixture, 59 probe preparation, 58-60 RNA isolation, 58 tissue culture, 58 Riboprobes, s e e a l s o Antisense RNA hydrolysis, 21-22, 50 labeling nonradioactive, 20-21 radioactive, 19-20, 60 purification, 21, 50, 60-61 in s i t u hybridization, 28, 50 synthesis, 17-20, 22-23, 47-48, 60 RNA, antisense, s e e Antisense RNA Scintillation proximity assay, endothelin-converting enzyme assay, 260-261 Size-exclusion chromatography, s e e High-performance liquid chromatography, size-exclusion chromatography Somatostatin, HPLC, 131-132 Substance P, HPLC, 131-132 Synaptosome membrane isolation, 287 preparation from human brain, 286-287,348-350 Thyrotropin-releasing hormone, HPLC, 1311 3 2 t r p E fusion vector, protein expression system, 310-311

Two-site immunometric assay antibody characterization, 143-145 concentration, 148 generation, 143 radiolabeling, 145 selection, 140-142, 147 solid phase coupling, 145-146 antigen selection, 142, 152 assay format, 142-143 calibration, 151 incubation time, 149 optimization, 147-149 peptide interference, 150-151 principles, 140-142 sensitivity, 140 specificity, 149-150 stability of peptides, 151 standards, 146

Unwindase biological functions, 116 effect on antisense RNA experiments, 116

Vaccinia virus DNA isolation, 72, 74 expression vector amplification, 79 construction, 67-69 DNA preparation, 74 drug selection, 75-76 marker transfer protocol, 74-75 plaque purification agarose overlays, 76-78 filter lifts, 78-79 screening slot blot, 80 Southern analysis, 80-81 immunoblot, 81 polymerase chain reaction, 81-82 genome, 66-67 infection cell culture, 70 efficiency, 66, 83 large-scale preparation, 70-72 life cycle, 66-67 recombinant protein expression, 65-66, 83, 85, 87-91

INDEX reconstitution of prohormone processing pathways, 87-90 safety in handling, 69 titering of stock solutions, 72 Vasopressin antibodies, 195 peptides extraction from tissue, 198-199, 201-203 HPLC, 203,205-206 high-voltage electrophoresis, 199-200

407 radioimmunoassay, 203,206 processing, 195-196 Western blot amyloid precursor protein, 323-325 carboxypeptidase E, 237-238 endopeptidase 24.16, 379-380 X-ray crystallography, protein requirements, 308

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FIG.6 Computer-generated pseudocolor images of ACE distribution in coronal sections through a normal rat basal ganglia (A-D) and through a rat basal ganglia 2 weeks after N-methyl-D-aspartic acid lesion of the right caudate nucleus (E-H). In the lesioned brain, the right caudate putamen (CPu), globus pallidus (GP), entopeduncular nucleus (EP), and substantia nigra pars recticulata (SNR) contain appreciably less ACE than the structures on the left side or in the control brain.