Protein Phosphatases

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Preface

Reversible phosphorylation affects proteins in all compartments of a cell and is crucial for life and death. It is therefore not surprising that this topic has been covered three times already in the Methods in Enzymology series: (1) in 1983 in Volume 99 on protein kinases, and (2)/(3) in 1991 in Volumes 200 and 201 (Part A and Part B) on protein phosphorylation. Much has happened since the last volume on protein phosphorylation was published in Methods in Enzymology. Recognition of the fundamental importance of reversible protein phosphorylation in cellular processes culminated in the awarding of the 1992 Nobel Prize for Physiology or Medicine to Edmond Fischer and Edwin Krebs for their discovery of the regulation of phosphorylase by phosphorylation in the early 1950s. In the year 2000, Paul Greengard, Eric Kandel, and Arvid Carlson received the Nobel Prize for their findings concerning signal transduction—including phosphorylation—in the nervous system. Finally, Timothy Hunt, Leland Hartwell, and Paul Nurse were awarded the Nobel Prize in 2001 for their achievements in studying cell cycle control (discovery of cyclins and cyclindependent kinases). This volume deals with protein phosphatases exclusively. The phosphatases had been overlooked for a long time. However, it is obvious by now that they are as equally important as the kinases, and the field is expanding rapidly. A major development over the past 10 years has been the realization that serine/threonine phosphatases acquire specificity and are controlled by association with targeting polypeptides that not only localize the enzymes to different cellular compartments, but also impart regulation and substrate recognition. To date, many phosphatase-binding proteins (including subunits, anchoring proteins, and regulatory proteins) have been identified. This applies for most of the beloved oldies such as type-1, type2A, etc., that continue to be hot spots in science. Protein tyrosine phosphatases that emerged much later also advanced to become major targets for further research in recent years. They have revealed an enormous structural diversity, and the ability of some of the protein tyrosine phosphatases also to act as a lipid phosphatase is most stunning. A novelty is the third class of phosphatases acting on labile, N-phosphorylated amino acids such as histidine: The first vertebrate histidine phosphatase has been discovered. Once more, technology and methodology have greatly advanced over the past years. Overexpression on the one side and downregulation or even

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PREFACE

knock out on the other side are just a few examples of what has emerged and what has been used in the field of protein phosphatases as represented in this volume. It is our pleasure to thank the authors for their cooperation, Drs. M. I. Simon and J. N. Ableson (the editors-in-chief) for their confidence, and the publisher for their generous support, thus making this project possible. SUSANNE KLUMPP JOSEF KRIEGLSTEIN

Table of Contents

CONTRIBUTORS TO VOLUME 366 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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PREFACE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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VOLUMES IN SERIES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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FOREWORD

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BY

PHILIP COHEN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Section I. Determination, Detection, and Localization 1. Monitoring of PP2A and PP2C by Phosphothreonyl Peptide Substrates

ARIANNA DONELLA-DEANA, MARCO BOSCHETTI, AND LORENZO A. PINNA

3

ANNA A. DEPAOLI-ROACH, PIER GIUSEPPE VILARDO, JONG-HWA KIM, NIRMALA MAVILA, BHARGAVI VEMURI, AND PETER J. ROACH

17

3. Measuring Protein Phosphatase Activity with Physiological Substrates

BO ZHOU AND ZHONG-YIN ZHANG

34

4. PTEN and Myotubularins: Families of Phosphoinositide Phosphatases

GREGORY S. TAYLOR AND JACK E. DIXON

43

5. Detection of Protein Histidine Phosphatase in Vertebrates

SUSANNE KLUMPP, JAN HERMESMEIER, JOSEF KRIEGLSTEIN

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2. Determination of Mammalian Glycogen Synthase Phosphatase Activity

AND

C. PETER DOWNES, ALEXANDER GRAY, STEPHEN A. WATT, AND JOHN M. LUCOCQ

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7. Mixed Peptide Sequencing and the FASTF/ FASTS Algorithms

RUPA RAY AND TIMOTHY A. J. HAYSTEAD

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8. Phosphoproteome Analysis in Yeast

RUPA RAY AND TIMOTHY A. J. HAYSTEAD

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6. Advances in Procedures for the Detection and Localization of Inositol Phospholipid Signals in Cells, Tissues, and Enzyme Assays

9. cDNA Microarray Analysis Reveals an Overexpression of the Dual-Specificity MAPK Phosphatase PYST2 in Acute Leukemia

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ORLEV LEVY-NISSENBAUM, ORIT SAGI-ASSIF, PIA RAANANI, ABRAHAM AVIGDOR, ISAAC BEN-BASSAT, AND ISAAC P. WITZ

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10. Protein Phosphatase Translocation in RBL-2H3 Cells

ALISTAIR T. R. SIM, JEFF HOLST, AND RUSSELL I. LUDOWYKE

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11. Whole Mount Analysis of Mammary Gland Structure in PTP Epsilon Transgenic Mice

ZOHAR TIRAN AND ARI ELSON

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Section II. Interacting Proteins and Subunits 12. Assay of Protein Phosphatase 1 Complexes

13. Validation of Interactions Phosphatase-1

with

Protein

14. Analysis of Specific Interactions of Native Protein Phosphatase 1 Isoforms with Targeting Subunits

15. Using the Ras Recruitment System to Identify PP2A–B55-Interacting Proteins 16. Altering the Holoenzyme Composition and Substrate Specificity of Protein Phosphatase 2A

PATRICIA T. W. COHEN, GARETH J. BROWNE, MIRELA DELIBEGOVIC, AND SHONAGH MUNRO

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ALEYDE VAN EYNDE AND MATHIEU BOLLEN

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ROGER J. COLBRAN, LEIGH C. CARMODY, PATRICIA A. BAUMAN, BRIAN E. WADZINSKI, AND MARTHA A. BASS

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HAIM M. BARR, RAKEFET SHARF, AND TAMAR KLEINBERGER

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THOMAS FELLNER, PATRICK PIRIBAUER, AND EGON OGRIS

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00

17. The Application of Fluorescence Resonance Energy Transfer to the Investigation of Phosphatases

JA´NOS SZO¨LLOSI AND DENIS R. ALEXANDER

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18. Receptor Protein-Tyrosine Phosphatase Dimerization

JEROEN DEN HERTOG, THEA VAN DER WIJK, LEON G. J. TERTOOLEN, AND CHRISTOPHE BLANCHETOT

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Section III. Inhibition, Stimulation, and Modulation of Activity 19. Phosphoprotein Inhibitors of Protein Phosphatase-1

MASUMI ETO, CRAIG LEACH, NIKOLAOS A. TOUNTAS, AND DAVID L. BRAUTIGAN

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20. Combinatorial Chemistry and Peptide Library Methods to Characterize Protein Phosphatases

STEFAN W. VETTER AND ZHONG-YIN ZHANG

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21. Activity of PP2C is Increased by Divalent Cations and Lipophilic Compounds Depending on the Substrate

JOSEF KRIEGLSTEIN, DAGMAR SELKE, ALEXANDER MAAßEN, SUSANNE KLUMPP

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AND

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TABLE OF CONTENTS

22. Regulation of Calcineurin by Oxidative Stress

23. Analysis of the Regulation of Protein Tyrosine Phosphatases in Vivo by Reversible Oxidation

MANIK C. GHOSH, XUTONG WANG, SHIPENG LI, AND CLAUDE KLEE

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TZU-CHING MENG AND NICHOLAS K. TONKS

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Section IV. Expression Systems 24. Preparation and Characterization of Recombinant Protein Phosphatase 1

TAKUO WATANABE, EDGAR F. DA CRUZ E SILVA, HSIEN-BIN HUANG, NATALIA STARKOVA, YOUNG-GUEN KWON, ATSUKO HORIUCHI, PAUL GREENGARD, AND ANGUS C. NAIRN

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25. An Inducible System to Study the Growth Arrest Properties of Protein Phosphatase 2C

PAULA OFEK, DANIELLA BEN-MEIR, AND SARA LAVI

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26. Use of Tetracycline-Regulatable Promoters for Functional Analysis of Protein Phosphatases in Yeast

JOAQUI´N

ARIN˜O AND

ENRIC HERRERO

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Section V. Knockdown and Knockout Technologies 27. Analysis of Protein Phosphatase Function in Drosophila Cells Using RNA Interference

ADAM M. SILVERSTEIN MARC C. MUMBY

28. Regulating the Expression of Protein Phosphatase Type 5

TERESA A. GOLDEN AND RICHARD E. HONKANEN

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29. Transgenic and Knockout Models of PP2A

JU¨RGEN GO¨TZ AND ANDREAS SCHILD

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30. Saccharomyces Gene Deletion Project: Applications and Use in the Study of Protein Kinases and Phosphatases

WAYNE A. WILSON PETER J. ROACH

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AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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SUBJECT INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors to Volume 366 Article numbers are in parentheses following the names of contributors. Affiliations listed are current.

DENIS R. ALEXANDER (17), Laboratory of Lymphocyte Signalling and Development, Molecular Immunology Programme, The Babraham Institute, Cambridge CB2 4AT, UK

MARCO BOSCHETTI (1), Dipartimento di Chimica Biologica, University of Padova, Viale G. Colombo 3, Padova 35121, Italy DAVID L. BRAUTIGAN (19), Center for Cell Signaling and Department of Microbiology, University of Virginia, School of Medicine, Charlottesville, Virginia 22908

JOAQUI´N ARIN˜O (26), Departament de Bioquı´mica i Biologia Molecular, Universitat Auto`noma de Barcelona Bellaterra, Barcelona, 08193, Spain

GARETH J. BROWNE (12), Medical Research Council Protein Phosphorylation Unit, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland, UK

ABRAHAM AVIGDOR (9), Institute of Hematology, The Chaim Sheba Medical Center, Tel-Hashomer, Israel, and Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, 69978, Israel

LEIGH C. CARMODY (14), Room 702, Light Hall, Vanderbilt University Medical Center, Nashville, Tennessee 37232-0615

HAIM M. BARR (15), Gonda Center for Molecular Biology, B. Rappaport Faculty of Medicine, Technion-Israel Institute of Technology, P.O.Box 9649, Bat-Galim, Haifa, 31096, Israel

PATRICIA T. W. COHEN (12), MRC Protein Phosphorylation Unit, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland, UK

MARTHA A. BASS (14), Room 702, Light Hall, Vanderbilt University Medical Center, Nashville, Tennessee 37232-0615

PHILIP COHEN (Foreword), MRC Protein Phosphorylation Unit, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland, UK

PATRICIA A. BAUMAN (14), Room 702, Light Hall, Vanderbilt University Medical Center, Nashville, Tennessee 37232-0615 ISAAC BEN-BASSAT (9), Institute of Hematology, The Chaim Sheba Medical Center, TelHashomer, Israel, and Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, 69978, Israel

ROGER J. COLBRAN (14), Room 702, Light Hall, Vanderbilt University Medical Center, Nashville, Tennessee 37232-0615 EDGAR F. DA CRUZ E SILVA (24), Centro de Biologia Celular, Universidade de Aveiro, 3810-193 Aveiro, Portugal

DANIELLA BEN-MEIR (25), Department of Cell Research and Immunology, Tel Aviv University, Tel Aviv, 69978, Israel CHRISTOPHE BLANCHETOT (18), McGill University, McIntyre Medical Sciences Building, 3655 Drummond Street, Montreal, Quebec, H3G 1Y6, Canada

MIRELA DELIBEGOVIC (12), Medical Research Council Protein Phosphorylation Unit, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland, UK

MATHIEU BOLLEN (13), Afdeling Biochemie, Faculteit Geneeskunde, Katholieke Universiteit Leuven, Herestraat 49, B-3000 Leuven, Belgium

JEROEN DEN HERTOG (18), Hubrecht Laboratory, Netherlands Institute for Developmental Biology, Uppsalalaan 8, Utrecht, 3584 CT, The Netherlands

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CONTRIBUTORS

ANNA A. DEPAOLI-ROACH (2), Department of Biochemistry and Molecular Biology, and Center for Diabetes Research, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, Indiana 46202 JACK E. DIXON (4), Departments of Pharmacology, Cellular and Molecular Medicine, Chemistry and Biochemistry, University of California, Basic Science Building 3092A-M/C 0636, 9500 Gilman Drive, La Jolla, California 92093-0636 ARIANNA DONELLA-DEANA (1), Dipartimento di Chimica Biologica, University of Padova, Viale G. Colombo 3, Padova 35121, Italy C. PETER DOWNES (6), School of Life Sciences, MSI/WTB Complex, Dow Street, University of Dundee, Dundee DD1 5EH, UK ARI ELSON (11), Department of Molecular Genetics, The Weizmann Institute of Science, Rehovot, 76100, Israel MASUMI ETO (19), Center for Cell Signaling and Department of Molecular Physiology and Biological Physics, University of Virginia, School of Medicine, Charlottesville, Virginia 22908 THOMAS FELLNER (16), Institute of Medical Biochemistry, Division of Molecular Biology, Vienna Biocenter, University of Vienna, Dr. Bohr-Gasse 9, Vienna A-1030, Austria MANIK C. GHOSH (22), Cell Biology and Metabolism Branch, National Institute of Child Health and Human Development, NIH, Bethesda, Maryland 20892 TERESA A. GOLDEN (28), Department of Biochemistry & Molecular Biology, College of Medicine, University of South Alabama, Mobile, Alabama 36688 JU¨RGEN GO¨TZ (29), Division of Psychiatry Research, University of Zurich, August Forel Str 1, Zurich CH 8008, Zu¨rich, Switzerland

TIMOTHY A. J. HAYSTEAD (7,8), Chief Scientific Founder, Serenex Inc., Durham, North Carolina 27710 JAN HERMESMEIER (5) Institut fu¨r Pharmazeutische & Medizinische Chemie, Westfa¨lische Wilhelms-Universita¨t, Hittorfstr. 58-62, 48149 Mu¨nster, Germany ENRIC HERRERO (26), Departament de Cie`ncies Me`diques Ba`siques, Universitat de Lleida, Lleida, 25198, Spain JEFF HOLST (10), Centre for Immunology, St. Vincent’s Hospital, University of New South Wales, Sydney, New South Wales, Australia RICHARD E. HONKANEN (28), Department of Biochemistry and Molecular Biology, MSB 2198, University of South Alabama, Mobile, Alabama 36688 ATSUKO HORIUCHI (24) Laboratory of Molecular and Cellular Neuroscience, The Rockefeller University, 1230 York Avenue, New York, New York 10021 HSIEN-BIN HUANG (24), Institute of Molecular Biology, National Chung Cheng University, Chia-Yi 621, Taiwan, ROC JONG-HWA KIM (2), Department of Biochemistry and Molecular Biology, and Center for Diabetes Research, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, Indiana 46202 CLAUDE KLEE (22), Laboratory of Biochemistry, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892 TAMAR KLEINBERGER (15), Gonda Center for Molecular Biology, B. Rappaport Faculty of Medicine, Technion-Israel Institute of Technology, P.O.Box 9649, Bat-Galim, Haifa, 31096, Israel

ALEXANDER GRAY (6), School of Life Sciences, MSI/WTB Complex, Dow Street, University of Dundee, Dundee DD1 5EH, UK

SUSANNE KLUMPP (5,21) Institut fu¨r Pharmazeutische & Medizinische Chemie, Westfa¨lische Wilhelms-Universita¨t, Hittorfstrasse 58-62, 48149 Mu¨nster, Germany

PAUL GREENGARD (24), Laboratory of Molecular and Cellular Neuroscience, The Rockefeller University, 1230 York Avenue, New York, New York 10021

JOSEF KRIEGLSTEIN (5,21) Institut fu¨r Pharmakologie und Toxikologie, PhilippsUniversita¨t, Ketzerbach 63, 35032 Marburg, Germany

CONTRIBUTORS

YOUNG-GUEN KWON (24), Department of Biochemistry, College of Natural Sciences, School of Medicine, Kangwon National University, Chunchon, Kangwon-Do, 200-701, Korea SARA LAVI (25), Department of Cell Research and Immunology, Tel Aviv University, Tel Aviv, 69978, Israel CRAIG LEACH (19), Center for Cell Signaling and Department of Microbiology, University of Virginia, School of Medicine, Charlottesville, Virginia 22908 ORLEV LEVY-NISSENBAUM (9), Department of Cell Research and Immunology, George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, 69978, Israel SHIPENG LI (22), Laboratory of Biochemistry, National Cancer Institute National Institutes of Health, Bethesda, Maryland 20892 JOHN M. LUCOCQ (6), School of Life Sciences, MSI/WTB Complex, Dow Street, University of Dundee, Dundee DD1 5EH, UK RUSSELL I. LUDOWYKE (10), Proteome Systems Ltd, 1/35-41 Waterloo Road, North Ryde, NSW 2113, Australia

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ANGUS C. NAIRN (24), Department of Psychiatry, Yale University School of Medicine, New Haven, Connecticut 06508 PAULA OFEK (25), Department of Cell Research and Immunology, Tel Aviv University, Tel Aviv, 69978, Israel EGON OGRIS (16), Institute of Medical Biochemistry, Division of Molecular Biology, Vienna Biocenter, University of Vienna, Dr. Bohr-Gasse 9, Vienna A-1030, Austria LORENZO A. PINNA (1), Dipartimento di Chimica Biologica, University of Padova, Viale G. Colombo 3, Padova 35121, Italy PATRICK PIRIBAUER (16), Institute of Medical Biochemistry, Division of Molecular Biology, Vienna Biocenter, University of Vienna, Dr. Bohr-Gasse 9, Vienna A-1030, Austria PIA RAANANI (9), Institute of Hematology, The Chaim Sheba Medical Center, Tel-Hashomer, Israel, and Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, 69978, Israel RUPA RAY (7,8), Department of Pharmacology and Cancer Biology, Duke University, Research Drive, C118 LSRC, Durham, North Carolina 27710-3686

ALEXANDER MAAßEN (21), Institut fu¨r Pharmazeutische & Medizinische Chemie, Westfa¨lische Wilhelms-Universita¨t, Hittorfstr. 58-62, 48149 Mu¨nster, Germany

PETER J. ROACH (2,30), Department of Biochemistry and Molecular Biology, and Center for Diabetes Research, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, Indiana 46202

NIRMALA MAVILA (2), Department of Biochemistry and Molecular Biology, and Center for Diabetes Research, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, Illinois 46202

ORIT SAGI-ASSIF (9) Department of Cell Research and Immunology, George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, 69978, Israel

TZU-CHING MENG (23), Cold Spring Harbor Laboratory, 1 Bungtown Road, Cold Spring Harbor, New York 11724

ANDREAS SCHILD (29), Division of Psychiatry Research, University of Zurich, August Forel Str 1, Zurich CH 8008, Zu¨rich, Switzerland

MARC C. MUMBY (27), Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, Texas 75390-9041 SHONAGH MUNRO (12), Medical Research Council Protein Phosphorylation Unit, School of Life Sciences MSI/WTB Complex, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland, UK

RAKEFET SHARF (15), Gonda Center for Molecular Biology, B. Rappaport Faculty of Medicine, Technion-Israel Institute of Technology, P.O. Box 9649, Bat-Galim, Haifa, 31096, Israel DAGMAR SELKE (21), Institut fu¨r Pharmazeutische & Medizinische Chemie, Westfa¨lische Wilhelms-Universita¨t, Hittorfstr. 58-62, 48149 Mu¨nster, Germany

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CONTRIBUTORS

ADAM M. SILVERSTEIN (27), Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, Texas 75390-9041 ALISTAIR T. R. SIM (10), School of Biomedical Sciences, University of Newcastle and Clinical Neuroscience Program, Hunter Medical Research Institute, Callaghan, NSW 2308, Australia NATALIA STARKOVA (24), Laboratory of Molecular and Cellular Neuroscience, The Rockefeller University, 1230 York Avenue, New York, New York 10021 00

JA´NOS SZO¨LLOSI (17), Department of Biophysics and Cell Biology, Faculty of Medicine, Medical and Health Science Center, University of Debrecen, Debrecen H-4012, Hungary GREGORY S. TAYLOR (4), Departments of Pharmacology, Cellular and Molecular Medicine, Chemistry and Biochemistry, University of California, Basic Science Building 3092A-M/C 0636, 9500 Gilman Drive, La Jolla, California 92093-0636 LEON G. J. TERTOOLEN (18), Hubrecht Laboratory, Netherlands Institute for Developmental Biology, Uppsalalaan 8, Utrecht, 3584 CT, The Netherlands ZOHAR TIRAN (11), Department of Molecular Genetics, The Weizmann Institute of Science, Rehovot, 76100, Israel NICHOLAS K. TONKS (23), Cold Spring Harbor Laboratory, 1 Bungtown Road, Cold Spring Harbor, New York 11724 NIKOLAOS A. TOUNTAS (19), Center for Cell Signaling and Department of Microbiology, University of Virginia, School of Medicine, Charlottesville, Virginia 22908 THEA VAN DER WIJK (18), Hubrecht Laboratory, Netherlands Institute for Developmental Biology, Uppsalalaan 8, Utrecht, 3584 CT, The Netherlands ALEYDE VAN EYNDE (13), Afdeling Biochemie, Faculteit Geneeskunde, Katholieke Universiteit Leuven, Herestraat 49, B-3000 Leuven, Belgium BHARGAVI VEMURI (2), Department of Biochemistry and Molecular Biology, and

Center for Diabetes Research, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, Indiana 46202 STEFAN W. VETTER (20), Department of Molecular Biology, The Scripps Research Institute, 10550 North Torrey Pines Blvd, La Jolla, California 92037 PIER GIUSEPPE VILARDO (2), Department of Biochemistry and Molecular Biology, and Center for Diabetes Research, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, Indiana 46202 BRIAN E. WADZINSKI (14), Room 702 Light Hall, Vanderbilt University Medical Center, Nashville, Tennessee 37232-0615 XUTONG WANG (22), Board of Governors, Federal Reserve System, 20th & C Street, Washington, District of Columbia 20551 TAKUO WATANABE (24), Department of Biochemistry and Molecular Vascular Biology, Kanazawa University Graduate School of Medical Science, 13-1, Takara-machi, Kanazawa 920-8640, Japan STEPHEN A. WATT (6), School of Life Sciences, MSI/WTB Complex, Dow Street, University of Dundee, Dundee DD1 5EH, UK WAYNE A. WILSON (30), Department of Biochemistry and Molecular Biology, and Center for Diabetes Research, Indiana University School of Medicine, Indianapolis, Indiana 46202 ISAAC P. WITZ (9), Department of Cell Research and Immunology, George S. Wise Faculty of Life Sciences, and The Ela Kodesz Institute for Research on Cancer Development and Prevention, Tel Aviv University, Tel Aviv, 69978, Israel ZHONG-YIN ZHANG (3,20), Department of Molecular Pharmacology, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Bronx, New York 10461 BO ZHOU (3), Department of Molecular Pharmacology, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Bronx, New York 10461

Foreword The Past and Future of Protein Phosphatase Research

I have had relatively little involvement with protein phosphatases over the past 10 years. Nevertheless, as it is still a subject that is dear to my heart, I was delighted to accept the Editors’ invitation to write this foreword. It has given me an opportunity to take a nostalgic look back at the early days of the field when protein phosphatases were my major preoccupation, as well as to consider what the future may hold. The first part is therefore a personal account of the events that led my laboratory to classify the serine/threonine protein phosphatases 20 years ago and an outline of the discoveries I have been most fascinated by over the past 20 years. A more detailed historical account containing the original references can be found elsewhere.1 The second half of this foreword contains a few thoughts about how the field of protein phosphatases might develop.

The Past

The study of protein phosphatases, like that of protein phosphorylation, originated from studies of the control of glycogen metabolism. Over 60 years ago, Carl and Gerty Cori found that glycogen phosphorylase could be isolated in two forms termed phosphorylase b and phosphorylase a. Since the b-form required 50 AMP for activity but the a-form was almost fully active without AMP, they reasoned (incorrectly) that the a-form must contain tightly bound 50 -AMP, and that the a to b converting enzyme they had discovered, must catalyze the release of 50 AMP from the a-form. It was only many years later, with the discovery by Edmond Fischer and Edwin Krebs that the conversion of phosphorylase b to a involved a phosphorylation event, that it became obvious that the a to b converting enzyme was a protein phosphatase. My own involvement with protein phosphatases began in 1972 when my first graduate student, John Antoniw, began to isolate the protein phosphatase that dephosphorylated and inactivated phosphorylase kinase in skeletal muscle, the enzyme that converts phosphorylase b to a. Surprisingly, John resolved the activity into two enzymes, one of which dephosphorylated the -subunit of the enzyme much faster than the 1

P. Cohen, Bioessays 16, 583–588 (1994).

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-subunit, the other being highly specific for the -subunit. Subsequently, John showed that the phosphatase specific for the -subunit was also the major phosphatase activity in skeletal muscle acting on glycogen synthase and glycogen phosphorylase and was therefore the original a to b converting enzyme. He and a postdoctoral fellow, Gillian Nimmo, also showed that this phosphatase was inhibited by two heat-stable proteins termed inhibitor 1 and inhibitor 2 that Walter Glinsmann had identified in 1976. The phosphatase was therefore renamed protein phosphatase-1 (PP1) to reflect the fact that it had a broad specificity and tissue distribution, and was likely to dephosphorylate many regulatory proteins in vivo. The phosphatase that dephosphorylated the -subunit was insensitive to inhibitors 1 and 2 and initially termed protein phosphatase 2 (PP2). It was only several years later that another graduate student, Lex Stewart, discovered that it was dependent on calcium ions and calmodulin for activity and, in collaboration with Claude Klee, that it was identical to calcineurin, a major calmodulin-binding protein in the brain of previously unknown function. Studies by a postdoctoral fellow, Tom Ingebritsen, and a graduate student, Gordon Foulkes, identified a protein phosphatase in the liver that, like PP2 from muscle, dephosphorylated the -subunit of phosphorylase kinase much faster than the -subunit and was insensitive to inhibitors 1 and 2. However, in contrast to PP2 from muscle, the hepatic protein phosphatase had a much higher activity towards phosphorylase. It was therefore termed PP2A to distinguish it from the muscle PP2, which was renamed PP2B. A Mg2 þ -dependent protein phosphatase that had been identified in several laboratories and purified to homogeneity by Shigeru Tsuiki in 1981 was also found to dephosphorylate the -subunit preferentially, was insensitive to inhibitors 1 and 2, and we therefore termed it PP2C. As a result of further investigations by Tom Ingebritsen, Gordon Foulkes and others in the laboratory, it became obvious that PP1, PP2A, PP2B and PP2C accounted for most, if not all, of the serine/threoninespecific protein phosphatases towards many regulatory proteins involved in controlling a variety of cellular processes. Although the publication of our ideas in 1983 was initially greeted with skepticism in some quarters, they came to be accepted because the simple methods introduced to distinguish each protein serine/threonine phosphatase enabled other investigators to rapidly classify the phosphatase that they had been studying. Over the past 20 years, there has been huge progress in understanding the structure, substrate specificity and regulation of these and many other protein phosphatases. One crucially important advance was the isolation and characterization of the first protein tyrosine phosphatase (PTPase) by

FOREWORD

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one of my former graduate students Nick Tonks, when he joined Edmond Fischer as a postdoctoral fellow. The PTPase subfamily encompasses many enzymes, including transmembrane receptor-like molecules whose activities are likely to be modulated by as yet unidentified ligands, and ‘‘dual specificity phosphatases that play key roles in the regulation of MAP kinase cascades and the cell division cycle.’’ The amino acid sequence of every human protein phosphatase is probably now known and structures of members of each subclass have been solved at atomic resolution by X-ray crystallography. We now know that the isoforms of PP1, PP2A and PP2B are members of the same subfamily (now known as the PPP subfamily), while the PP2Cs are structurally unrelated (the PPM family). A third subfamily of protein serine/threonine phosphatases (the FCP family), of which there are at least five members, was only identified in the late 1990s by Jack Greenblatt and his colleagues.2 The amino acid sequences of the FCP phosphatases are unrelated to either the PPP or PPM subfamilies or to the PTPases. Although Harry Matthews has shown that members of the PPP and PPM subfamilies dephosphorylate histidine residues in proteins very efficiently, the first eukaryotic protein histidine phosphatase that is unable to dephosphorylate serine and threonine residues was recently identified by Susanne Klumpp and her colleagues.3 The structure of this protein histidine phosphatase is also unrelated to other protein phosphatases. Thus there are at least five structurally distinct subfamilies of protein phosphatases in eukaryotic cell, or more if there are separate phosphatases that dephosphorylate lysine and arginine residues in proteins. We now appreciate that protein kinases and protein phosphatases do not find their substrates by simple diffusion, but are directed to specific locations by targeting subunits and/or by the presence of ‘‘docking’’ sites located on the phosphatases and their substrates. This has become an enormous area of research in which the protein phosphatases can be said to have paved the way, starting with the finding in 1985 by Peter Stralfors, a postdoctoral fellow in my laboratory, that PP1 is directed to glycogen by a specific glycogen-targeting subunit. A surprising development at the time was the discovery of connections between protein phosphatases and human disease, starting with the report by Akira Takai in 1987 that okadaic acid was a potent inhibitor of PP1 and PP2A. This complex polyketal is a major toxic component associated with diarrhetic seafood poisoning, which has poisoned thousands of people and 2

J. Archambault, G. Pan, G. K. Dahmus, M. Cartier, N. F. Marshall, S. Zhang, M. E. Dahmus, and J. Greenblatt, J. Biol. Chem. 273, 27593–27601 (1998). 3 S. Klumpp, J. Hermesmeier, D. Selke, R. Baumeister, R. Kellner, and J. Krieglstein, J. Cereb. Blood Flow Metab. 22, 1420–1424 (2002).

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damaged the shellfish industries of many countries. Microcystins were subsequently found to be even more potent inhibitors of PP1 and PP2A. These cyclic heptapeptides are liver toxins produced by blue-green algae and the most potent liver carcinogens known to man. A plethora of other naturally occurring inhibitors of PP1 and PP2A have been identified subsequently. A further interesting connection between protein phosphatases and cancer was made by Gernot Walter, who found that the small T-antigens of simian virus 40 and polyomavirus, which enhance the transformation of mammalian cells, form complexes with a particular species of PP2A. Marc Mumby showed that this decreases their activity towards some substrates, explaining their oncogenicity. No less remarkable was the discovery by Stuart Schreiber in 1991 that cyclosporin and FK506, when bound to immunophilins, are potent and specific inhibitors of PP2B. These immunosuppressants, which have permitted the widespread use of organ transplantation, block the production of interleukin-2 and hence the proliferation of T cells. Remarkable connections between PTPases and disease were also made by Jack Dixon in the early 1990s. He showed that bacteria of the genus Yersinia produce a PTPase which is a virulence factor for several diseases, including pseudotuberculosis, a range of gastrointestinal syndromes and the bubonic plague. The last mentioned has been responsible for several pandemics over the past millennium, the most recent being the ‘‘Black Death’’ which killed no less than a quarter of the population of Europe in the 17th century.

The Future

So what does the future of protein phosphatase research hold and what are the key issues that still need to be resolved? One of the most challenging problems in the postgenomic era will be to identify all the physiological substrates of each protein phosphatase. This is a formidable task bearing in mind that about a third of the 30,000 proteins encoded by the human genome are phosphorylated frequently at multiple sites. Since about 150 protein phosphatases are encoded by the human genome, one can calculate that ‘‘on average’’ a protein phosphatase must have 60–70 substrates in vivo. However, this is an underestimate since many substrates, like phosphorylase kinase, are dephosphorylated by two or more phosphatases. Clearly, powerful new methodologies will be needed to tackle this problem, such as the substrate trapping mutants of PTPases developed by Nick Tonks, which are described in one of the chapters of this volume.

FOREWORD

xlix

Disruption or mutation of the genes encoding some protein phosphatases has been useful in defining some of the physiological processes that they regulate. For example, Mitsuhiro Yanagida showed that the mutation of PP1 in yeast caused chromosome separation, while Tricia Cohen showed that the disruption of PP4 in Drosophila caused defective nucleation of microtubules from centrosomes. However, inactivating a protein phosphatase catalytic subunit may frequently not be that helpful in identifying the direct substrates of a protein phosphatase in vivo. One reason for this is that many protein kinases are activated by phosphorylation. Therefore suppressing the activity of a phosphatase, whether by gene disruption, RNAi or the use of a specific inhibitor, will activate many protein kinases and therefore lead to the hyperphosphorylation of proteins that are not themselves direct substrates of the phosphatase under investigation. A more effective approach may therefore be to prevent the expression of a particular targeting or regulatory subunit. For example, Anna de Paoli-Roach and Tricia Cohen have both generated mice that do not express the gene encoding the GM glycogen-targeting subunit of PP1, providing genetic evidence that the PP1–GM complex is indeed the major phosphatase acting on glycogen synthase and glycogen phosphorylase in muscle. Extending this approach to other targeting subunits will undoubtedly aid the identification of other substrates of PP1, although ‘‘knock-in’’ mutations expressing a modified targeting subunit in which the PP1-interacting ‘‘RVXF’’ motif is disabled may be even better. Methodological problems associated with studying PP1-targeting subunits complexes are addressed in Section II of this volume. Many naturally occurring inhibitors of the PPP subfamily have been identified and exploited to demonstrate that a number of physiological processes are regulated by reversible phosphorylation. However, their usefulness is limited, because they all inhibit several PPP family members. Despite the caveats outlined earlier, far more specific inhibitors would be valuable. The recent report from Kunimi Kikuchi and his colleagues that tautomycetin is a relatively specific inhibitor of PP1 may therefore be a significant development.4 Finally, what are the chances that specific inhibitors of protein phosphatases will be developed for the treatment of disease? Enthusiasts of this idea received a major boost five years ago when Michel Trembly reported that mice deficient in PTP1B were not only hypersensitive to insulin, but also did not gain weight when fed on a high carbohydrate, high fat diet. For this reason, many pharmaceutical companies are attempting to 4

S. Mitsuhashi, N. Matsuura, M. Ubukata, H. Oikawa, H. Shima, and K. Kikuchi, Biochem. Biophys. Res. Commun. 287, 328–331 (2001).

l

FOREWORD

develop drugs that target this enzyme, which may be useful in the treatment of diabetes and obesity. Some relatively specific inhibitors have already been generated but, to my knowledge, no PTP1B inhibitor has yet entered human clinical trials. However, there are other protein phosphatases that are attractive drug targets, such as CD45, a PTPase expressed in cells of the immune system that is essential for T-cell activation. Inhibitors of CD45 could be effective immunosuppressants. However, simply inhibiting the catalytic subunit of most protein phosphatases will be toxic or oncogenic. The future of drug discovery in this area may therefore lie in the development of compounds that disrupt the interaction of a phosphatase with one of its regulatory or targeting subunits, or the interaction between a targeting subunit and another regulatory protein. For example, the form of PP1 associated with hepatic glycogen is complexed to the liver glycogen-targeting subunit GL. The binding of phosphorylase a to the extreme C-terminus of GL prevents PP1 from dephosphorylating and activating glycogen synthase.5 A drug that disrupted the interaction of phosphorylase a with GL would be expected to activate glycogen synthase and hence stimulate the conversion of blood glucose into hepatic glycogen. Cell permeant inhibitors of the protein kinase GSK3, which also stimulate the activation of hepatic glycogen synthase, have recently been shown by Gerald Schulman to normalize blood glucose levels in animal models of diabetes. Pharmaceutical companies will tell you that it is extremely difficult (and therefore unattractive) to develop a drug that disrupts a protein–protein interaction. However, it was only ten years ago that it was considered virtually impossible to develop a really specific protein kinase inhibitor, and yet these enzymes have now become the second most studied family of drug targets after G protein-coupled receptors. It only takes one success to overcome a dogma and it may well be that, in the not too distant future, drugs that target components of protein phosphatases will become a reality. PHILIP COHEN MRC Protein Phosphorylation Unit University of Dundee Scotland

5

C. G. Armstrong, M. J. Doherty, and P. T. W. Cohen, Biochem. J. 336, 699–704 (1998).

Section I Determination, Detection, and Localization

[1]

DEPHOSPHORYLATION OF PHOSPHOTHREONINE PEPTIDES

3

[1] Monitoring of PP2A and PP2C by Phosphothreonyl Peptide Substrates By ARIANNA DONELLA-DEANA, MARCO BOSCHETTI, and LORENZO A. PINNA Reversible phosphorylation of Ser, Thr, and Tyr residues occurring through the concerted action of protein kinases and protein phosphatases affects about one-third of eukaryotic proteins and represents the most frequent and general mechanism by which nearly all biological functions are regulated. The human genome encodes more than 600 protein kinases and slightly less than 100 protein phosphatases. This figure by itself, in conjunction with the observation that protein phosphatases operate on a minority of potential phosphoacceptor residues preselected by protein kinases, would indicate that the specificity of phosphatases needs to be less stringent than that of protein kinases. This is especially true of site specificity, i.e., the ability to recognize consensus sequences defined by local structural features surrounding the phosphoacceptor aminoacid. This is a prominent feature of most protein kinases, with special reference of Ser/Thr specific ones, and has been exploited for the design of hundreds of specific peptide substrates. These proved useful for the selective monitoring of individual kinases or classes of kinases, whose structural features responsible for site specificity in many instances have been disclosed by mutational and structural analyzes (for a compilation of consensus sequences and peptide substrates of kinases see Pinna and Ruzzene1 and Ruzzene and Pinna2). In contrast many attempts to define consensus sequences recognized by individual protein phosphatases were less rewarding and consequently the development of phosphopeptides useful for the specific assay of different categories of protein phosphatases proved rather unsuccessful3 consistent with the notion that residues surrounding the phosphoaminoacid may play a marginal role in substrate recognition, which is mainly mediated by targeting subunits and/or structural modules outside the catalytic core of the phosphatase.4–7 In a way this was expectable if it is 1

L. A. Pinna and M. Ruzzene, Biochim. Biophys. Acta 131, 191 (1996). M. Ruzzene and L. A. Pinna, in ‘‘Protein Phosphorylation: A Practical Approach’’ (G. Hardy, ed.), 2nd Ed., p. 221. Oxford University Press, Oxford, UK, 1999. 3 L. A. Pinna and A. Donella-Deana, Biochim. Biophys. Acta 1222, 415 (1994). 4 M. J. Hubbard and P. Cohen, Trends Biochem. Sci. 18, 172 (1993). 5 F. Shibasaki, E. R. Price, D. Milan, and F. McKeon, Nature 382, 370 (1996). 6 H. Song, N. Hanlon, N. R. Brown, N. E. M. Noble, L. N. Johnson, and D. Barford, Mol. Cell 7, 615 (2001). 7 N. K. Tonks and B. G. Neel, Curr. Opin. Cell Biol. 13, 182 (2001). 2

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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considered that an individual phosphatase is committed to the dephosphorylation of a variety of proteins previously phosphorylated by kinases that recognize different consensus sequences. Calcineurin, e.g., is implicated in the dephosphorylation of several transcription factors, whose phosphoacceptor sites are very variable for being phosphorylated by different kinases. Likewise it would be hard to figure out any common denominator around the phosphorylated residues which are affected by PP2A.8 A telling example is also provided by two physiological targets of PP2C, hydroxymethylglutaryl Coenzyme A (HMG-CoA) reductase9 and BAD (Klumpp and Krieglstein, personal communication) whose dephosphorylated residues display sequences (HNRpSKIN and PAGpTEED, respectively) having nothing in common. The circumstance that phosphopeptides are scarcely selective substrates does not hamper their usage for handy and sensitive assays of protein phosphatase activities whenever specificity is not a priority requirement. This concept especially applies to Tyr specific protein phosphatases (PTPs), whose activity is in fact often monitored and scrutinized with the aid of phosphopeptides displaying kinetic parameters comparable to those of protein substrates.3 A sample of outstanding phosphopeptide substrates of PTPs is provided in Table I with their Kcat/Km ratio values and pertinent references. Clearly the successful usage of phosphopeptides as tools for monitoring and characterizing PTPs is due to a concomitance of properties which are not shared by Ser/Thr specific protein phosphatases (PPs). First, PTPs are extremely proficient enzymes with Kcat/Km ratios approaching in some cases the efficiency limited by diffusion events, a circumstance that makes easily applicable nonradioactive methods for monitoring their activity. Second, the dephosphorylation of tyrosyl residues can be monitored by spectrophotometric methods (taking advantage of the different absorption spectra of phosphorylated vs unphosphorylated tyrosine) and specific immunochemical assays (based on fluorescent antiphosphotyrosine antibodies) which are not applicable to P-Ser and P-Thr residues. Third, the promiscuity of PTPs is such that they are able to dephosphorylate even free phosphotyrosine10 and related compounds if these are added at appropriate concentration; thus a broadly employed PTPs substrate is p-nitrophenyl phosphate (pNPP) whose dephosphorylation correlates with a change in color which can be followed spectrophotometrically. 8

T. A. Millward, S. Zolnierowicz, and B. A. Hemmings, Trends Biochem. Sci. 24, 186 (1999). Y. P. Ching, T. Kobayashi, S. Tamura, and D. G. Hardie, FEBS Lett. 411, 265 (1997). 10 Z. Zhao, N. F. Zander, D. A. Malencik, S. R. Anderson, and E. H. Fischer, Anal. Biochem. 202, 361 (1992). 9

[1]

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DEPHOSPHORYLATION OF PHOSPHOTHREONINE PEPTIDES

TABLE I PHOSPHOPEPTIDES USED FOR SENSITIVE ASSAYS OF PROTEIN TYROSINE PHOSPHATASES Phosphopeptide AFLEDFFTSTEPQpYQPGENL NIDGDEVNpYEE DADEpYLIPQQG TAEPDpYGALYE Ac-ELEFpYMDE-NH2 RRApTVAf

PPs

Kcat/Km (S
1  M
1)  107

Ref.

TC-PTP TC-PTP PTPU323 Yersinia PTP HPTPb PTP1B PP2A

3.70 4.42 2.88 2.22 5.71 2.20 0.02

a a b b c d e

a

M. Ruzzene, A. Donella-Deana, O. Marin, J. W. Perich, P. Ruzza, G. Borin, A. Calderan, and L. A. Pinna, Eur. J. Biochem. 211, 289 (1993). b J. Zhang, Z. Zhang, K. Brew, and E. Y. Lee, Biochemistry 35, 6276 (1996). c H. Cho, R. Krishnaraj, M. Itoh, E. Kitas, W. Bannwarth, H. Saito, and C. T. Walsh, Protein Sci. 2, 977 (1993). d S. W. Vetter, Y. F. Keng, D. S. Lawrence, and Z. Y. Zhang, J. Biol. Chem. 275, 2265 (2000). e P. Agostinis J. Goris, E. Waelkens, L. A. Pinna, F. Marchiori, and W. Merlevede, J. Biol. Chem. 262, 1060 (1987). f For sake of comparison the Kcat/Km ratio of one of the best peptide substrates for Ser/Thr PPs is also reported.

By sharp contrast to PTPs, Ser/Thr PPs, whose catalytic efficiency is on the average much lower than that of PTPs (see Table I), are generally assayed by means of phosphoradiolabeled protein substrates. Commonly used are phosphorylase-a and histones (for the assay of PP1 and PP2A), RII (regulatory subunit of cAMP-dependent protein kinase) and dopamine-andcAMP-regulated phosphoprotein (for assaying calcineurin/PP2B) and casein phosphorylated by PKA (cAMP-dependent protein kinase) (for the assay of PP2C). Some PPs, notably calcineurin (which also dephosphorylates phosphotyrosyl residues) and, under special circumstances, PP2A, display significant activity toward pNPP. Unless the phosphatase is highly purified, however, pNPP cannot be exploited for the reliable monitoring of its activity owing to the much higher pNPP-ase activity of PTPs and nonspecific phosphatases which might contaminate the sample. It should be noted on the other hand that PP1 is nearly inactive on phosphopeptides,3,11–13 while calcineurin needs quite large phosphopeptides 11

S. J. McNall and E. H. Fischer, J. Biol. Chem. 263, 1893 (1988). P. Agostinis, J. Goris, L. A. Pinna, F. Marchiori, J. W. Perich, H. E. Meyer, and W. Merlevede, Eur. J. Biochem. 189, 235 (1990). 13 P. Agostinis, J. Goris, E. Waelkens, L. A. Pinna, F. Marchiori, and W. Merlevede, J. Biol. Chem. 262, 1060 (1987). 12

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[1]

(>20 residues long) in order to display an activity comparable to that observed with its protein substrates.14 At variance with PP1 and calcineurin, PP2A and PP2C do dephosphorylate a variety of short phosphopeptides with efficiencies comparable to protein substrates routinely used for monitoring their activity.12,15 Some of these peptides were subjected to structural modifications with the aim to disclose local structural features eventually acting as positive or negative determinants, irrespective of their actual physiological relevance. An unexpected outcome of these studies was that in general the replacement of phosphothreonine for phosphoserine dramatically improved dephosphorylation efficiency by PP2A, PP2C, and PP1. In particular the phosphorylated form of the peptide RRATVA, i.e., the threonyl derivative of the ‘‘kemptide’’ (RRASVA), reproducing the phosphoacceptor site of pyruvate kinase, a substrate of PP2A,16 turned out to be dephosphorylated much more readily than its phosphoseryl counterpart and also faster than the protein substrates commonly employed for monitoring PP2A and PPC, phosphorylase-a and casein phosphoradiolabeled by PKA, respectively (see Table II). A quite high P-Thr vs PSer dephosphorylation rate was also observed with PP1; in this case however the dephosphorylation rate of the peptides is negligible as compared to that of phosphorylase-a. Calcineurin does not display any marked preference for the phosphothreonyl kemptide whose dephosphorylation rate is similar to that of its phosphoseryl derivative, both being much slower than that of the protein substrate RII. By sharp contrast, as also reported in Table II, a number of nonspecific phosphatases, either acidic or alkaline, invariably display a remarkable preference for the phosphoseryl kemptide. Collectively taken these data show that the phosphothreonyl kemptide, RRApTVA, represents a first choice substrate for the assay of PP2A and PP2C and, in combination with its pS counterpart, it also provides a valuable tool for discriminating between phosphatase activities accounted for either by bona fide protein phosphatases of the PP2A and PP2C classes or by nonspecific alkaline/acidic phosphatases. The general applicability of these criteria was validated by showing that the marked preference for pT vs pS peptides is not restricted to the catalytic subunit of PP2A but is also shared by all its oligomeric forms.12,13 Likewise the pT-kemptide is by far preferred over its pS congener by both the alpha and beta isoforms of animal PP2C as well as by PP2C from the protozoan Paramecium 14

A. Donella-Deana, M. H. Krinks, C. Klee, M. Ruzzene, and L. A. Pinna, Eur. J. Biochem. 219, 109 (1994). 15 A. Donella-Deana, C. H. Mac Gowan, P. Cohen, F. Marchiori, H. E. Meyer, and L. A. Pinna, Biochim. Biophys. Acta 1051, 199 (1990). 16 M. Nishimura and K. Uyeda, J. Biol. Chem. 270, 26341 (1995).

[1]

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DEPHOSPHORYLATION OF PHOSPHOTHREONINE PEPTIDES

TABLE II PREFERENTIAL DEPHOSPHORYLATION OF PHOSPHOTHREONYL VS PHOSPHOSERYL PEPTIDE BY SOME CLASSES OF PPs Relative activitya

Dephosphorylation ratio

RRApTVA

RRApSVA

(pT/pS)

Protein phosphatases PP1 PP2A PP2B (calcineurin) PP2C

2.0 712.1 1.6 253.2

0.1 11.0 0.5 8.8

20.02 64.69 3.20 28.71

Alkaline phosphatases S. cerevisiae Intestinal E. coli

0.0 40.2 0.0

72.4 326.3 563.9

0.00 0.12 0.00

Acid phosphatases S. cerevisiae Prostatic Potato Wheat germ

2.4 32.4 0.4 1.4

51.4 205.8 5.1 8.6

0.04 0.15 0.08 0.16

a Expressed as percent of the dephosphorylation rate of the protein substrates phosphorylase a, RII and phosphocasein in the case of protein phosphatases PP1/PP2A, PP2B and PP2C, respectively, and of pNPP in the case of alkaline and acid phosphatases. Drawn from A. Donella-Deana, M. H. Krinks, C. Klee, M. Ruzzene, and L. A. Pinna, Eur. J. Biochem. 219, 109 (1994), and A. Donella-Deana, H. E. Meyer, and L. A. Pinna, Biochim. Biophys. Acta 1094, 130 (1991), respectively.

tetraurelia17 and by SpoIIE, a bacterial protein phosphatase related to the PP2C class of eukaryotic protein phosphatases.18 Interestingly, moreover, preferential dephosphorylation of P-Thr residues is not restricted to peptide substrates, since PP2C specifically dephosphorylated only pT residues present in a sample of 32P-casein evenly radiolabeled at Ser and Thr residues,15 and a mutant of HMG-CoA reductase with the phosphorylatable serine replaced by threonine, once phosphorylated by AMP-activated protein Kinase (AMPK) was dephosphorylated by PP2A much more readily than the phospho-wild type protein.9 This does not necessarily mean that preferential P-Thr dephosphorylation reflects a physiological situation. As a matter of fact P-Ser is by far predominant over P-Thr in naturally 17

S. Klumpp, C. Hanke, A. Donella-Deana, A. Beyer, R. Kellner, L. A. Pinna, and J. E. Schultz, J. Biol. Chem. 269, 32774 (1994). 18 E. Adler, A. Donella-Deana, F. Arigoni, L. A. Pinna, and P. Stragier, Mol. Microbiol. 23, 57 (1997).

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[1]

occurring phosphoproteins19 and, to the best of our knowledge there is no in vivo data supporting the view that the latter are turning over more rapidly than the former. However, it may be pertinent to note that phosphothreonine is frequently found at sites which have been reported to be affected by PP2C, notably the regulatory phosphothreonines found in the glycine rich loop of cyclin-dependent kinases (CDKs)20 and at the activation loop of MAP kinases21,22 and of AMPK23 and the aforementioned T-117 of the BAD protein. This may suggest that PP2C has been adopted as a specialized ‘‘phosphothreonyl phosphatase.’’ On the other hand, the preference of PP2C for P-Thr vs P-Ser residues is also deeply influenced by the surrounding aminoacids. In fact the replacement of pT for pS in the context of the HMG-CoA reductase phosphosite (HNRpSKINL) which accelerates >10 fold the rate of dephosphorylation by PP2A, has only a modest effect on dephosphorylation by PP2C.9 Likewise the phosphopeptide RRREEpTEEE, which is an excellent substrate of PP2A by virtue of its phosphothreonyl residue, is almost unaffected by PP2C.15 This provides a tool for discriminating PP2A from PP2C activities using peptide substrates. The structural features underlying the striking preference of PP2A and PP2C for phosphothreonyl over phosphoseryl residues are still unknown, since structural data of complexes between Ser/Thr protein phosphatases and peptide substrates are not available. Kinetic data show that with PP2A and, to a lesser extent, with PP2C the beneficial effect of phosphothreonine is mostly accounted for by increased Vmax, whereas the Km values of the pT and pS peptides are similar (see Table III), suggesting that the methyl group of threonine accelerates the catalytic event rather than increasing the binding affinity. Paradoxically the opposite might be expectable assuming a catalytic mechanism based on protonation of the leaving hydroxyaminoacid side chain, since the methyl group is a better electron donor than an hydrogen atom. This would make the P–O ester bond of phosphothreonine less polarized toward the oxygen which would therefore be less susceptible to protonation as compared to the oxygen of phosphoserine. Consequently phosphothreonyl residues, on the average, are more acid stable than phosphoserine ones.24 The reason why 19

T. Mustelin, ‘‘Src Family Tyrosine Kinases in Leukocytes.’’ MBIU, R.G. Landes Co, Austin, TX, 1994. 20 A. Cheng, P. Kaldis, and M. J. Solomon, J. Biol. Chem. 275, 34744 (2000). 21 C. C. Fjeld and J. M. Denu, J. Biol. Chem. 274, 20336 (1999). 22 J. Warmka, J. Hanneman, J. I. Lee, D. Amin, and I. Ota, Mol. Cell Biol. 21, 51 (2001). 23 A. E. Marley, J. E. Sullivan, D. Carling, W. M. Abbott, G. J. Smith, I. W. Taylor, F. Carey, and R. K. Beri, Biochem. J. 320, 801 (1996). 24 D. B. Bylund and T. S. Huang, Anal. Biochem. 73, 477 (1976).

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DEPHOSPHORYLATION OF PHOSPHOTHREONINE PEPTIDES

TABLE III KINETIC CONSTANTS FOR THE ENZYMATIC DEPHOSPHORYLATION OF pT AND pS KEMPTIDES BY PP2A AND PP2C PP2Aa

RRApTVA RRApSVA

PP2Cb

Vmaxc

Kmd

Vmaxc

Kmd

1250 131

16 16

2400 416

1.0 2.5

a

P. Agostinis, J. Goris, L. A. Pinna, F. Marchiori, J. W. Perich, H. E. Meyer, and W. Merlevede, Eur. J. Biochem. 189, 235 (1990). b S. Klumpp, C. Hanke, A. Donella-Deana, A. Beyer, R. Kellner, L. A. Pinna, and J. E. Schultz, J. Biol. Chem. 269, 32774 (1994). c Expressed as nmol/min/mg. d Expressed as M.

they conversely are much proner to cleavage by some classes of protein phosphatases, notably 2A and 2C, remains unexplained. One possibility would be that the methyl group becomes a steric hindrance once the phosphoester bond has been cleaved, thus accelerating the release of the dephosphorylated product, assuming this is the rate limiting step in the overall catalytic reaction. This may apply to PP2A, whose mechanism of catalysis is unknown, while in the case of PP2C the rate limiting step seems to be the release of the phosphate.21 To sum up, while the mechanistic features underlying preferential dephosphorylation of phosphothreonyl residues by PP2A and PP2C, and, even more the possible physiological significance of such a selection are still enigmatic, the practical usefulness of phosphothreonyl peptides for assaying their activity is self evident and it may deserve more attention than it has been given in the past.

Experimental Procedures Assay with Radioactive Phosphopeptides

1a Preparation of Radiolabeled Phosphopeptides Solutions Solution A: A buffer, suitable for the specific kinase, containing 10 mM unlabeled ATP and [ 32P]ATP or [ 33P]ATP (Amersham Pharmacia Biotech) to reach the specific activity of 2000 cpm/pmol, 10 mM MgCl2 or MnCl2, 1 mM peptide, the kinase (in the case of

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DETERMINATION, DETECTION, AND LOCALIZATION

[1]

kemptides the catalytic subunit of PKA) and the appropriate cofactors/ activators. Solution B: 30% (v : v) acetic acid. Procedure 1.

2.

3.

4.

5.

6.

7. 8.

The standard peptide phosphorylation is carried out in 200 l (or more depending on the amount of phosphopeptide required) of solution A. The reaction is stopped by addition of 85 l of 100% (v : v) acetic acid plus 215 l of 30% (v : v) acetic acid to reach a 0.5 ml volume and the acetic acid concentration of 30% (v : v). A number of Pasteur pipettes, corresponding to the number of phosphorylated peptides are prepared. Each pipette is plugged with glass wool and held by a suitable rack. AG 1-X8 resin (0.5 ml) is poured onto each pipette and the resin is extensively washed with 30% (v : v) acetic acid. Each sample is loaded on a pipette and the flow through fraction is collected. The resin is washed three times with 1 ml of 30% (v : v) acetic acid and the eluate is collected in the same test tube as the flow through to a total volume of 3.5 ml. 20 l of each sample is counted by scintillation counter to estimate the degree of peptide phosphorylation and to calculate the concentration of the phosphopeptide. The pipette eluate, containing a mixture of the unphosphorylated and the phosphorylated peptide, is supplemented with an equal volume of water to allow its freezing at
80 C, lyophilized, resuspended with water and lyophilized three times to remove traces of acetic acid. The lyophilized phosphopeptide is redissolved at the desired concentration in water, or in buffer at the appropriate pH. 2 l of the resuspended phosphopeptide are counted in a scintillation counter and the concentration (pmol) of the phosphopeptide is calculated by dividing the cpm of the radioactive peptide by the specific radioactivity of the [ 32P]ATP or [ 33P]ATP (cpm/pmol) used for the phosphorylation reaction.

Generally it is not necessary to separate the phosphorylated peptide from its nonphosphorylated counterpart since it has been shown that even large excess of the latter does not appreciably affect the kinetic parameters of protein phosphatases, either Ser/Thr or Tyr specific. If the mixture of phosphorylated and nonphosphorylated peptides is going to be used in the subsequent phosphatase assay it is especially important to know exactly the

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11

specific radioactivity of 32P-ATP or 33P-ATP in the phosphorylation reaction since the concentration of the phosphopeptide will be assumed to be equal to that of the radiolabeled phosphate incorporated into the peptide. 1b Separation of the Phosphopeptide from its Nonphosphorylated Form If for any reason one wants to use the purified phosphopeptide, free of its nonphosphorylated form, in the phosphatase assay, separation can be readily achieved by RP-HPLC on C18 column where the phosphoforms in general tend to be eluted before their nonphosphorylated derivatives. In particular, separation of RRApTVA from RRATVA, exemplified in Fig. 1, can be attained as follows: Solutions Solution A: 0.1% aqueous trifluoroacetic acid (TFA) (Fluka). Solution B: acetonitrile (Fluka) containing 0.08% TFA. Procedure 1.

The dried mixture of nonphosphorylated and 33P-phosphorylated peptide obtained from procedure 1a (points 1–6) is dissolved in

FIG. 1. RP-HPLC separation of phosphorylated from nonphosphorylated peptide. Twentyfive microgram each of synthetic RRATVA and RRApTVA were mixed with partially phosphoradiolabeled RRATVA obtained as described in the experimental procedure 1a (steps 1–7) and loaded on a C18 Symmetry column eluted as described in the text (1b).

12

DETERMINATION, DETECTION, AND LOCALIZATION

2. 3.

4.

[1]

solution A and subjected to analytical Reverse-Phase-HPLC using a C18 symmetry 300 column (4.6  250 mm) (Waters), equilibrated with solution A. The column is eluted at a flow rate of 1 ml/min with a linear gradient of solution B from 0 to 30% for 60 min. The elution of both phosphorylated and nonphosphorylated peptides is monitored by absorbance at 220 nm. The radioactivity of the fractions collected every 30 sec is measured by counting an aliquot in liquid scintillation counter. As shown in Fig. 1 radioactivity overlaps the less retarded absorbance peak. The fractions containing the phosphopeptide are lyophilized.

2a Dephosphorylation Reaction and Assay of Radioactive Inorganic Phosphate Solutions Solution A: 10% (w : v) TCA (trichloroacetic acid) (Mallinckrodt Baker). Solution B: 5% (w : v) ammonium molybdate; 10 g of ammonium molybdate tetrahydrated (Mallinckrodt Baker) are added in small amounts to a solution containing 22.4 ml of concentrated H2SO4 and 60 ml of H2O. After solubilization, the solution volume is adjusted to 200 ml with H2O. Solution C: water-saturated isobutanol–toluene (1:1); 1 part of isobutanol (Mallinckrodt Baker) is stirred with 1 part of toluene (Mallinckrodt Baker) and about 0.5 part of H2O. The organic phase can be used when the aqueous and organic phases are completely separated. Solution D: scintillation liquid (Packard). Procedure 1.

2.

3. 4.

Dephosphorylation assay is performed in 30 l of incubation medium containing the buffer specific for the phosphatase of choice, the radioactive peptide (3–20 M) dissolved in water or in a convenient buffer, and the appropriate cofactors/activators. The dephosphorylation reaction is stopped by adding 1.5 ml of 10% TCA plus 0.5 ml of 5% ammonium molybdate, to convert inorganic phosphate into phosphomolybdate. 2.5 ml of isobutanol/toluene are added. The tubes are capped and shaken for at least 30 sec by vortex or by a rotational shaker. This extracts phosphomolybdate into the inorganic phase, while nonreacted phosphopeptide(s) remain in the aqueous phase.

[1]

DEPHOSPHORYLATION OF PHOSPHOTHREONINE PEPTIDES

5.

13

When the two phases are separated, 2 ml of the upper organic phase are withdrawn, added to 2 ml of scintillation liquid and counted in scintillation counter. This provides the basis for the calculation of inorganic phosphate released since its specific radioactivity is known.

2b Simplified Procedure for Quantifying Phosphate Release The above procedure is the most reliable, since it provides the absolute assurance that radioactivity measured is accounted for by inorganic phosphate alone. If, however, the protein phosphatase preparation to be assayed is sufficiently pure, conversion of inorganic phosphate into phosphomolybdate complex and the extraction of this latter with organic solvent can be omitted, and the phosphate generated through the phosphatase reaction can be directly determined after quantitative removal of excess nonreacted phosphopeptide. This can be done, in the case of the phospho-kemptides and other monophosphorylated peptides with two or more arginine (or eventually lysine) residues, using phosphocellulose paper absorption method.25 Solutions Solution A: 0.5% (v : v) phosphoric acid (J. T. Baker). Solution B: 20% (w : v) tricloacetic acid (TCA) (J. T. Baker). Procedure 1.

2.

3.

4.

The dephosphorylation reaction performed in 30 l of incubation mixture as described above is stopped with 7 l of 20% tricloacetic acid. 30 l of each sample are spotted onto P81 phosphocellulose paper (Whatman) strips (2  1 cm), which bind the phosphopeptide but not inorganic phosphate. Each paper strip is stirred for 30 sec in a tube containing 1 ml 0.5% phosphoric acid in order to extract inorganic phosphate released during the reaction. The amount of phosphate is quantified by counting 0.8 ml of phosphoric acid in a liquid scintillator.

The applicability of this method is critically dependent on the lack of proteolytic activity which otherwise would cleave the peptide and remove the basic residues required for its binding to phosphocellulose. The addition 25

D. B. Glass, R. A. Masaracchia, J. R. Feramisco, and B. E. Kemp, Anal. Biochem. 87, 566 (1978).

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DETERMINATION, DETECTION, AND LOCALIZATION

[1]

of a protease inhibitor cocktail to the reaction mixture can obviate this drawback. In the case of PP2A assay it is advisable to include a control with okadaic acid (0.1 M) or another specific inhibitor which will validate the implication of the phosphatase in the generation of the soluble radioactive pool, ruling out any artifact due to other activities. This is not possible with PP2C, for which a potent and specific inhibitor comparable to okadaic acid is not available. With the caveats and limitations just outlined, this method provides results quite comparable to those of procedure 2a with the advantage of skipping the troublesome steps of generating the phosphomolybdate complex and extracting it with hazardous organic solvents. If the phosphopeptide does not bind to P-cellulose paper for not being positively charged at low pH values, advantage could be taken of the tendency of peptides having aromatic side chains to absorb on charcoal, a property which has been exploited to remove nonreacted phosphotyrosyl peptides at the end of PTPase reaction.26 While aromatic residues are not present in the pT-kemptide, (RRApTVA), a tyrosyl or a tryptophanyl residue could be replaced for its valine without suppressing its ability to be phosphorylated by PKA and dephosphorylated by PP2A and PP2C.

Nonradioactive Assay (Malachite Green)

Methods based on colorimetric determination of inorganic phosphate released by the phosphatase reaction have the advantage of avoiding both the hazard of manipulating radioactive compounds and the aleatority of having to rely on enzymatic reactions to prepare the phosphosubstrate, whose yield, moreover, is often quite low. Nowadays synthetic procedures are available which allow to produce in principle any kind of phosphopeptide, including phosphothreonyl ones27 with yields approaching 100% and without the limitation imposed by the specificity and efficiency of available protein kinases. The main limit of these nonradioactive methods is the relatively low sensitivity of colorimetric determination of inorganic phosphate. This restricts their application to the assay of at least partially purified phosphatases and not of phosphatase activity in crude lysates where high background of inorganic phosphate is likely to be present. In general the highest acceptable background should not exceed 10% of the 26

M. Streuli, N. X. Krueger, A. Y. Tsai, and H. Saito, Proc. Natl. Acad. Sci. U.S.A. 86, 8698 (1989). 27 D. D. Williams, O. Marin, L. A. Pinna, and C. G. Proud, FEBS Lett. 448, 86 (1999).

[1]

DEPHOSPHORYLATION OF PHOSPHOTHREONINE PEPTIDES

15

phosphate released enzymatically. Note on the other hand that if initial dephosphorylation rates are to be determined phosphate released must not exceed a minor proportion ( 60 and almost 30-fold higher than with the latter, respectively (see Table II). Malachite Green Assay Solutions Solution A: concentrated sulfuric acid (60 ml) is slowly added to 300 ml of water. The solution is then cooled at room temperature and supplemented with 0.44 g of malachite green (Merck). Solution B: 15% (w/v) ammonium molybdate tetrahydrate (Merck). Solution C: 11% (w/v) Tween 20 (Merck). Solution D: malachite green solution is prepared on the day of use by stirring 10 ml of solution A with 2.5 ml of solution B and 0.2 ml of solution C. Solution E: Pi standard solution; potassium phosphate monobasic (Sigma), equivalent to 20 g P/ml is dissolved in 0.05 M HCl. Procedure 1. 2. 3.

28

50 l of solution D is mixed with 200 l of the solution to be analyzed. The color is allowed to develop for 30 min at 22 C. The absorbance of each reaction mixture is measured at 650 nm, using the appropriate blank.

T. P. Geladopoulos, T. G. Sotiroudis, and A. E. Evangelopoulos, Anal. Biochem. 192, 112 (1991). 29 K. Itaya and M. Ui, Clin. Chim. Acta 14, 361 (1966). 30 A. A. Baykov, O. A. Evtushenko, and S. M. Avaeva, Anal. Biochem. 171, 266 (1988). 31 D. F. McCain and Z. Y. Zhang, Methods Enzymol. 345, 507 (2002).

16

DETERMINATION, DETECTION, AND LOCALIZATION

4.

[1]

Quantification of the released Pi is obtained by constructing an appropriate calibration curve with Pi standard solution E.

Perspectives

While the empirical usage of phosphothreonyl peptides as substrates to assay Ser/Thr specific protein phosphatases with special reference to PP2A and PP2C, is grounded on solid arguments, progress is desirable in two directions: (i) the understanding of the structural features rendering phosphothreonyl residues much more susceptible to dephosphorylation as compared to phosphoseryl ones; (ii) technical improvements aimed at making the assay more sensitive and performable by colorimetric and/or continuous spectrophotometric techniques, avoiding radioactive reagents. Structural insights into the mode of binding of phosphothreonyl peptides to the active site of PP2A and PP2C, in conjunction with mutational analysis will disclose the rationale for the beneficial effect of the methyl group of threonine on catalytic efficiency and will pave the road toward the design of phosphopeptides and/or peptidomimetics susceptible to faster and more selective dephosphorylation as compared to the ones available to date. On the other hand methodological improvements to the assay could also come from alterations in the sequence surrounding phosphothreonine, aimed at further increasing dephosphorylation efficiency. One strategy would be that of reinforcing the beneficial effect of phosphothreonine by additional structural features which have been shown to improve dephosphorylation by independent mechanisms. Pertinent to this may be the observation that a cluster of four arginines at position spanning between n-3 and n-6 increases 20-fold the dephosphorylation of a phosphoseryl peptide related to kemptide by PP2A.12 An alternative approach would be that of designing phosphothreonyl peptide substrate unrelated to the kemptide, either drawn from the phosphosites of proteins which are readily dephosphorylated by PP2A and/or PP2C (e.g., AMPK, MAP kinases etc.) or artificially constructed by the ‘‘inverse alanine scanning’’ approach, which proved useful for designing one of the best substrates for PTPs32 (see Table I). While these improvements are expected to increase the sensitivity of methods based on colorimetric determination of inorganic phosphate, different methodological approaches, commonly exploited 32

S. W. Vetter, Y. F. Keng, D. S. Lawrence, and Z. Y. Zhang, J. Biol. Chem. 275, 2265 (2000).

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17

for the assay of PTPs, could become applicable also to PP2A and PP2C if the phosphopeptide substrate displays fluorescence properties affected by the phosphate. This could be done by placing a tryptophan adjacent to the phosphorylated threonine. In particular, if the Val at n þ 1 in the kemptide is replaced by a tryptophan, the fluorescence intensity of this latter is expected to change in response to dephosphorylation of the adjacent residue, by analogy with the behavior of phosphoseryl peptides.33 Even a derivatization of the phosphothreonyl residue itself could be considered, e.g., by replacing the hydrogen of the  carbon with aromatic groups whose fluorescence spectra are modified upon dephosphorylation. This would make applicable also in the case of Ser/Thr PPs continuous nonradioactive assays which, owing to their rapidity and precision, are often exploited, instead of the traditional quenched assays, for monitoring PTPs.31

Acknowledgments The skilful technical assistance of Mr. G. Tasinato is gratefully acknowledged. This work was supported by the Italian Association for Cancer Research (AIRC), the Ministero dell’Istruzione dell’Universita’ della Ricerca (MIUR) (COFIN 2000 and 2001) and the Centro Nazionale della Ricerca (CNR) (no. 98.03280.ST74 and Target Project on Biotechnology).

33

D. J. Wright, E. S. Noiman, P. B. Chock, and V. Chau, Proc. Natl. Acad. Sci. USA 78, 6048 (1981).

[1] Determination of Mammalian Glycogen Synthase Phosphatase Activity By ANNA A. DEPAOLI-ROACH, PIER GIUSEPPE VILARDO, JONG-HWA KIM, NIRMALA MAVILA, BHARGAVI VEMURI, and PETER J. ROACH

Glycogen serves as a repository of glucose and is a major source of energy in many cell types. In mammals, the major stores are in skeletal muscle and liver, but other tissues such as cardiac, adipose, kidney and brain are capable of synthesizing glycogen. Because of its large mass in the whole body, skeletal muscle plays an important role in blood glucose homeostasis and in fact,  80% of the glucose taken up postprandially is converted to glycogen in

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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for the assay of PTPs, could become applicable also to PP2A and PP2C if the phosphopeptide substrate displays fluorescence properties affected by the phosphate. This could be done by placing a tryptophan adjacent to the phosphorylated threonine. In particular, if the Val at n þ 1 in the kemptide is replaced by a tryptophan, the fluorescence intensity of this latter is expected to change in response to dephosphorylation of the adjacent residue, by analogy with the behavior of phosphoseryl peptides.33 Even a derivatization of the phosphothreonyl residue itself could be considered, e.g., by replacing the hydrogen of the  carbon with aromatic groups whose fluorescence spectra are modified upon dephosphorylation. This would make applicable also in the case of Ser/Thr PPs continuous nonradioactive assays which, owing to their rapidity and precision, are often exploited, instead of the traditional quenched assays, for monitoring PTPs.31

Acknowledgments The skilful technical assistance of Mr. G. Tasinato is gratefully acknowledged. This work was supported by the Italian Association for Cancer Research (AIRC), the Ministero dell’Istruzione dell’Universita’ della Ricerca (MIUR) (COFIN 2000 and 2001) and the Centro Nazionale della Ricerca (CNR) (no. 98.03280.ST74 and Target Project on Biotechnology).

33

D. J. Wright, E. S. Noiman, P. B. Chock, and V. Chau, Proc. Natl. Acad. Sci. USA 78, 6048 (1981).

[2] Determination of Mammalian Glycogen Synthase Phosphatase Activity By ANNA A. DEPAOLI-ROACH, PIER GIUSEPPE VILARDO, JONG-HWA KIM, NIRMALA MAVILA, BHARGAVI VEMURI, and PETER J. ROACH

Glycogen serves as a repository of glucose and is a major source of energy in many cell types. In mammals, the major stores are in skeletal muscle and liver, but other tissues such as cardiac, adipose, kidney and brain are capable of synthesizing glycogen. Because of its large mass in the whole body, skeletal muscle plays an important role in blood glucose homeostasis and in fact,
80% of the glucose taken up postprandially is converted to glycogen in

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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DETERMINATION, DETECTION, AND LOCALIZATION

[2]

this tissue. Glycogen synthesis involves various steps,1,2 starting with transport of glucose inside the cell, where it is rapidly phosphorylated by hexokinase to glucose-6-P (G-6-P). Through the action of phosphoglucomutase and UDP-glucose pyrophosphorylase, the hexose phosphate is converted to UDP-glucose, the glycosyl donor for glycogen synthesis by glycogen synthase (GS). The branching enzyme introduces branchpoints as -1,6 glycosidic linkages. Glycogen degradation is catalyzed by glycogen phosphorylase (Ph) and debranching enzyme, yielding primarily glucose-1-P. In skeletal muscle, glycogen metabolism is regulated by hormones, such as insulin and epinephrine3,4 as well as by neuronal stimuli during muscle contraction.5 Defects in glycogen metabolism are associated with diabetes.1 Within the cell, glycogen metabolism is controlled largely by the coordinated regulation of the two enzymes responsible for its synthesis and breakdown, GS and Ph, respectively. Both enzymes are controlled by covalent phosphorylation and by allosteric effectors.2,4 In vitro, muscle GS undergoes a complex multisite phosphorylation at nine serine residues by ten or more protein kinases2,6–8 (Fig. 1) including, cAMP-dependent protein kinase (PKA), casein kinase I (CKI), casein kinase II (CKII), glycogen synthase kinase-3 (GSK-3), phosphorylase kinase (PhK), protein kinase C (PKC), AMP-activated protein kinase (AMPK) and calmodulin-dependent protein kinaseII (CaMK). CKI and GSK-3 act in a hierarchical manner9 in the sense that they require prior phosphorylation by other protein kinases to generate their recognition sites. CKI requires phosphorylation at Ser7 (site 2) in order to act on Ser10 (site 2a). GSK-3 recognizes phosphate groups 4 residues COOH-terminal to the target site. CKII forms the recognition site (site 5) for GSK-3 which then progressively phosphorylates 1

A. V. Skurat and P. J. Roach, in ‘‘Diabetes Mellitus: A Fundamental and Clinical Text’’ (D. LeRoith, J. E. Olefsky, and S. Taylor, eds.), 2nd Ed., p. 251. J.B. Lippincott Company, Philadelphia, USA, 2000. 2 P. J. Roach, Curr. Mol. Med. 2, 101 (2002). 3 J. C. Lawrence, Jr., J. F. Hiken, A. A. DePaoli-Roach, and P. J. Roach, J. Biol. Chem. 258, 10710 (1983). 4 P. Cohen, in ‘‘The Enzymes’’ (P. Boyer and E. G. Krebs, eds.), 3rd Ed., p. 461. Academic Press, Orlando, 1986. 5 W. G. Aschenbach, Y. Suzuki, K. Breeden, C. Prats, M. F. Hirshman, S. D. Dufresne, K. Sakamoto, P. G. Vilardo, M. Steele, J. H. Kim, S. L. Jing, L. J. Goodyear, and A. A. DePaoli-Roach, J. Biol. Chem. 276, 39959 (2001). 6 C. Picton, A. Aitken, T. Bilham, and P. Cohen, Eur. J. Biochem. 124, 37 (1982). 7 L. Poulter, S. G. Ang, B. W. Gibson, D. H. Williams, C. F. Holmes, F. B. Caudwell, J. Pitcher, and P. Cohen, Eur. J. Biochem. 175, 497 (1988). 8 A. A. DePaoli-Roach, Z. Ahmad, M. Camici, J. C. Lawrence, Jr., and P. J. Roach, J. Biol. Chem. 258, 10702 (1983). 9 P. J. Roach, J. Biol. Chem. 266, 14139 (1991).

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GLYCOGEN SYNTHASE PHOSPHATASE ACTIVITY

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FIG. 1. Diagram of glycogen synthase. The rabbit skeletal muscle GS (743 amino acids and Mr 85,000) is shown with its NH2 and COOH-termini phosphorylation sites. The kinases that phosphorylate the various sites in vitro are also listed. The asterisks denote the sites that affect enzyme activity. PKA, cAMP-dependent protein kinase; AMPK, AMP-activated kinase; PhK, phosphorylase kinase; PKC, protein kinase C; PKG, protein kinase G; S6KII, S6 kinase II; CaMK, calmodulin-dependent protein kinase; MAPKAPK-2, mitogen-activated protein kinase activated protein kinase-2; CK-I, casein kinase I; CK-II, casein kinase-II; GSK-3, glycogen synthase kinase-3.

sites 4, 3a, 3b, and 3c. The most important regulatory phosphorylation sites are distributed between the NH2- (sites 2 and 2a) and the COOHtermini (sites 3a and 3b) of the GS molecule (Fig. 1; Refs. 10–12). In general, phosphorylation results in inactivation which is antagonized by binding of the allosteric effector G-6-P. Therefore, the ratio of the GS activity in the absence and in the presence of G-6-P provides an index of the phosphorylation state of the enzyme. Ph is activated by phosphorylation of a single site, Ser14, by PhK to form Ph a.13 The less active, dephosphorylated form (Ph b) acquires full activity in the presence of the allosteric effector AMP. The / þ AMP activity ratio is used as a kinetic index of the phosphorylation state of Ph. Mutagenesis studies have revealed the regulatory phosphorylation sites on GS10–12 important for modulation of activity. Sites 1a and 1b are not essential for inactivation, whereas sites 2 and 3a are the most effective in controlling enzyme activity. These results are consistent with previous studies in which epinephrine-induced inactivation of GS correlated with

10

A. V. Skurat, Y. Wang, and P. J. Roach, J. Biol. Chem. 269, 25534 (1994). A. V. Skurat and P. J. Roach, J. Biol. Chem. 270, 12491 (1995). 12 A. V. Skurat, A. D. Dietrich, and P. J. Roach, Diabetes 49, 1096 (2000). 13 P. Cohen, Biochem. Soc. Trans. 15, 999 (1987). 11

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DETERMINATION, DETECTION, AND LOCALIZATION

[2]

increased phosphorylation at the site 2 and site 3 regions.3,14,15 Similarly, activation of GS by insulin was associated with decreased phosphorylation in the same regions.3,15,16 Therefore, neither the direct action of PKA, activated by b-adrenergic agonists, nor that of GSK-3, inhibited by insulin,17,18 can account for the modulation of GS activity by the hormones that alter phosphorylation at sites not substrates for the cognate kinases. These observations led to the postulation that both hormones may elicit their effect on GS by regulating protein phosphatases that are able to dephosphorylate sites at both the NH2- and COOH-termini.19 Although the same phosphatases may also act on Ph, their action is not strictly required for hormonal control of this enzyme. Phosphorylase activation by epinephrine can proceed through phosphorylation and activation of PhK by PKA, which in turn phosphorylates and activates Ph. Insulin has little effect on Ph activity. The glycogen-associated phosphatases, PP1Gs, have been implicated in the hormonal control of glycogen synthase.19 In rodent skeletal muscle, three forms have been identified and in human muscle a liver-specific form, GL, may also be present.20–22 They all consist of a catalytic subunit of protein phosphatase 1 (PP1c) and a regulatory/targeting subunit which localizes the enzyme to glycogen in proximity of the glycogen metabolizing enzymes.20 The RGL/GM subunit is specifically expressed in striated muscle23 whereas the protein targeting to glycogen (PTG) and R6 are more ubiquitous. Genome-wide searches have identified three additional potential glycogen-binding subunits24 whose significance is completely unknown. Both PP1G/RGL and PTG have been implicated in insulin control of glycogen metabolism.25 While a mechanism for PTG has not been 14

P. J. Parker, N. Embi, F. B. Caudwell, and P. Cohen, Eur. J. Biochem. 124, 47 (1982). V. S. Sheorain, H. Juhl, M. Bass, and T. R. Soderling, J. Biol. Chem. 259, 7024 (1984). 16 P. J. Parker, F. B. Caudwell, and P. Cohen, Eur. J. Biochem. 130, 227 (1983). 17 D. A. Cross, P. W. Watt, M. Shaw, J. van der Kaay, C. P. Downes, J. C. Holder, and P. Cohen, FEBS Lett. 406, 211 (1997). 18 J. C. Lawrence, Jr., and P. J. Roach, Diabetes 46, 541 (1997). 19 M. J. Hubbard and P. Cohen, Trends Biochem. Sci. 18, 172 (1993). 20 A. A. DePaoli-Roach, in ‘‘Handbook of Cellular Signaling’’ (R. A. Bradshaw and E. D. Dennis, eds.). Academic Press, San Diego, 2003, in press. 21 P. T. Cohen, J. Cell. Sci. 115, 241 (2002). 22 S. Munro, D. J. Cuthbertson, J. Cunningham, M. Sales, and P. T. Cohen, Diabetes 51, 591 (2002). 23 Y. Suzuki, C. Lanner, J.-H. Kim, P. G. Vilardo, H. Zhang, J. Jie Yang, L. D. Cooper, M. Steele, A. Kennedy, C. Bock, A. Scrimgeour, J. C. Lawrence, Jr., and A. A. DePaoli-Roach, Mol. Cell. Biol. 21, 2683 (2001). 24 H. Ceulemans, W. Stalmans, and M. Bollen, Bioessays 24, 371 (2002). 25 M. J. Brady and A. R. Saltiel, Recent Prog. Horm. Res. 56, 157 (2001). 15

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21

elucidated, PP1G/RGL would be activated by phosphorylation of Ser48 by an insulin-stimulated protein kinase.19,26 The activated phosphatase would then dephosphorylate and activate glycogen synthase. However, studies in our laboratory utilizing RGL knockout mice have shown that this enzyme is not required for insulin activation of muscle GS.23 In both wild type and RGL null mutant mice, an insulin-stimulated glycogen synthase phosphatase activity was still present.23 RGL has also been postulated to play a role in epinephrine control of glycogen metabolism.19 Phosphorylation at Ser67 by PKA may cause dissociation of PP1c from RGL with the released PP1c being less active towards GS and Ph. Furthermore, activation of PKA would lead to phosphorylation of phosphatase inhibitor-1 (I-1), which then becomes a potent inhibitor of the free PP1c. By this mechanism, the phosphatase activity towards the glycogen metabolizing enzymes would be reduced, resulting in inhibition of glycogen synthesis and stimulation of glycogen breakdown. Studies with RGL and I-1 knockout mice, though, indicate that neither is essential for epinephrine-induced inactivation of GS and activation of Ph.27,28 However, the possibility that other protein phosphatases mediate epinephrine control of glycogen metabolism, as is most likely for insulin action, cannot be ruled out. It is clear that protein phosphatases are equal partners with protein kinases in controls by reversible phosphorylation even though understanding of their regulation is not as advanced as that of protein kinases. Technically, studies of protein phosphatases are more complex because phosphorylated substrates are required. This means that not only must protein substrates be available but also the kinases that phosphorylate them. Another complication is that, if the phosphatase is itself regulated by phosphorylation, it is difficult to prevent its dephosphorylation during preparation of a cell or tissue extract and still retain enzyme activity. Compounds that block dephosphorylation may also inhibit the activity of the phosphatase of interest. Nevertheless, it is critical to determine phosphatase activity towards specific physiological substrates. Most importantly, it is especially desirable to be able to detect the effect of the phosphatase at sites on substrates that modulate activity or function. This is particularly true for GS in which a large number of sites are phosphorylated but not all contribute equally to changes in activity. 26

P. Dent, A. Lavoinne, S. Nakielny, F. B. Caudwell, P. Watt, and P. Cohen, Nature 348, 302 (1990). 27 A. G. Scrimgeour, P. B. Allen, A. A. Fienberg, P. Greengard, and J. C. Lawrence, Jr., J. Biol. Chem. 274, 20949 (1999). 28 A. A. DePaoli-Roach, Y. Suzuki, C. Lanner, J.-H. Kim, W. G. Aschenbach, C. Prats, P. G. Vilardo, M. Steele, M. F. Hirshman, and L. J. Goodyear, Diabetes 50, Supp. 2, 1136 (2001).

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[2]

Principles

The most commonly used substrate for the determination of Ser/Thr protein phosphatase activity is Ph. The protein is abundantly expressed in skeletal muscle and gram quantities can be easily prepared from rabbits. In addition, it is phosphorylated at a single site by Phk, which is also expressed at high level and is relatively easy to isolate. Phosphatase activity can be monitored either by 32Pi release from 32P-Ph a or changes in activity29 after dephosphorylation. A similar approach can also be used to monitor GS phosphatase activity, although GS is not as abundant as Ph, is phosphorylated at multiple sites by different protein kinases and the phosphorylation sites do not contribute equally to changes in activity.

Advantages and Disadvantages

Insulin, epinephrine and muscle contraction regulate GS activity at least in part through control of protein phosphatase(s).19,25 Utilizing 32P-Ph a as a substrate, we were unable to detect insulin-stimulated phosphatase activity in mouse skeletal muscle. The use of an assay in which dephosphorylation of GS was coupled to changes in its activity, however, did reveal the activated phosphatase.23 Two GS phosphatase assays will be described, one in which phosphatase activity is determined by 32P release from 32P-GS phosphorylated at six different sites and the other, a ‘‘coupled assay,’’ in which phosphatase activity is monitored by its ability to dephosphorylate and activate fully phosphorylated and inactive GS. The first assay allows determination of the activity of protein phosphatases that dephosphorylate any of the six phosphorylation sites while the second assay detects activity of phosphatases that act on sites that control GS activity. Although the sensitivity of the first assay can be high, depending on the specific radioactivity of the substrate, detection of a specific phosphatase that acts at distinct sites may be difficult to discern over background dephosphorylation at multiple sites. On the other hand, the ‘‘coupled assay’’ could more readily detect phosphatase activity specific for GS activating sites. The disadvantage of these assays is that they are labor intensive, require several protein kinases and purification of the substrate from rabbit skeletal muscle. Expression of mammalian GS in bacteria yields only 100 g of protein per liter of culture,30 an amount insufficient for extensive study of GS phosphatases. 29 30

D. L. Brautigan and C. L. Shriner, in Methods Enzymol., 159, 339 (1988). W. Zhang, A. A. DePaoli-Roach, and P. J. Roach, Arch. Biochem. Biophys. 304, 219 (1993).

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Purification of GS

Methods for purification of rabbit skeletal muscle GS have been established by several laboratories, mostly based on isolation of glycogen to which the enzyme binds.31–33 Our currently used procedure is a modification of that previously reported by Smith et al.,34 and Takeda et al.33 For determination of phosphatase activity by 32P release from 32P-labeled GS, the dephosphorylated form of GS is purified, whereas for the ‘‘coupled assay’’ the phosphorylated form is isolated. Since the two procedures are similar, the method for isolation of the dephosphorylated form will be described in detail and only the differences will be highlighted for the purification of the phosphorylated form.

Buffers and Other Materials

TEG pH 7.8: 50 mM Tris–HCl, 5 mM EDTA, 2 mM EGTA. TEGS pH 7.8: same as TEG plus 2 mM Na2SO4. TEGS pH 6.8: same as TEGS above except pH 6.8. TEGF pH8.2: same as TEG plus 0.1 M KF. dH2O: distilled H2O. DEAE-cellulose: The DEAE-cellulose chromatography step in the purification is very critical. A course grade DEAE-cellulose is strongly recommended. The resin used in our laboratory is Selectacel DEAEstandard, available from Polysciences, Inc. (Warrington, PA). The preparation of the resin requires a lengthy procedure, 4–6 hr. At the end of the enzyme preparation, the resin is not discarded and is recycled by regeneration. Regeneration is required also for new, unused resin. 1. 2.

3.

31

Wash resin, sufficient to pack one 500 ml column, four times with 4 liter dH2O to remove fines by decantation. To the settled resin, add 4 liters of 0.3 M NaOH, mix in the beaker and filter on a Buchner funnel layered with miracloth, under vacuum generated by a H2O pump. Transfer the resin to a 4 liter beaker and repeat the alkali wash three times. Wash the resin with dH2O on the funnel until the pH of the eluate is neutral.

H. G. Nimmo, C. G. Proud, and P. Cohen, Eur. J. Biochem. 68, 21 (1976). T. R. Soderling, J. Biol. Chem. 251, 4359 (1976). 33 Y. Takeda, H. B. Brewer, Jr., and J. Larner, J. Biol. Chem. 250, 8943 (1975). 34 C. H. Smith, N. E. Brown, and J. Larner, Biochim. Biophys. Acta 242, 81 (1971). 32

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DETERMINATION, DETECTION, AND LOCALIZATION

4. 5. 6. 7. 8. 9. 10. 11. 12.

[2]

Wash twice with 1.5 liters 95% ethanol. Wash with 4 liter dH2O. Transfer the resin to 4 liters of 0.3 N HCl, mix and filter. Repeat this step three times. Wash with dH2O on funnel till eluate is neutral. Repeat the alkali wash as in step 2. Wash with dH2O till neutral. Wash with 4 liters of 2x concentrated TEGS pH 7.8. Resuspend the resin in 4 liters of TEGS pH 7.8 and stir gently overnight at 4 C. Pack 500 ml resin in a column (5  25 cm) and equilibrate with 2 liters of TEGS pH 7.8 containing 1 mM phenylmethylsulfonyl fluoride (PMSF) and 50 mM b-mercaptoethanol (b-ME).

During the HCl and NaOH washes, the DEAE-cellulose should not be kept for prolonged periods in the presence of the acid or alkali. Washes should be very rapid and the resin should only be left when neutral. Sepharose 4B column: 2.5  90 cm Sepharose CL-4B column, equilibrated overnight at room temperature with TEGS pH 7.8 containing 1 mM PMSF, 50 mM b-ME and 25% glycerol at a flow rate of 22 ml/hr.

Purification of Dephospho-GS

The procedure outlined below results in almost completely dephosphorylated and glycogen-free GS. All procedures are conducted at 4 C unless indicated otherwise. Extraction A standard preparation utilizes 4 New Zealand white rabbits, 6–8 lb each, which yield 2.5–3.0 kg muscle. The animals are injected through the marginal ear vein with lethal doses of pentobarbital (150 mg/6–8 lb) and exanguinated by severing of the jugular veins. The entire legs are quickly snapped from the joint and immediately submerged under ice. The back muscle is removed by cutting along the spine to minimize sectioning of fibers. Tissue harvesting should not exceed 8–10 min per animal, as longer times will cause prolonged muscle twitching and glycogenolysis. Loss of glycogen will result in inefficient precipitation in subsequent steps and poor enzyme recovery. The use of commercially available frozen tissue is not recommended because it leads to proteolytically degraded enzyme33 and low yields due to decreased glycogen content. After cooling, the muscle is cut into one inch pieces and homogenized in 2.5 volumes of TEG, pH 7.8 plus 1 mM b-ME using a Waring Blender, for 30 sec at low speed and then

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GLYCOGEN SYNTHASE PHOSPHATASE ACTIVITY

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60 sec at medium speed. The homogenate is centrifuged at 9000g for 50 min at 4 C and the supernatant is filtered through miracloth. Ethanol Precipitation The filtered supernatant is cooled to 0–1 C in an ice/salt bath. Care must be taken not to freeze the extract. Ethanol (95%) is then added slowly under constant stirring to a final concentration of 30%. It is critical to monitor the temperature and to keep it between 1 and 5 C during the precipitation. The use of ethanol that has been cooled at 80 C ensures that the temperature does not rise. After the final addition of ethanol, continue to stir for 10–15 min on the ice/salt bath. The suspension is then centrifuged at 9000g for 60 min at 3 C. GS Dephosphorylation The supernatant is discarded and the pale pink precipitate is suspended to a final volume of 500–600 ml of TEGS pH 6.8, 1 mM PMSF, and 50 mM b-ME, using a teflon pestle and a 50 ml glass vessel kept in ice-H2O to maintain low temperature. This procedure is facilitated by the use of a vertically mounted drill to drive the pestle. The suspension is dialyzed in a large dialysis bag for 4 hr at 4 C against 4 liters of the same buffer with one change after the first 2 hr. This step allows for dephosphorylation of GS by the endogenous phosphatases. The enzyme solution is then adjusted to pH 7.8 by the addition of
250 ml of unneutralized TEGS, 1 mM PMSF and 50 mM b-ME. The preparation is incubated at 30 C for 30 min. In fact the sample is initially placed in a 37 C H2O bath and stirred constantly till it reaches 30 C and then is transferred for 30 min to a 30 C H2O bath. Centrifuge at 9000g for 45 min at 4 C to remove precipitated material. DEAE-Cellulose Chromatography The supernatant is recovered and applied to a DEAE-cellulose column (5  25 cm, prepared as described above) at a flow rate of 60 ml/hr. After loading the sample, the column is washed first with two bed volumes of TEGS, pH 7.8, containing 1 mM PMSF and 50 mM b-ME, followed by 15 bed volumes of the same buffer plus 0.12 M NaCl at a flow rate of 120 ml/hr. This step takes usually 2.5 days. The extensive washing at this ionic strength ensures that all the glycogen phosphorylase is removed. GS is then eluted with the same buffer containing 0.25 M NaCl and 12 ml fractions are collected. Rabbit liver glycogen (Sigma), to a final concentration of 0.25 mg/ml, is added to the collection tubes, and GS activity is measured as described below, but only in the

26

DETERMINATION, DETECTION, AND LOCALIZATION

[2]

presence of G-6-P in order to expedite the analysis of the chromatographic profile. Ethanol Precipitations Fractions with GS activity are pooled and cooled to 0–1 C. Ethanol cooled to 80 C is added slowly to a final concentration of 15%. During the addition, the temperature should be kept around 2 C to facilitate precipitation of the glycogen to which the GS is bound, but the sample must not freeze. Centrifuge at 9000g for 60 min at 3 C. The supernatant is discarded and the pellet resuspended in
35 ml of TEGS pH 7.8, 1 mM PMSF, and 50 mM b-ME using a glass vessel and teflon pestle kept on ice. The ethanol precipitation is repeated as described above and the suspension is centrifuged at 22,500g for 60 min at 3 C. Alpha-amylase digestion The pellet is resuspended in 8 ml of TEGS pH 6.8, 1 mM PMSF, 50 mM b-ME, and 25% v/v glycerol and -amylase from Bacillus amyloliquefaciens (Roche Molecular Biochemicals, Indianapolis, IN) is added to a final concentration of 0.12 mg/ml. The enzyme solution is dialyzed against 500 ml of the same buffer for two days at room temperature, changing buffer every 12 hr. Visible changes in light scattering properties and viscosity of the solution should occur. During this step the glycogen is degraded and the degradation products, which could inhibit the reaction, are removed by dialysis. The presence of 25% glycerol prevents enzyme aggregation and allows this and the next step to be conducted at room temperature. Sepharose CL-4B Chromatography After dialysis, the sample is centrifuged for 15 min on a bench top centrifuge to remove insoluble material and the supernatant is applied at room temperature to a 2.5  90 cm Sepharose CL-4B column equilibrated overnight with TEGS pH 7.8, 1 mM PMSF, 50 mM b-ME, and 25% v/v glycerol at a flow rate of 20 ml/hr. The column is developed at the same flow rate and 4 ml fractions are collected. The OD280 of the fractions is monitored and glycogen synthase activity is measured. Fractions containing enzyme with similar specific activity are pooled and stored at 80 C. Under these conditions the enzyme is stable for at least one year. Note that at all steps during the preparation, GS activity, protein concentration and volumes should be closely monitored in order to estimate enzyme yield, purification and specific activity. As a rule of thumb, for measurement of GS activity, an aliquot of the initial extract is diluted 1:10, the sample after the first ethanol precipitation 1:100, the

[2]

27

GLYCOGEN SYNTHASE PHOSPHATASE ACTIVITY

TABLE I PURIFICATION OF DEPHOSPHORYLATED GLYCOGEN SYNTHASE FROM RABBIT SKELETAL MUSCLEa

Extract 30% Ethanol Loaded onto DEAE-Cellulose DEAE-Cellulose 15% Ethanol Sepharose-CL-4B

Protein (mg)

/ þ G-6-P ratio

Specific activity (U/mg)

Yield (%)

112,650 32,400 14,820

0.54 0.44 0.71

0.04 0.14 0.12

100 92 37

313 25.6 11.1

0.70 0.60 0.80

1.21 22.3 40.9

8 12 10

a This preparation utilized 2.7 kg of rabbit skeletal muscle. GS activity is measured as described in the text.

DEAE-cellulose column fractions 1 : 5, the sample after the second 15% ethanol precipitation 1 : 300 and the fractions from the Sepharose CL-4B chromatography 1:100, in order to be in the linear range of the reaction. On average, 10–20 mg of GS with a specific activity between 30–40 U/mg and a / þ G-6-P ratio of 0.8 as measured in the absence of SO2 4 can be obtained from 2.5 to 3.0 kg of muscle (Table I). As judged by SDS-PAGE and Coomassie Blue staining the preparation is more than 95% pure (Fig. 2) and the phosphate content less than 0.5 mol/mol GS protein.35 It should also be noted that in all the procedures described in this chapter, the rabbit liver glycogen used has been deionized by passage through a mixed bed ionic exchange resin (Amberlite MB-3A, Aldrich). The deionization will remove ions that may otherwise affect GS and phosphatase activity. The eluted glycogen is precipitated with 66% ethanol, completely dried to remove all traces of ethanol and stored at 20 C.

Purification of Phospho-GS

The purification procedure for phosphorylated GS is very similar to that of the dephosphorylated form and only the differences are described. A.

35

The extraction step is the same as for the dephospho-GS except that the buffer contains 0.1 M KF and the pH is 8.2. These conditions

P. J. Roach, Y. Takeda, and J. Larner, J. Biol. Chem. 251, 1913 (1976).

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DETERMINATION, DETECTION, AND LOCALIZATION

[2]

FIG. 2. SDS-polyacrylamide gel electrophoresis of purified phospho- and dephospho-GS. Lanes 1 and 2, 0.64 g each of dephospho-GS before and after phosphorylation by CKII, GSK-3 and PhK. Note the reduction in the electrophoretic mobility of the phosphorylated form compared to the dephosphorylated protein. Lanes 3 and 4, 1 g each of the partially purified phospho-GS before and after further phosphorylation, respectively. Note that this form of GS shows a reduced mobility that is not significantly affected by further phosphorylation. Lane M corresponds to the molecular weight marker proteins whose masses in kilodaltons (kDa) are indicated. The samples were separated on a 6.5% polyacrylamide gel according to U. K. Laemmli, Nature 227, 680 (1970) and stained with Coomassie Blue. Protein concentration was determined by the method of M. M. Bradford, Anal. Biochem. 72, 248 (1976).

will inhibit phosphatase activity and hence the dephosphorylation of GS during the homogenization. B. After the 30% ethanol precipitation, the precipitate is resuspended in TEG pH 7.8, 1 mM PMSF, and 50 mM b-ME and directly incubated at 30 C for 30 min without prior dialysis. The latter step would have favored dephosphorylation which is not desirable for the preparation of the phospho-GS. C. During the DEAE-cellulose chromatography, there is no Na2SO4 in the buffers and the enzyme is eluted with 0.3 M NaCl. D. The DEAE-cellulose fractions containing GS activity are pooled and precipitated with 30% ethanol. The precipitate is resuspended in
30 ml of TEG pH 7.8, 1 mM PMSF, and 50 mM b-ME and incubated overnight at 4 C in the presence of 5 mM ATP, 10 M cAMP, 12 mM MgCl2. This step will allow phosphorylation of the GS by copurified protein kinases. E. Finally, the enzyme is precipitated twice with 15% ethanol, resuspended in 10 ml of TEG pH 7.8, 1 mM PMSF, 50 mM b-ME, and 10% v/v glycerol and stored at 80 C. Further purification of the enzyme is not required for the ‘‘coupled assay.’’ Starting with
1.5 kg of skeletal muscle, this preparation results in GS with / þ G-6-P ratio of 0.1 and specific activity 7.8 U/mg (Table II and Fig. 2).

[2]

29

GLYCOGEN SYNTHASE PHOSPHATASE ACTIVITY

TABLE II PURIFICATION OF PHOSPHORYLATED GLYCOGEN SYNTHASE FROM RABBIT SKELETAL MUSCLEa

Extract 30% Ethanol Loaded onto DEAE-Cellulose DEAE-Cellulose 30% Ethanol 15% Ethanol

Protein (mg)

/ þ G-6-P ratio

Specific activity (U/mg)

Yield (%)

53,360 16,764 6,798 214 94 63

0.2 0.1 0.1 0.1 0.1 0.1

0.02 0.06 0.07 2.2 6.0 7.8

100 95 46 46 52 46

a

This preparation utilized 1.5 kg of rabbit skeletal muscle. GS activity is measured as described in the text.

Determination of GS Activity

GS activity is measured by incorporation of [14C]glucose from UDP[U-14C]glucose into glycogen as described by Thomas et al.36 The reaction mixture (90 l) contains 50 mM Tris–HCl pH 7.8, 15 mM EDTA, 0.6 mM EGTA, 50 mM KF, 7 mg/ml deionized rabbit liver glycogen, 4.4 mM UDP[U-14C]glucose, specific radioactivity 400 cpm/nmol in the absence or presence of 7.2 mM G-6-P and appropriately diluted enzyme. The reaction is started by addition of 30 l GS immediately diluted in ice-cold buffer containing 50 mM Tris–HCl pH 7.8, 10 mM EDTA, 2 mM EGTA, 100 mM KF, and 50 mM b-ME. The reaction is carried out at 30 C for 10–15 min and terminated by placing 75 l of the reaction mixture on a 2  2 cm 31ET filter paper, which is immediately dunked into 20 C cold 66% ethanol. The 31ET paper is thicker and more porous than the 3MM and therefore traps better the precipitated glycogen. The filters are washed in 66% ethanol three times with stirring for 10, 60, and 30 min, respectively, followed by 10 min in acetone, dried and counted in a liquid scintillation counter. The second and third washes are conducted in room temperature ethanol. No more than 15% of the substrate should be used in the reaction to ensure linearity. One unit of GS is the amount of enzyme that incorporates 1 mol of glucose from UDP-glucose into glycogen/min.

36

J. A. Thomas, K. K. Schlender, and J. Larner, Anal. Biochem. 25, 486 (1968).

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DETERMINATION, DETECTION, AND LOCALIZATION

[2]

Phosphorylation of GS Further Phosphorylation of Phospho-GS for the ‘‘Coupled Assay’’

The preparation of the phospho-GS results in enzyme with a / þ G-6-P ratio of
0.1, indicating that the enzyme is not fully phosphorylated. In order to lower the activity ratio, which is desirable for determination of phosphatase activity based on activation of GS, the partially phosphorylated GS (3.2 mg/ml) is incubated for 2 hr at 30 C in the presence of 8 g/ml CKII (specific activity 830 nmol/min/mg), 5 g/ml GSK-3b (specific activity 820 nmol/min/mg), 5 g/ml PhK (specific activity 565 nmol/min/mg), 0.1 M NaCl, 1.5 mM CaCl2, 0.1 mM ATP, 10 mM MgCl2 in 50 mM Tris–HCl pH 7.5, 0.5 mM EDTA, 1 mM EGTA, 0.5 mM PMSF, 50 mM b-ME, and 5% v/v glycerol. The reaction is terminated by addition of EDTA to a final concentration of 10 mM. GS is then precipitated twice with 15% ethanol and finally resuspended in 50 mM Tris–HCl pH 7.8, 1 mM EGTA, 0.5 mM EDTA, 4 nM okadaic acid, 1 mM PMSF, 50 mM b-ME and 10% v/v glycerol and stored in 100 l aliquots at 80 C. After the phosphorylation and two 15% ethanol precipitations, the / þ G-6-P activity ratio is decreased to 0.02 and the specific activity increased to 13.9 U/mg. 32

P-labeling of Dephospho-GS

Dephospho-GS is dialyzed overnight against 4 liters of 30 mM Tris–HCl pH 7.5, 1 mM EDTA, 0.5 mM EGTA, 50 mM b-ME, 1 mM PMSF, and 10% v/v glycerol with two changes of buffer. The dialyzed GS is incubated, at a concentration of 0.2 mg/ml, for 2 hr at 30 C in a 4 ml reaction containing 50 mM Tris–HCl pH 7.5, 0.25 mM EGTA, 0.5 mM EDTA, 0.5 mM PMSF, 5% v/v glycerol, 5 g/ml GSK-3b, 8 g/ml CKII, 5 g/ml PhK, 0.1 M NaCl, 0.1 mM [ -32P]ATP (
4000 cpm/pmol), 10 mM MgCl2 and 1.5 mM CaCl2. The level of phosphorylation is monitored by placing 10 l of the reaction mixture 1.0 cm from the bottom of a 2  10 cm instant thin-layer-chromatography paper strip (Gelman Sciences) and developing the strip in 5% TCA and 0.2 M KCl for 7 min. By this procedure, the unreacted [ -32P]ATP migrates to the top while the phosphorylated GS remains at the origin. The strips are dried and the region containing the phosphorylated protein is cut and counted. Appropriate controls for [ -32P]ATP blanks are also processed and their value is subtracted from that obtained for the phosphorylated samples. Under these conditions, normally
6 mol of 32P per mol GS are incorporated. As shown in Fig. 2, phosphorylation results in a reduced electrophoretic mobility. The unreacted [ -32P]ATP in the preparation is

[2]

GLYCOGEN SYNTHASE PHOSPHATASE ACTIVITY

31

removed by size exclusion chromatography on two 10 ml Sephadex G50-150 columns (0.8  20 cm). The fractions (1 ml each) containing the 32P-GS are pooled, concentrated by ultrafiltration using Centricon 100 (Amicon), dialyzed for 36 hr against 30 mM Tris–HCl pH 7.5, 1 mM EDTA, 0.5 mM EGTA, 50 mM b-ME, and 5% v/v glycerol and stored on ice for up to 1 month. Protein Kinases

Phk, specific activity of 565 nmol/min/mg with Ph b as substrate, is purified from rabbit skeletal muscle by the procedure of Cohen.37 Recombinant GSK-3b expressed in E. coli is purified as previously reported.38 The purified enzyme has a specific activity of 820 nmol/min/mg utilizing as substrate 0.1 mM of the synthetic phosphopeptide RRAAEELDSRAGS(P)PQL, derived from the GSK-3 phosphorylation site of the eukaryotic initiation factor 2B.39 The  and b subunits of CKII expressed in E. coli are purified as described.40 After purification of the individual subunits, the holoenzyme is reconstituted by combining equimolar amounts of the two subunits. The resulting enzyme has a specific activity of 830 nmol/min/mg with 0.1 mM of the peptide substrate RRRDDDSDDD. GS Phosphatase Assays 32

P Release from

32

P-GS

GS phosphatase activity is measured in extracts of mouse skeletal muscle rapidly frozen in liquid N2 and stored at 80 C. Powdered tissue samples (
40 mg) are homogenized in 10 volumes of 50 mM Tris–HCl pH 7.5, 0.1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1 mM N-p-tosyl-L-lysine-chloromethyl ketone, 2 mM benzamidine, 0.5 mM PMSF, 10 g/ml leupeptin, 50 mM b-ME, 2 mg/ml deionized rabbit liver glycogen, for 20 sec with a BioSpec Tissue Tearor at 30,000 rpm and centrifuged at 600g for 10 min. The supernatant is filtered through glass wool and diluted 1:5 in homogenization buffer. Ten microliters are then added to 10 l of a solution containing 50 mM Tris–HCl pH 7.5, 0.5 mM EDTA, 37

P. Cohen, Eur. J. Biochem. 34, 1 (1973). Q. M. Wang, C. J. Fiol, A. A. DePaoli-Roach, and P. J. Roach, J. Biol. Chem. 269, 14566 (1994). 39 G. I. Welsh, J. C. Patel, and C. G. Proud, Anal. Biochem. 244, 16 (1997). 40 D. Li, G. Dobrowolska, and E. G. Krebs, J. Biol. Chem. 271, 15662 (1996). 38

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DETERMINATION, DETECTION, AND LOCALIZATION

[2]

FIG. 3. GS phosphatase activity. GS phosphatase activity was measured in mouse skeletal muscle extracts by monitoring the release of 32P from 32P-GS (Panel A) and by the dephosphorylation/activation coupled assay (Panel B). The activity was determined in the absence (C) or the presence of 4 nM okadaic acid (OA) and/or 1 M inhibitor-2 (I-2) as described in the text. Statistical significance was assessed by Student’s t-test (* , P 590 nm), respectively. Since the emission spectra of the fluorescein and rhodamine overlap, and both molecules can be excited by the use of both laser beams, correction factors have to be introduced. These factors are S1, S2, and S3. The definition of these parameters are: S1 ¼ I2/I1 S2 ¼ I2/I3 S3 ¼ I3/I1

determined using only donor labeled cells determined using only acceptor labeled cells determined using only donor labeled cells.

The three detected intensities can be expressed as: I1 ð488 ! 535Þ ¼ IF ð1
E Þ

ð11Þ

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APPLICATION OF FRET TO PHOSPHATASES

209

I2 ð488 ! > 590Þ ¼ IF ð1
EÞS1 þ IR S2 þ IF E

ð12Þ

S3 IF E S1

ð13Þ

I3 ð514 ! > 590Þ ¼ IF ð1
EÞS3 þ IR þ

I1 is smaller than IF because the energy transfer causes donor quenching. I2 consists of three additive terms: (i) the overlapping fraction of the quenched fluorescein intensity, (ii) the direct contribution of rhodamine and (iii) sensitized emission due to energy transfer. The proportionality factor  is the ratio of I2 for a given number of excited rhodamine molecules and I1 for the same number of excited fluorescein molecules.  is constant for each experimental setup, and has to be determined for every defined case. I3 is a sum of (i) a fraction of the quenched fluorescein intensity, (ii) the rhodamine intensity and (iii) the sensitized emission of rhodamine due to energy transfer corrected for the lower molar extinction coefficient of fluorescein at 514 nm than at 488 nm. From the Eqs. (11)–(13) E can be expressed as follows: E¼

A 1þA

ð14Þ

Based on Eq. (14) the transfer efficiency can be calculated on a cell-by-cell basis, resulting in frequency distribution histograms. The generated distribution histogram of FRET efficiency will provide information about the heterogeneity of the cell population with high statistical accuracy. Usually it is assumed that cellular autofluorescence is negligible compared to the specific fluorescence. If autofluorescence is not negligible, appropriate corrections should be performed. When the autofluorescence is substantial, but the signal to autofluorescence ratio is above four, the correction for autofluorescence can be done using the average autofluorescence intensities of the cell population. If the autofluorescence is more significant, such that its value is comparable with the specific signal, a more elaborate correction method is also possible, since in most cell types there is a good correlation between the autofluorescence detected at different spectral regions. In such situations a further independent parameter should be detected, a fourth fluorescence intensity, and the autofluorescence content of specific fluorescence signals can then be corrected on a cell-by-cell basis. In general it should be remembered that naturally high autofluorescence is likely to decrease the precision of the measurements. Using this approach, flow cytometry can provide quantitative measurements on individual cells, allowing convenient determination of the distribution of energy transfer efficiency values in a population. In most

210

INTERACTING PROTEINS AND SUBUNITS

[17]

flow cytometric energy transfer experiments fluorescein is used as donor and rhodamine as acceptor. However, the equations shown are equally valid for other donor acceptor pairs, providing that the different spectral characteristics are considered. Applying the Cy3 and Cy5 donor/acceptor pair has several advantages over the classical fluorescein–rhodamine pair: (i) Cy3 and Cy5 have higher absorption coefficients and better photostability, (ii) both the donor and the acceptor emissions are shifted towards the red region, so autofluorescence is much less of a problem, (iii) even in commercial flow cytometers, the acceptor (Cy5) can be excited independently from Cy3, so that the S3 value becomes zero, making simpler the equations used for analysis. Based on such considerations, a detailed description of an improved flow cytometric energy transfer method has recently been published.8 The main advantage of the approach is reduction of autofluorescence-related errors, which is achieved by application of long wavelength dyes, such as Cy3 and Cy5, and by cell-by-cell correction of autofluorescence. The improved flow cytometric energy transfer method allows energy transfer efficiencies to be determined on cellular systems even when the expression of cell-surface molecules is low. In addition, the lower variance of energy transfer efficiency distributions enables a much more accurate discrimination of subpopulations having distinct FRET efficiencies. Method Outline for FRET Measurements Using Flow Cytometry

Although experimental protocols will vary considerably depending on the precise cellular system being analyzed, the following method can be used for the analysis of cell-surface associations between different cell surface proteins. Step 1: Prepare minimum set of samples: unlabeled cells, cells labeled with donor (e.g., Cy3) only, cell labeled with acceptor (e.g., Cy5) only, and cells labeled with both donor and acceptor. Use antibodies, preferably F(ab0 ) fragments, directly conjugated with fluorescent dyes. If the expression levels of proteins under investigation are comparable, prepare an additional sample for which proteins originally labeled with the donor-conjugated antibody will be labeled with acceptor-conjugated antibody and vice versa. If the expression level of the two proteins under investigation is different, then the protein with higher expression level should be chosen as acceptor. Step 2: Analyze the samples on a flow cytometer having at least two excitation wavelengths, one for the donor (e.g., 488 nm) and one for the 8

Z. Sebestye´n, P. Nagy, G. Horva´th, G. Va´mosi, R. Debets, J. W. Gratama, D. R. Alexander, 00 and J. Szo¨llosi, Cytometry 48, 124 (2002).

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acceptor (e.g., 632 nm). Choose the appropriate filters according to your donor and acceptor dye. Collect at least three fluorescence intensities (for autofluorescence correction, collect four fluorescence intensities). Whenever possible use linear gain, avoiding logarithmic gain. Do not use hardware compensation. Step 3: Store the data in list mode. Analyze the unlabeled and single labeled cell to determine the spectral overlap factors. Step 4: Finally analyze the double labeled (FRET) sample, and calculate the FRET efficiency distribution histogram.

Donor and Acceptor Photobleaching FRET Microscopy

While many cells can be analyzed by flow cytometry within a short timeframe, flow cytometry lacks the ability to reveal the distribution of protein associations within a single cell. Donor photobleaching fluorescence resonance energy transfer (pbFRET) is a quantitative approach with the added value of not requiring sophisticated and expensive instrumentation. Jovin and Arndt-Jovin introduced the pbFRET method to determine transfer efficiency on a pixel-by-pixel basis in a microscope.9 FRET efficiency is calculated from the photobleaching kinetics of the donor in the presence and absence of the acceptor. Photobleaching is an irreversible conversion of a dye from a fluorescent to a nonfluorescent molecule, starting with the excited state of the molecule. In the case of energy transfer, there is an extra possibility for de-excitation of the donor, therefore the donor will spend less time in the excited state. Since the photobleaching rate depends upon how long the donor is in the excited state, the presence of acceptor within the FRET distance will slow down the photobleaching rate of the donor. Assuming that the photobleaching of the donor occurs from the excited singlet state, FRET efficiency can be calculated as follows:

E ¼1


TD , TDA

ð15Þ

where E is the FRET efficiency, TD-s are the photobleaching time constants of the donor and the upper index indicates the presence of the acceptor. Photobleaching kinetics in pixels or regions of interests (ROIs) is measured on donor labeled and donor–acceptor double-labeled samples. After completing the photobleaching, the residual intensity is subtracted 9

T. M. Jovin and D. J. Arndt-Jovin, Ann. Rev. Biophys. Chem. 18, 271 (1989).

212

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INTERACTING PROTEINS AND SUBUNITS

from all images on a pixel-by-pixel basis and the resulting bleaching curves are fitted to a double exponential equation, which usually yields better results than single exponential fits. From the two exponential time constants an effective photobleaching time constant can be calculated by taking an amplitude weighted average: IðtÞ ¼ I0 þ I1  e
ðt=1 Þ þ I2  e
ðt=2 Þ , Teff ¼

I1 1 þ I2 2 I1 þ I2

ð16Þ

where I0 is the residual unbleachable intensity, which should be subtracted from each pixel before the bleaching curves are fitted to the double exponential equation. I1 and I2 are the amplitudes of the components having different bleaching time constants. Frequency distribution histograms of the time constant can be prepared and mean values of the histograms are calculated. From these mean values the average transfer efficiency can be calculated using Eq. (16). From the average energy transfer efficiency, the FRET efficiency assigned to each pixel can also be determined and a two dimensional FRET map can be generated showing the intracellular heterogeneity of this parameter. The photophysics of bleaching is far from being understood and sometimes results give anomalously high values, especially when FRET efficiency is measured between very dense cell surface antigens. In the transition region where the photobleaching changes from a unimolecular to a bimolecular reaction, interpretation of results may be difficult. The sensitivity of pbFRET to environmental factors (such as O2 concentration) may also raise some problems if this parameter cannot be controlled in the donor only and the donor–acceptor double-labeled samples in the same way. Since photobleaching of common dyes is irreversible, it is not possible to repeat pbFRET measurements on the same cells, so the kinetics of the association processes of membrane components cannot be monitored. Some of these disadvantages can be overcome by applying the intensity based FRET microscopic method (described later). Method Outline for Photobleaching FRET Microscopy

Step 1: Prepare a minimum set of samples: cells labeled with donor (e.g., fluorescein) only, and cells labeled with both donor and acceptor (rhodamine or Cy3). (Unlabeled cells should also be used occasionally to check the signal level for the donor.) Choose a fluorescent donor which can be bleached at a convenient rate (not too fast and not too slow) with the available fluorescence microscope. If the association of other protein

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APPLICATION OF FRET TO PHOSPHATASES

213

pairs is investigated, the samples should be prepared in pairs: the same donor-conjugated mAbs or F(ab0 ) fragments should be used in the donor only, and also in the donor and acceptor samples. Step 2: Take sequential donor images of the donor-only sample and the donor and acceptor labeled sample (double labeled sample), using excitation for the donor while bleaching away the donor signal. Step 3: Check the images for pixel shift. If pixel shift can be observed, the shifted images should be corrected before data processing. Step 4: Generate fluorescence intensity curves for each pixel of the images and fit the curves to double exponential equations. For data processing select the areas of the image where the donor signal is above background. Step 5: Calculate the effective rate constant by taking the amplitudeweighted time constant of the exponential terms. Step 6: Calculate the FRET efficiency on a pixel-by-pixel basis by comparing the rate of photobleaching in the double-labeled sample to that in the donor-only sample. The acceptor bleaching based method, using Cy3 and Cy5 as the donor– acceptor pair, was applied successfully for determination of FRET efficiency on a pixel-by-pixel basis by Wouters et al.10 Microscopic images of donor and acceptor labeled cells were used in the following sequence. A prephotobleach donor (Cy3) image was acquired by scanning at the excitation wavelength of the donor (543 nm for Cy3) in the confocal laser scanning microscope (CLSM). Then an acceptor (Cy5) image was taken by scanning at the acceptor excitation wavelength (633 nm for Cy5). On the basis of acceptor image areas of the original image, fields were selected in which the acceptor fluorophores were subsequently photobleached by repeated scanning at the excitation wavelength of the acceptor, thereby abolishing FRET. After acceptor photobleaching, a second donor image was taken at the donor excitation wavelength. An increase of donor fluorescence intensity should be observed in the region of acceptor photobleaching where cellular structures exhibited FRET. After correction for image registration, the FRET efficiencies (Ei) in pixel i were calculated from image arithmetic of the two (prephotobleach, Ipre,i and postphotobleach Ipost,i) donor images: Ei ¼

Ipost,i
Ipre,i : Ipost,i

ð17Þ

If the acceptor is photobleached only in part of the image, the complementary part can be used for donor photobleaching. This approach 10

F. S. Wouters, P. I. H. Bastiaens, K. W. A. Wirtz, and T. M. Jovin, EMBO J. 17, 7179 (1998).

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INTERACTING PROTEINS AND SUBUNITS

[17]

can provide a second independent assessment of FRET in cells. Combinations of the donor and acceptor photobleaching can provide more solid bases for the association of proteins under investigation.

Intensity Based FRET Microscopy

Recently an intensity based FRET measurement was elaborated using acceptor sensitization to estimate FRET efficiency on a pixel-by-pixel basis without losing temporal resolution.11 In order to achieve quantitative FRET measurements the autofluorescence of cells had to be corrected on a pixelby-pixel basis. The fraction of autofluorescence appearing in various detection channels (blue emission, i.e., pure autofluorescence, fluorescein, rhodamine and energy transfer) can be determined, since the autofluorescence spectrum of cells has a wide excitation and emission spectrum and is fairly constant. It is possible to define ratios that describe the ratio of fluorescence intensities recorded in fluorescein, energy transfer and rhodamine detection channels to that detected in the blue channel using unlabeled cells and UV excitation. This approach is valid because the fluorescent dyes applied have negligible absorption in the relevant UV range. This correction eliminates the contribution of autofluorescence from the fluorescein, rhodamine and FRET images on a pixel-by-pixel basis. From the fluorescein, rhodamine and FRET images corrected for autofluorescence, the FRET efficiency can be calculated on a pixel-bypixel basis using an equation set similar to that applied in flow cytometry (Eqs. (11)–(13)). The exact experimental setup and the detailed description of calculations are given in reference.11 Although photobleaching is useful in pbFRET, it presents a problem in the case of intensity-based measurements. When the energy transfer values are calculated using the intensity-based approach, correction for photobleaching should be taken into account. Since the bleaching rate of rhodamine is very low with illumination intensities commonly used in such experiments, correction should be applied only for the fluorescein and the energy transfer images. Fluorescein and energy transfer images are sequentially recorded. When calculating the energy transfer in an image, the relevant fluorescein intensity is computed as the average of the previous and the next fluorescein image. This linear approximation of bleaching is reasonable only for short exposure times. From these images the FRET efficiency can be calculated on a pixel-by-pixel basis. 11

00

P. Nagy, G. Va´mosi, A. Bodna´r, S. J. Lockett, and J. Szo¨llosi, Eur. Biophys. J. 27, 377 (1998).

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Comparison between flow cytometric FRET measurements and the different image cytometric energy transfer calculation methods has revealed that consistently higher transfer values are obtained with the pbFRET method. It has been shown that some of this discrepancy may be attributed to the different weighting of the energy transfer values by the pbFRET and the intensity-based energy transfer methods. Using mathematical models applying Monte Carlo simulation it was demonstrated that this overestimation in pbFRET is proportional to the heterogeneity in the pixel-bypixel FRET efficiency values. Therefore differences in FRET efficiency values provided by the pbFRET and the intensity-based approaches should be interpreted with caution.11

Method Outline for Intensity Based FRET Microscopy

Step 1: Prepare a minimum set of samples: unlabeled cells, cells labeled with donor (e.g., fluorescein) only, cell labeled with acceptor (e.g., rhodamine) only, and cells labeled with both donor and acceptor. Step 2: Choose the appropriate excitation, emission and dichroic filters according to your donor and acceptor dye. Collect at least three images (donor, acceptor, and FRET). (For autofluorescence correction collect four images, one extra for autofluorescence.) Take the autofluorescence image first, then the acceptor (rhodamine) and then the donor and FRET images alternatively if there is some photobleaching. Step 3: Correct all the images for camera dark current by subtracting images taken with closed shutters. Step 4: Because of the filter changes (especially for dichroic filters) check images for image registration. Correct images for any pixel shift. Step 5: Correct images for the background caused by the fluorescence of the optical elements of the microscope, using cell free areas of the images. Step 6: Analyze unlabeled cells first, determining the spectral overlap factors of autofluorescence. Step 7: Analyze donor-only and acceptor-only samples. Using all four images, correct the images for autofluorescence first, then determine the spectral overlap factors originating from the donor spectrum in the case of the donor-only sample and from the acceptor spectrum in the case of the acceptor-only sample. Step 8: Analyze all four images of the double-labeled samples, correct for autofluorescence, then for donor emission and acceptor emission. After all the corrections, calculate the FRET efficiency on a pixel-by-pixel basis. If there was photobleaching during image recording, use the

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average of donor images and the average of FRET images taken alternatively.

Limitations of FRET Studies

For FRET applications to have appropriate experimental interpretations, it is important to understand its limitations as well. The most serious drawback of FRET is its modest capacity to determine absolute distances. It is quite good at determining relative distances, namely, whether two points are getting closer or further apart following a stimulus. This is caused by the fact that FRET efficiency depends not only on the distance between the donor and acceptor, but also on the relative orientation (2) of the dyes as well (as discussed above). Even when measuring relative distances, care must be taken to ensure that the orientation factor (2) does not change between the two systems to be compared. In addition, the system can be more complicated when randomly conjugated fluorophore is used. For example, in a substantial fraction of experiments, fluorophore-conjugated mAbs are used to label cell surface antigens. According to common practice, covalent coupling reactions occur frequently between isothiocyanate or succinimidyl ester reactive groups of appropriate derivatives of the fluorophore and the "-amino groups of lysine side chains of immunoglobulins. The number of the exposed lysine side chains of comparable reactivity of an antibody molecule usually exceeds one. Each antibody molecule may therefore carry several fluorophores and the labeled lysine side chains of the individual antibody molecules may be different. Reactive groups are often attached to the fluorophores via an n-carbon linker, for which n typically ranges from 2 to 12. The linker can allow relatively free rotation of the dye, which minimizes uncertainty of 2. It also minimizes quenching of the dye by the protein, especially if that is due to a hydrophobic environment. The linker, however, has a disadvantage of adding uncertainty to the exact position of the dye. In general the minimal length that allows free rotation of the dyes and does not cause quenching is around six-carbon atoms long. Indirect immunofluorescent labeling strategies may be applied to FRET measurements if suitable directly fluorophore-conjugated mAbs are not available, or as an approach to enhancing the specific fluorescence signal. In such cases special attention should be paid to the fact that the size of the antibody complexes used affects the measured FRET efficiency values (Fig. 2). Application of a larger antibody complex causes a decrease in FRET efficiency due to the geometry of the antibody complexes since when antibody or F(ab0 ) complexes become larger, the actual distance between the donor and acceptor fluorophores increases.8 This explains the decreased

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FIG. 2. Labeling strategy affects the efficiency of FRET. The top half of the figure shows the labeling strategy. The heavy chain and light chain (b2m) of class I HLA molecules were labeled directly with fluorescently tagged primary mAb, indirectly with unlabeled mAb and subsequently with fluorescent F(ab0 ) fragments of goat-anti-mouse IgG or whole rat-antimouse IgG molecules. In the case of indirect staining more than one polyclonal antibody or F(ab0 ) fragment can bind to a single primary antibody. For the sake of simplicity only one secondary antibody or F(ab0 ) fragment is shown in the scheme. In these measurements Cy3 dyes were used as donors and Cy5 dyes as acceptors. The bottom half of the figure shows histograms displaying FRET efficiency distributions measured between the light and the heavy chains of class I HLA molecules using direct and indirect labeling. The first distribution curve on the left represents the FRET distribution of cells in the absence of energy transfer when cells were labeled only with a donor-conjugated primary mAb. Note that upon increasing the complexity of the labeling scheme by introducing secondary antibodies the transfer efficiency values decrease, the FRET efficiency histograms shifting to the left.

FRET efficiency when fluorescent secondary F(ab0 ) fragments are used on both the donor and the acceptor side relative to primary fluorescent antibodies. It also explains a further decrease in FRET efficiency when intact fluorescent secondary antibodies are used. Such findings underline the fact that FRET values cannot be directly compared to each other if they are obtained using different labeling strategies.8

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Another problem is that FRET has a very sharp distance dependence. It is therefore difficult to measure relatively long distances because the signal becomes very weak. At the same time energy transfer tends to be all or none: if the donor and acceptor are within 1.63  Ro distance there is energy transfer, if they are farther apart, energy is transferred with very little efficiency. For these reasons the failure to detect FRET from a pair of labeled proteins carries no intrinsic information regarding the association of the proteins. An absence of detectable FRET cannot be taken as formal exclusion of protein–protein associations. When studying cells labeled with donor and acceptor conjugated mAbs, averaging is performed at different levels. The first averaging follows from the random conjugation of the fluorescent label. An additional averaging is brought about by the eventual distribution of separation distances between the epitopes labeled with mAbs. This multiple averaging, an inevitable consequence of the nonuniform stoichiometry, explains why the experimental goals of FRET measurements vary depending on whether purified molecular systems are being used, or whether the associations of cell-surface molecules are being examined. In the former case FRET efficiency values can be converted into absolute distances. Calculation of distance relationships from energy transfer efficiencies is easy in the case of a single-donor single-acceptor system if the localization and relative orientation of the fluorophores are known. However, if cell membrane components are investigated, a twodimensional restriction applies to the labeled molecules. Analytical solutions for randomly distributed donor and acceptor molecules and numerical solutions for nonrandom distribution have been elaborated by different groups.12–14 In order to differentiate between random and nonrandom distributions, energy transfer efficiencies have to be determined at different acceptor concentrations.

The Application of FRET to Studying Phosphatases CD45 and CD45 Isoforms

CD45 is a transmembrane phosphotyrosine phosphatase that is expressed on all nucleated hematopoietic cells. Its phosphatase activity is required for efficient signal transduction by the antigen receptors that

12

B. Snyder and E. Freire, Biophys. J. 40, 137 (1982). J. Yguerabide, Biophys. J. 66, 683 (1994). 14 A. K. Kenworthy and M. Edidin, J. Cell Biol. 142, 69 (1998). 13

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mediate lymphocyte development and activation.15–17 Alternative splicing of CD45 exons 4–6 (A, B and C) generates up to eight different CD45 isoforms of which five are expressed at significant levels in T cells. The isoforms vary only in their ectodomains which are markedly different in terms of size and glycosylation status. All T cells express more than one CD45 isoform, and differential isoform expression is tightly controlled during thymic development and the activation of matured T cells (reviewed in Refs. 15 and 16). It has been suggested that the actions and/ or activity of CD45 phosphotyrosine phosphatase might be regulated by the extracellular domain of CD45 through influencing dimerization. Homo- and heterodimerization of CD45 molecules were previously detected using immunoprecipitation and chemical crosslinking.17–19 The application of FRET techniques to live cells in such contexts has the advantage of avoiding the use of detergent and the consequent disruption of membranes. The first FRET measurement between cell surface molecules using CD45 as one of the possible association partners was performed by Mittler et al.20 In their previous study flow cytometric energy transfer values demonstrated physical association between T cell antigen receptor (TCR) and CD4 molecules on helper T cells. Energy transfer was not detected between the TCR-CD3 complex and CD45, despite of the fact that the CD45 is abundantly expressed on the T cell surface.21 However, when human peripheral blood lymphocytes were activated using solid phase CD3 mAbs, significant energy transfer could subsequently be measured between CD4 or CD8 molecules (labeled with fluoresceinated mAbs) and CD45 (labeled with rhodaminated mAbs).20 Maximal association occurred 72–96 hr after exposure to CD3 mAbs on both CD4 þ and CD8 þ T cells. Flow cytometric FRET data were confirmed by immunoprecipitation of dithiobis succinimidyl propionate or disuccinimidyl suberate cross-linked resting or activated T cells labeled with 125I. It should be noted that the authors did not report whether the various CD45 isoforms differentially associated with CD4/CD8.

15

D. R. Alexander, in ‘‘Lymphocyte Signalling: Mechanism, Subversion, and Manipulation’’ (M. M. Harnett and K. P. Rigley, eds.), p. 107. John Wiley and Sons, New York, 1997. 16 D. R. Alexander, Sem. Immunol. 12, 349 (2000). 17 Z. Xu and A. Weiss, Nature Immunol. 3, 764 (2002). 18 M. Bonnard, C. R. Maroun, and M. Julius, Cell. Immunol. 175, 1 (1997). 19 A. Takeda, J. J. Wu, and A. L. Maizel, J. Biol. Chem. 267, 16651 (1992). 20 R. S. Mittler, B. M. Rankin, and P. A. Kiener, J. Immunol. 147, 3434 (1991). 21 R. S. Mittler, S. J. Goldman, G. L. Spitalny, and S. J. Burakoff, Proc. Natl. Acad. Sci. U.S.A. 86, 8531 (1989).

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Using both immunoprecipitation and flow cytometric FRET techniques, Lazarovits et al. studied the association pattern of CD7 on the cell surface.22 CD7 is a 40 kDa glycoprotein that is expressed on a major subset of human peripheral blood T cells. Ligation of CD7 using a cross-linked mAb contributed to mitogenesis, and signals delivered via CD7 stimulated integrin-mediated adhesion. Coimmunoprecipitation data suggested that CD7 associates with CD3 and CD45. To confirm this observation flow cytometric energy transfer measurements were performed using fluorescein (F) and rhodamine (R) labeled mAbs as donor acceptor pairs. There was significant increase in the sensitized emission when F-CD7 and R-CD45, or F-CD7 and R-CD3 interaction was investigated, indicating molecular associations between these entities. These data supported the hypothesis that CD7 exists in an oligomeric complex with CD3/TCR, CD45, and a tyrosine kinase, thereby providing a physical basis for the accessory role of the CD7 molecule in T cell activation.22 In this study the shifts in the mean values of fluorescence distribution histograms were determined rather than measuring FRET efficiency on a cell-by-cell basis. This approach provided enough evidence to support the coimmunoprecipitation data, although the homoassociation of the CD45 molecules could not be addressed because of the inherent limitations of this approach. In another FRET study the CD45 served as a negative control. Intensity based FRET microscopy was used to explore the transmembrane association of leukocyte function associated antigen-1 (LFA-1) with microfilaments in response to TCR ligation.23 Fluorescein conjugated LFA-1 F(ab0 ) fragments served as donors and rhodamine phalloidins as acceptors. Microscopic FRET was quantitated both using imaging and photon counting techniques. When cells were incubated on anti-CD3 coated glass surfaces both fluorescence images and photon count rates were significantly enhanced. This enhancement was not due to a general effect of T cell activation on transmembrane cytoskeletal proximity since there was no FRET observed between CD45 and phalloidin upon CD3 ligation. The authors proposed that the proximity or linkage between LFA-1 and microfilaments may contribute to the formation of stable integrin mediated mechanical linkages between cells.23 An investigation into the role of CD45 isoforms in T cell antigen receptor signal transduction was carried out by transfecting CD45-negative CD4 þ CD8 þ HPB-ALL T cells with the CD45R0, CD45RBC and

22

A. I. Lazarovits, N. Osman, C. E. Le Feuvre, S. C. Ley, and M. J. Crumpton, J. Immunol. 153, 3956 (1994). 23 H. Poo, B. A. Fox, and H. R. Petty, J. Cell. Physiol. 159, 176 (1994).

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CD45RABC isoforms.24 The flow cytometric FRET method was used to determine the association patterns of different CD45 isoforms, although using donor-quenching both accuracy and reproducibility of FRET measurements were compromised due to the low expression level of CD45 molecules. In an attempt to increase the accuracy of flow cytometric FRET measurements, the newly developed method based on the application of Cy3 and Cy5 donor acceptor pair and the cell-by-cell correction of autofluorescence was applied.8 Flow cytometric FRET analysis showed that the CD45R0 isoform, but not the CD45RBC or CD45RABC isoforms, was found as homodimers in these cells, and also preferentially associated with CD4 and CD8 at the cell-surface. These differential associations were correlated with the actions of CD45 in activating the CD4/CD8-associated p56lck tyrosine kinase, thereby regulating the efficiency of TCR signal transduction coupling.24 The mechanism of CD45R0 association with CD4 and CD8 remains unknown. The association could be mediated by an intermediary molecule or more likely by direct interactions between binding motifs in the respective polypeptide back-bones and/or oligosaccharide interactions. It is possible that CD45-CD4/CD8 binding represents the default association state and that this is prevented by the A–C exon products. Similarly CD45R0 homoassociation might represent the default position of CD45 molecules that is also prevented, for example, by the heavy sialylation associated with the A-C exon region.

RPTP

Structural, functional and biochemical studies have suggested that the receptor tyrosine phosphatase  (RPTP) is inhibited by dimerization, but direct evidence that RPTP actually dimerizes in living cells was provided only recently by Tertoolen et al.25 Instead of using fluorescently conjugated mAbs, the authors used fluorescent fusion proteins. In order to assess RPTP homodimerization, FRET was measured between chimeric proteins of cyan (CFP) and yellow (YFP) emitting derivatives of green fluorescent protein, fused to RPTP, using three different techniques: dual wavelength excitation (i.e., intensity based FRET microscopy), spectral imaging and fluorescence lifetime imaging. All three methods suggested that FRET occurred between RPTP-CFP and RPTP-YFP fusion proteins, and thus that RPTP constitutively dimerized in living cells. FRET results were 24

S. Dornan, Z. Sebestye´n, J. Gamble, P. Nagy, A. Bodna´r, L. Alldrige, S. Doe, N. Holmes, 00 L. K. Goff, P. Beverley, J. Szo¨lllosi, and D. R. Alexander, J. Biol. Chem. 277, 1912 (2002). 25 L. G. J. Tertoolen, C. Blanchetot, G. Jiang, J. Overvoorde, T. W. J. Gadella Jr., T. Hunter, and J. den Hertog, BMC Cell Biol. 2, 8 (2001).

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consistent with experiments using chemical cross-linkers. Using deletion mutants, the site of interaction was mapped to the transmembrane domain, and the results showed that the transmembrane domain was sufficient to drive dimerization. These data provide strong support for the model that dimerization is involved in the regulation of RPTP activity.25 GPI-anchored Phosphatase

Indirect evidence suggests that the GPI anchor in GPI-anchored proteins may function to hold the protein close to the plasma membrane. The FRET technique has now been used to test this theory directly for the GPI-anchored ectoenzyme placental alkaline phosphatase (PLAP).26 For this study PLAP was fluorescently labeled at the N-terminus with 7(dimethylamino)coumarin-4-acetic acid succinimidyl ester (DMACA-SE) or Oregon Green 488 succinimidyl ester (OG488-SE), and each was reconstituted by detergent dilution into defined lipid bilayer vesicles containing an increasing mole fraction of a fluorescent lipid probe. The fluorescence of the labeled PLAP donors was quenched in a concentrationdependent manner by the lipid acceptors. The energy transfer data were analyzed using an approach that describes FRET between a uniform distribution of donors and acceptors in an infinite plane, and the distance of closest approach between the protein moiety of PLAP and the lipid-water interfacial region of the bilayer was estimated to be smaller than 10–14 A˚. These results supported the concept that the protein portion of PLAP is located very close to the lipid bilayer, possibly resting on the surface.26 Protein Phosphatase 1

Protein phosphatase 1 (PP1) is expressed in mammalian cells as three closely related isoforms, known as , b/ and 1, all encoded by separate genes. It is not yet known whether these isoforms play distinct roles in vivo. All isoforms have the potential to interact with an inhibitor known as NIPP1 (nuclear inhibitor of PP1) which targets PP1 to interchromatin granule clusters in the nucleoplasm.27 When PP1- and PP1- were tagged with fluorescent probes and coexpressed with NIPP1 they were both retargeted to these structures. FRET analysis revealed that direct interaction of PP1- with NIPP1 occurred predominantly at or near interchromatin granule clusters. This provides an example of the way in which FRET can be used to 26 27

M. T. Lehto and F. J. Sharom, Biochemistry 41, 8368 (2002). L. Trinkle-Mulcahy, J. E. Sleeman, and A. I. Lamond, J. Cell Sci. 114, 4219 (2001).

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generate information about intracellular phosphatase interactions and localization.27 PTP1B

A further useful application of FRET has been to identify a novel trafficking pathway whereby tyrosine phosphorylated versions of the EGF and PDGF receptors internalize and transit to the endoplasmic reticulum where they are dephosphorylated by the ER-associated PTP1B phosphotyrosine phosphatase. To directly measure molecular interactions between PTP1B and receptor tyrosine kinases, FRET was measured between EGFR-GFP, PDGFR-GFP and Cy3-FG6 mAb bound to PTP1B, using a special application of acceptor photobleaching. Fluorescence lifetime of the donor was determined before and after acceptor photobleaching using fluorescence lifetime imaging microscopy (FLIM) on a pixel-by-pixel basis. FLIM is a robust method for determining FRET in cells, because fluorescence lifetimes are independent of fluorophore concentrations and light path length but dependent on excited state reactions such as FRET. The fluorescence lifetimes were determined using the phase modulation approach. No FRET was observed between PTP1B and EGFR or PDGFR in resting cells. However, after stimulation punctate structures with FRET could be detected, indicative of localized interactions. In a complementary experiment the classical acceptor photobleaching approach was applied in a confocal microscope in order to generate high-resolution 3D image of sites of EGFR or PDGFR interactions with PTP1B. (The FLIM approach is still not available in combination with confocal microscopy.) With this combined experimental approach, where the FLIM provided more robust data and the confocal FRET imaging better 3D resolution, it was successfully demonstrated that a high percentage of the activated receptor tyrosine kinases (such as EGFR and PDGFR) become associated with PTP1B in the ER prior to degradation or recycling.28 It is clear that the use of antibodies and fusion proteins in combination with intensity based FRET imaging, spectral imaging and fluorescence lifetime imaging, together offer the opportunity to study the complex behavior of key regulatory proteins in their natural environment within the living cell. It seems very likely that FRET will see increasing application in the investigation of phosphatases within their natural habitats. 28

F. G. Haj, P. J. Verveer, A. Squire, B. G. Neel, and P. I. H. Bastiaens, Science 295, 1708 (2002).

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Acknowledgments This work was supported by the Biotechnology and Biological Sciences Research Council, by the British Council British-Hungarian Academic Research Programs 069 and GB-1/2003, and by the Hungarian Academy of Sciences Grant OTKA 30399/1999.

[17] Receptor Protein-Tyrosine Phosphatase Dimerization By JEROEN DEN HERTOG, THEA VAN DER WIJK, LEON G. J. TERTOOLEN, and CHRISTOPHE BLANCHETOT

Introduction

The receptor protein-tyrosine phosphatases (RPTPs) form a subfamily of the classical protein-tyrosine phosphatase family.1 The extracellular domains of the RPTPs are diverse, varying from very small (e.g., RPTP", 23 amino acids) to very large with multiple Fibronectin type III-like and immunoglobulin domains (e.g., LAR). RPTPs have a single membranespanning domain and most RPTPs contain two cytoplasmic catalytic domains. The N-terminal membrane-proximal domain, RPTP-D1, contains most of the catalytic activity, while the C-terminal membrane-distal PTP domain, RPTP-D2, appears to play a regulatory role. Little is known about how RPTP activity is regulated. It is established that the enzymatic counterparts of the RPTPs, the receptor protein-tyrosine kinases (RPTKs), are regulated by ligand binding to their extracellular domain, e.g., epidermal growth factor (EGF) binding activates intrinsic PTK activity of the EGF receptor (EGFR). Similarly, ligand binding to the ectodomain of RPTPs may be involved in regulation of their activity. By now, several ligand-RPTP pairs have been identified. Both soluble factors were identified as ligands, e.g., Pleiotrophin binding to RPTPb/,2 as well as extracellular matrix components, e.g., Heparan Sulphate Proteoglycan binding to RPTP.3 Whether ligand binding affects intracellular PTP activity remains to be determined definitively. Moreover, RPTPs may be regulated by other means as well, including posttranslational modifications such as phosphorylation and oxidation. 1

N. K. Tonks and B. G. Neel, Curr. Opin. Cell Biol. 13, 182–195 (2001). K. Meng, A. Rodriguez-Pena, T. Dimitrov, W. Chen, M. Yamin, M. Noda, and T. F. Deuel, Proc. Natl. Acad. Sci. U.S.A. 97, 2603–2608 (2000). 3 A. R. Aricescu, I. W. McKinnell, W. Halfter, and A. W. Stoker, Mol. Cell Biol. 22, 1881–1892 (2002). 2

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Acknowledgments This work was supported by the Biotechnology and Biological Sciences Research Council, by the British Council British-Hungarian Academic Research Programs 069 and GB-1/2003, and by the Hungarian Academy of Sciences Grant OTKA 30399/1999.

[18] Receptor Protein-Tyrosine Phosphatase Dimerization By JEROEN DEN HERTOG, THEA VAN DER WIJK, LEON G. J. TERTOOLEN, and CHRISTOPHE BLANCHETOT

Introduction

The receptor protein-tyrosine phosphatases (RPTPs) form a subfamily of the classical protein-tyrosine phosphatase family.1 The extracellular domains of the RPTPs are diverse, varying from very small (e.g., RPTP", 23 amino acids) to very large with multiple Fibronectin type III-like and immunoglobulin domains (e.g., LAR). RPTPs have a single membranespanning domain and most RPTPs contain two cytoplasmic catalytic domains. The N-terminal membrane-proximal domain, RPTP-D1, contains most of the catalytic activity, while the C-terminal membrane-distal PTP domain, RPTP-D2, appears to play a regulatory role. Little is known about how RPTP activity is regulated. It is established that the enzymatic counterparts of the RPTPs, the receptor protein-tyrosine kinases (RPTKs), are regulated by ligand binding to their extracellular domain, e.g., epidermal growth factor (EGF) binding activates intrinsic PTK activity of the EGF receptor (EGFR). Similarly, ligand binding to the ectodomain of RPTPs may be involved in regulation of their activity. By now, several ligand-RPTP pairs have been identified. Both soluble factors were identified as ligands, e.g., Pleiotrophin binding to RPTPb/,2 as well as extracellular matrix components, e.g., Heparan Sulphate Proteoglycan binding to RPTP.3 Whether ligand binding affects intracellular PTP activity remains to be determined definitively. Moreover, RPTPs may be regulated by other means as well, including posttranslational modifications such as phosphorylation and oxidation. 1

N. K. Tonks and B. G. Neel, Curr. Opin. Cell Biol. 13, 182–195 (2001). K. Meng, A. Rodriguez-Pena, T. Dimitrov, W. Chen, M. Yamin, M. Noda, and T. F. Deuel, Proc. Natl. Acad. Sci. U.S.A. 97, 2603–2608 (2000). 3 A. R. Aricescu, I. W. McKinnell, W. Halfter, and A. W. Stoker, Mol. Cell Biol. 22, 1881–1892 (2002). 2

METHODS IN ENZYMOLOGY, VOL. 366

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Dimerization is a well-known regulatory mechanism for single membrane-spanning receptors.4 For instance, ligand-induced RPTK activation involves dimerization.5 The RPTPs may be regulated by dimerization as well. Ligand-induced dimerization of EGFR/CD45, a chimeric protein, consisting of the extracellular domain of the EGFR and the intracellular domain of the RPTP, CD45, leads to functional inactivation of CD45,6 suggesting that dimerization inhibits CD45 activity. The crystal structure of RPTP-D1 provided structural support for dimerization-induced inhibition of RPTP activity.7 A helix-turn-helix wedge-like segment to the N-terminal side of RPTP-D1 that is conserved in RPTPs, but not in cytoplasmic PTPs, interacts with the dyad-related monomer, thus forming a dimer in which both catalytic sites are occluded.7 Mutation of a single residue in the corresponding wedge region of CD45 abolishes dimerization-induced functional inactivation, strongly suggesting that the wedge plays a role in dimerizationinduced inactivation of CD45.8 Forced dimerization of RPTP by introduction of a disulfide bridge in the extracellular domain leads to inactivation of RPTP catalytic activity, which is dependent on an intact wedge.9 However, three other RPTP mutants with a disulfide bond in the extracellular domain dimerize constitutively, but are active like wild type RPTP, demonstrating that dimerization per se does not lead to inactivation. The positions of the disulfide bonds suggest that rotational coupling between the two monomers in a dimer is an important determinant for dimer activity. Perhaps RPTPs dimerize constitutively and their activity is regulated by small changes in rotational coupling as a result of ligand binding or posttranslational modification. In conclusion, evidence is accumulating that RPTPs can be regulated by dimerization. We have addressed the question whether RPTPs dimerize in living cells using four different techniques: (i) chemical cross-linking, (ii) genetic crosslinking by introduction of disulfide bonds, (iii) fluorescence resonance energy transfer (FRET) and (iv) analysis of interactions of RPTP fragments by coimmunoprecipitation. These techniques will be reviewed in this chapter. Chemical Cross-linkers

Chemical cross-linkers have been widely used to investigate protein– protein interactions, including RPTP dimerization. For instance, Takeda 4

A. Weiss and J. Schlessinger, Cell 94, 277–280 (1998). M. A. Lemmon and J. Schlessinger, Trends Biochem. Sci. 19, 459–463 (1994). 6 D. M. Desai, J. Sap, J. Schlessinger, and A. Weiss, Cell 73, 541–554 (1993). 7 A. M. Bilwes, J. den Hertog, T. Hunter, and J. P. Noel, Nature 382, 555–559 (1996). 8 R. Majeti, A. M. Bilwes, J. P. Noel, T. Hunter, and A. Weiss, Science 279, 88–91 (1998). 9 G. Jiang, J. den Hertog, J. Su, J. Noel, J. Sap, and T. Hunter, Nature 401, 606–610 (1999). 5

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et al.10 used the thiol-cleavable and homobifunctional chemical cross-linker, dithiobis succinimidyl propionate (DSP) to show dimerization of CD45. Noncell permeable cross-linkers may be used to cross-link surface molecules on living cells. We have used the homobifunctional noncell permeable crosslinker, bis[sulfosuccinimidyl]-suberate (BS3), on living cells to demonstrate cross-linking of RPTP.11,12 A panel of deletion mutants and BS3-mediated cross-linking were used to map the site(s) of interaction in RPTP. The cells were biotinylated prior to cross-linking to ensure that only surface expressed RPTP was detected.11 Finally, we used BS3-mediated cross-linking with different experimental conditions to assess the effects of oxidative stress on RPTP dimerization.13 Xu and Weiss14 used a similar membrane-insoluble cross-linker, ethylene glycol bis[sulfosuccinimidylsuccinate] (sulfo-EGS), to show cross-linking of CD45. Taken together, chemical cross-linking using membrane-insoluble cross-linkers is a versatile method to show dimerization of RPTPs.

Cross-linking Using bis[sulfosuccinimidyl]-suberate (BS3)

Routinely, we use transiently or stably transfected cells overexpressing epitope tagged RPTP(s) of interest for our analyses of dimerization, facilitating detection of the RPTP(s) by immunoblotting and/or immunoprecipitation. We use 293 human embryonic kidney, mouse SK-N-MC neuroepithelioma and mouse embryo fibroblast cells for these experiments. Using standard transfection procedures, we obtain 2 to 5-fold overexpression of RPTP in stably transfected cells and 5 to 10-fold in transiently transfected cells. Immediately prior to harvesting, the cells are washed (2
) in ice-cold phosphate buffered saline (PBS) and incubated with freshly prepared BS3 (1 mg/ml) in PBS for 1 hr on ice. Subsequently, the cells are washed (2
) with PBS and lysed in Tris-buffered cell lysis buffer, quenching excess BS3 for 20 min on ice (T-CLB: 50 mM Tris, pH 7.5, 150 mM NaCl, 1 mM MgCl2, 1 mM EGTA, 1% Triton X-100, 10% glycerol and protease inhibitors: 10 U/ml aprotinin, 1 M PMSF). Cells are scraped, lysates collected and cell debris removed by centrifugation at 4 C. Aliquots of the cell lysates are loaded onto sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) gels directly or RPTP is 10

A. Takeda, J. J. Wu, and A. L. Maizel, J. Biol. Chem. 267, 16651–16659 (1992). G. Jiang, J. den Hertog, and T. Hunter, Mol. Cell Biol. 20, 5917–5929 (2000). 12 L. G. Tertoolen, C. Blanchetot, G. Jiang, J. Overvoorde, T. W. J. Gadella, T. Hunter, and J. den Hertog, BMC Cell Biol. 2, 8 (2001). 13 C. Blanchetot, L. G. Tertoolen, and J. den Hertog, EMBO J. 21, 493–503 (2002). 14 Z. Xu and A. Weiss, Nat. Immunol. 3, 764–771 (2002). 11

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immunoprecipitated using anti-hemagglutinin (HA) epitope tag MAb 12CA5, directed against the HA-tag in the extracellular domain of RPTP, immediately C-terminal to the signal sequence of RPTP. After washing 4
with HNTG (50 mM HEPES, pH 7.4, 150 mM NaCl, 0.1% Triton X100, 10% glycerol), reducing Laemmli sample buffer is added to the immunoprecipitates, the samples are heated at 95 C for 5 min and loaded onto an SDS-PAGE gel. After electrophoresis, the material on the gel is transferred to polyvinyldifluoride (PVDF) membranes by semidry blotting and probed with polyclonal anti-RPTP antibodies or MAb 12CA5. The blots are developed by enhanced chemiluminescence (ECL) according to standard protocols. A typical example of a cross-linking experiment is depicted in Fig. 1. Genetic Cross-linking

Introduction of a cysteine residue in the extracellular juxtamembrane region of a transmembrane protein may lead to the formation of a disulfide bond if two molecules are close enough together. In some respects, this is similar to the action of chemical cross-linkers and therefore introduction of a disulfide bond may be viewed as genetic cross-linking. Sorokin et al.15 found that introduction of a cysteine residue in the extracellular domain of the EGFR by itself is not sufficient to induce dimerization. However, EGF-treatment leads to disulfide bond formation of mutant EGFR, which persists after removal of EGF. We demonstrated that introduction of Cys residues in the extracellular domain of RPTP induces constitutive dimerization, as assessed by SDS-PAGE on nonreducing gels. We engineered four mutants with Cys-residues in the juxtamembrane domain of RPTP, just outside the plasma membrane at positions 135, 137, 139, and 141 (the first residue of the transmembrane domain being Ile143), ensuring that the four mutants cover at least one complete -helical turn.9 All four mutants dimerize constitutively to a similar extent (cf. Fig. 2). Similarly, introduction of Cys residues in the extracellular domain of three distinct splice isoforms of CD45 leads to constitutive dimerization as well.14 Disulfide bond formation between two mutant proteins with a Cys residue in the ectodomain, close to the membrane, does not formally prove that the wild type proteins dimerize in living cells. However, it does indicate that these two proteins have some affinity for each other, in that they have been very close to allow formation of the disulfide bond. The EGFR mutant suggests that introduction of a Cys residue by itself is not sufficient for 15

A. Sorokin, M. A. Lemmon, A. Ullrich, and J. Schlessinger, J. Biol. Chem. 269, 9752–9759 (1994).

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FIG. 1. Chemical cross-linking of RPTP dimers. Human embryonic kidney 293 cells were transfected with vector (), or with an SV40 promoter driven expression vector for hemagglutinin-tagged RPTP-
250-YFP (RPTP), a chimeric construct encompassing the extracellular domain, transmembrane domain and a short segment of the intracellular domain of RPTP (residues 1–250), fused to YFP. The HA-epitope tag is inserted in the extracellular domain, immediately following the signal sequence. The chemical cross-linker BS3 was used to cross-link cell surface expressed proteins as described in the text. Subsequently, the cells were lysed and aliquots of the cell lysate were run on a 7.5% SDS-PAGE gel. The material on the gel was transferred to polyvinylidene fluoride (PVDF) membrane by semidry blotting. The blots were probed with anti-HA-tag MAb 12CA5 and developed by enhanced chemiluminescence using standard protocols. The molecular weights of marker proteins that were coelectrophoresed with the samples are indicated on the left in kDa. The position of HA-RPTP-
250-YFP monomer and dimer is indicated on the right. Nonspecific background bands in the lysate are also indicated on the right (NS).

disulfide bond formation, since EGF treatment is required for dimerization and disulfide bond formation. Detection of Enforced Dimers

Whether there is an optimal position for engineered disulfide bonds in transmembrane proteins remains to be determined. In the EGFR, RPTP and CD45 disulfide bonds were successfully introduced in the extracellular juxtamembrane domain—within the first 10 residues from the

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FIG. 2. Genetic cross-linking of RPTP. Phe135 and Pro137 in the extracellular juxtamembrane region of RPTP, were replaced by Cys by site directed mutagenesis, rendering RPTP-F135C and RPTP-P137C, respectively. SV40 promoter driven expression vectors for HA-tagged wild type RPTP (WT), RPTP-F135C (F135C) or RPTP-P137C (P137C) were transfected into HEK 293 cells. The cells were lysed in the presence of 20 mM iodoacetic acid (IAA) to protect the disulfide bonds. Whole cell lysates (WCL) were resuspended in Laemmli sample buffer without b-mercaptoethanol (nonreducing conditions), or with b-mercaptoethanol (2% v/v), heated for 5 min at 95 C and loaded onto 5% SDS-PAGE gels. Separate gels were used for the reducing and nonreducing conditions to avoid inadvertent reduction of disulfide bonds. Following electrophoresis, the material on the gels was transferred to PVDF membranes, and the blots were probed with anti-RPTP polyclonal antibody (5478) and developed using ECL. The blots depicted here show extensive dimerization of RPTP-F135C and RPTP-P137C under nonreducing conditions, but not under reducing conditions. Wild-type RPTP dimers are not detectable under these conditions. The position of RPTP dimers, monomers and of a nonspecific background band (NS) are indicated on the right. Molecular weights of marker proteins that were coelectrophoresed with the samples are indicated in kDa on the left.

transmembrane domain—either by point mutation or by insertion of a short linker encoding a Cys. Detection of dimers after disulfide bond formation is done by nonreducing SDS-PAGE. The cells are transfected with mutants, containing Cys residues in the extracellular domain, and the cells are harvested by lysis

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in CLB (50 mM HEPES, pH 7.4, 150 mM NaCl, 1 mM MgCl2, 1 mM EGTA, 1% Triton X-100, 10% glycerol and protease inhibitors: 10 U/ml aprotinin, 1 M PMSF) containing 20 mM iodoacetic acid (IAA) to protect the disulfide bond from reduction (20 min on ice). Monomers and dimers are detected by immunoblotting of the whole cell lysate, or the protein of interest is first immunoprecipitated. To avoid interfering bands of the intact antibody in the molecular weight range of monomeric or dimeric protein of interest, the antibodies are cross-linked to Protein-A Sepharose beads. We routinely cross-link antibodies to Protein A-Sepharose beads with dimethylpimelimidate (DMP), using an improved version of a protocol for cross-linking of Glutathione-S-Transferase fusion proteins to Glutathione beads:16 the antibody is allowed to bind to Protein-A Sepharose beads for 1 hr at room temperature; wash 3
0.2 M Na2B4O7 buffer (pH 9.0); cross-link with 20 mM DMP (fresh) in Na2B4O7 buffer (30 min, room temperature); wash 3
0.2 M ethanolamine (pH 8.0); incubate 1 hr room temperature in 0.2 M ethanolamine (to quench excess DMP); wash 3
Tris Buffered Saline (TBS; 50 mM Tris, pH 7.5; 150 mM NaCl). Cross-linked antibodies may be stored for later use at 4 C in TBS. The immunoprecipitates are washed 4
in HNTG and the beads are resuspended in an equal volume of 2
Laemmli sample buffer without b-mercaptoethanol (nonreducing conditions), or with b-mercaptoethanol (2% v/v; reducing conditions). The samples are heated at 95 C for 5 min and loaded onto separate 5% SDS-PAGE gels to avoid reduction of the disulfide bond in the nonreduced samples. Similarly, whole cell lysates are mixed with nonreducing or reducing sample buffer and loaded on separate SDS-PAGE gels. After electrophoresis, the gels are blotted by semidry blotting and the blots are probed with antibodies and developed with ECL according to standard procedures.

Fluorescence Resonance Energy Transfer

FRET is considered to be a spectroscopic ruler17 that may be used to assess distances at the molecular level in intact living cells and is used to analyze protein–protein interactions in vivo. FRET is a direct radiation-less transfer of energy from the excited donor fluorophore to the acceptor fluorophore, which occurs if the two fluorophores are in very near vicinity of

16

A. Koff, A. Giordano, D. Desai, K. Yamashita, J. W. Harper, S. Elledge, T. Nishimoto, D. O. Morgan, B. R. Franza, and J. M. Roberts, Science 257, 1689–1694 (1992). 17 L. Stryer, Annu. Rev. Biochem. 47, 819–846 (1978).

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each other, usually less than 60 A˚.18–20 A major advantage of this technique is that it can be used to analyze interactions between genetically encoded fluorophores, e.g., derivatives of Green Fluorescent Protein (GFP).21,22 For instance, if the proteins of interest are fused to Cyan Fluorescent Protein (CFP, donor fluorophore) on the one hand and Yellow Fluorescent Protein (YFP, acceptor fluorophore) on the other, analysis of FRET between the two GFP derivatives (GFP-FRET) can be used to assess interactions between the two proteins. Dimerization of G protein coupled receptors has been assessed using FRET23 and we have analyzed RPTP dimerization in living cells by assessing FRET between fusion proteins of RPTP and the CFP-YFP fluorophore pair.12 FRET has also been used to assess dimerization of CD45, using anti-CD45 specific antibodies, coupled to fluorophores.24 GFP-FRET may be used to investigate conformational changes in fragments of proteins as well. CFP is fused to one end of the protein fragment, YFP to the other and FRET measurements are done under different experimental conditions. Initially, GFP-FRET was used to assay changes in cytosolic Ca2 þ -concentrations using ‘‘cameleons,’’ chimeric proteins consisting of the Ca2 þ -binding domain of Calmodulin, flanked by CFP and YFP. Ca2 þ binding to cameleon leads to a conformational change, bringing the two fluorophores closer together, leading to an increase in FRET. Therefore, cameleons are genetically encoded, spectrometric Ca2 þ sensors.25,26 Since then, many FRET-based ‘‘sensor’’ proteins have been designed, including indicators of protein (tyrosine) kinase activity and protease activity. We have used FRET to measure a conformational change in RPTP-D2 in response to oxidative stress.13 CFP and YFP were fused to the N- and C-terminus of RPTP-D2, respectively, resulting in high FRET levels (cf. Fig. 3C), which is consistent with the crystal structure of RPTP-D2, in which the N- and C-terminus are very close together. H2O2 treatment led to a dramatic reduction in FRET, as a result of a conformational change. The catalytic site Cys in RPTP-D2 is 18

R. M. Clegg, in ‘‘Fluorescence Imaging Spectroscopy and Microscopy’’ (X. F. Wang and B. Herman, eds.), pp. 179–252. John Wiley & Sons Inc., Oxford, 1966. 19 P. R. Selvin, Meth. Enzymol. 246, 300–334 (1995). 20 P. Wu and L. Brand, Anal. Biochem. 218, 1–13 (1994). 21 T. W. J. Gadella, G. N. van der Krogt, and T. Bisseling, Trends. Plant Sci. 4, 287–291 (1999). 22 B. A. Pollok and R. Heim, Trends Cell Biol. 9, 57–60 (1999). 23 M. C. Overton and K. J. Blumer, Curr. Biol. 10, 341–344 (2000). 24 S. Dornan, Z. Sebestyen, J. Gamble, P. Nagy, A. Bodnar, L. Alldridge, S. Doe, N. Holmes, L. K. Goff, P. Beverley, J. Szollosi, and D. R. Alexander, J. Biol. Chem. 277, 1912–1918 (2002). 25 A. Miyawaki, J. Llopis, R. Heim, J. M. McCaffery, J. A. Adams, M. Ikura, and R. Y. Tsien, Nature 388, 882–887 (1997). 26 R. Y. Tsien, Annu. Rev. Biochem. 67, 509–544 (1998).

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FIG. 3. RPTP dimerization demonstrated by FRET. (A) Emission spectra of CFP and YFP. Full length RPTP, fused to CFP (RPTP-CFP) or RPTP, fused to YFP (RPTP-YFP) were transfected into 293 HEK cells. Single cell measurements were done to establish the emission spectra of the two fluorophores in our system. CFP was excited at 430 nm (slit width 8 nm) and YFP at 490 nm (slit width 8 nm). Emission spectra were recorded as described in detail in the text. (B) Equal amounts of RPTP-CFP and RPTP-YFP were cotransfected into 293 HEK cells and emission spectra of single cells (excitation at 430 nm) were recorded (RPTP). Similarly, equal amounts of EGFR-CFP and EGFR-YFP, with CFP or YFP fused to Ile923 to the C-terminal side of the kinase domain in the EGFR were transfected into 293 HEK cells and emission spectra were recorded (excitation at 430 nm). Note the shift in the optimum of the RPTP spectrum to higher wavelengths, reflecting higher sensitized emission (FRET) in RPTP than in the EGFR. In addition, note that both emission spectra are shifted toward higher wavelengths as compared to the CFP spectrum in (A), suggesting FRET occurs in both EGFR and RPTP. (C) H2O2-induced conformational change in RPTP-D2. A fusion construct of the membrane-distal PTP domain of RPTP, RPTP-D2, flanked by CFP to the N-terminal side and YFP to the C-terminal side was transfected into 293 HEK cells. The emission spectrum of resting cells (rest) was recorded (excitation at 430 nm), showing very high levels of FRET, which is consistent with the N- and C-terminus of RPTP-D2 being close together, as seen in the crystal structure. H2O2-treatment (1 mM, 10 min) led to a dramatic reduction in FRET (H2O2). The sensitized emission peak (maximum of 535 nm) is much reduced and at the same time the CFP peak is increased, which is expected since less energy is transferred to YFP by photon-less transfer and more is directly emitted as CFP emission.

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essential for the conformational change, since mutation of this site completely abolished the H2O2-induced reduction in FRET.13 Taken together, FRET is a usefull technique that can be used to investigate constitutive as well as inducible inter- and intramolecular protein–protein interactions in vivo.

Detection of FRET

There are several ways to detect FRET. These are all based on excitation of the donor fluorophore and detection of the acceptor fluorophore. Fluorescence lifetime imaging (FLIM) uses the difference in fluorescence life time to discriminate between the donor and acceptor fluorophore. FLIM is very reliable and accurate for detection of FRET but it is not as sensitive as other methods and it requires highly sophisticated equipment for the analysis. A relatively simple method to detect FRET is to make use of the differences in spectral properties of the two fluorophores. Excitation of the donor fluorophore will lead to excitation of the acceptor fluorophore by FRET if the two fluorophores are close enough together (< 60 A˚). Therefore, spectrophotometric analysis of sensitized emission of the acceptor upon excitation of the donor is a direct measure for FRET. Ideally, the spectral properties of a donor– acceptor fluorophore pair are such that there is no overlap between the spectra of these fluorophores. However, usually there is overlap between the spectra, leading to inadvertent direct excitation of the acceptor, or to direct detection of the donor, leading to an overestimation of FRET. Corrections can be made for spectral overlap. However, if the relative concentrations of the two fluorophores are not known, which is often the case if the two fluorophores are not in the same molecule, major errors may be introduced. In fact, if the acceptor fluorophore expression is high compared to the donor fluorophore, signals from direct acceptor excitation may be mistaken for FRET.12 We routinely determine emission spectra rather than emission at single wavelengths, using narrow bandpass filters. The spectra show a shift towards the acceptor emission optimum, or even two peaks as a result of sensitized emission (FRET) as compared to the CFP spectrum (Fig. 3). If one of the fluorophores is not expressed properly, the emission spectra loose their characteristic shape, and this would go unnoticed if only emission intensities at 480 nm (CFP) and 535 nm (YFP) would be detected. Nevertheless, determining the ratio between emission intensities at two wavelengths is very useful when changes in FRET over time in response to stimuli are investigated, since the relative amounts of the fluorophores are constant and do not affect the change

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in FRET. Moreover, ratiometric FRET analysis allows imaging of FRET in living cells. FRET Analysis Hardware

The hardware that is required for spectral measurements of FRET consists of a powerful light source and a filter set that allows discrimination between the donor and acceptor fluorophores. FRET measurements can be done in a bulk of cells, using a spectrophotometer, but in our experience, the detection levels in transiently transfected cells are too low. In addition, FRET measurements can be done in cell lysates, using a spectrophotometer, or even using purified proteins. However, the strength of the FRET technique lies in analysis of protein–protein interactions in living cells. To this end, a FACS (Fluorescence Assisted Cell Sorter) has been used successfully.23,24 The most powerful set-up is a fluorescence microscope, allowing single cell spectral measurements (SCSM), detection of fluorescence and FRET in specific subcellular locations and single cell spectral imaging of FRET. The fluorescence microscope should be equipped with filter sets that allow excitation of only the donor or acceptor. Moreover, the filters should discriminate between emission of the donor and acceptor. For single cell spectral imaging a spectrophotometer is required for spectral imaging on the detector side. We routinely use a Leitz orthoplan upright microscope (Leitz GMBH, Wetzlar, Germany) with a Xenon light source (450 W). We use four different configurations of the microscope for four different types of measurements: For static dual wavelength excitation measurements, two filter sets (Ploemopak) are used, the ‘‘CFP’’ filter set (filter #1) with an RKP455 dichroic mirror and a 490 nm long-pass emission filter, and the ‘‘FRET’’ filter (filter #2), equipped with a dichroic mirror RKP510 (reflection shortpass filter) and a BP530–560 (band-pass) emission filter (Leitz GMBH, Wetzlar, Germany). The fluorescence intensity is quantified with a Photon Counting Tube (type 9862, EMI Limited, Middlesex, England). The fluorescence intensities (obtained after excitation at 440 nm or 490 nm) are corrected for differences in excitation light intensities, using the reference photomultiplier. Fluorescence intensities are recorded from single living cells and corrected for background, using adjacent nontransfected cells. For real time dual excitation measurements, the excitation wavelengths are rapidly alternated between 430(8) and 490(8) nm. A RKP 515 nm dichroic mirror is used with an emission barrier filter 535(30) nm. Fluorescence is quantified via a photomultiplier. An image intensifier coupled to a progressive scan CCD camera synchronized with the chopper module is used to collect time series of dual wavelength excitation image pairs.

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For real time dual emission measurements, two filters 480(20) nm and 535(30) nm mounted in a fast rotating filter wheel are used in combination with a JP4 dichroic mirror (Chroma Technology, Brattleboro, USA), excitation at 430(8) nm. An image intensifier coupled to a progressive scan CCD camera synchronized with the filter wheel is used to collect time series of dual wavelength image pairs. For single cell spectral analysis, a filter cube fitted with a 455 nm dichroic mirror at 430(8) nm excitation is used. Emission spectra are recorded from 450 to 600 nm (8 nm bandwidth). The YFP spectra are measured with a dichroic mirror >510 nm at 490(8) nm excitation from 510 to 600 nm (8 nm bandwidth), with an integration time of 0.5 s/nm, 8 nm bandwidth. CFP and YFP Fusion Proteins

For our FRET analyses, we have used the CFP-YFP donor–acceptor fluorophore pair. The mutations that were introduced in GFP to generate CFP and YFP are listed in Table I. The spectral properties of these two GFP derivatives make them a good pair for FRET analysis (Table I, Fig. 3A). Currently, we use an improved version of YFP, YFP2.1 from cameleon2.1,27 with two extra point mutations (Table I) that is relatively

GFP DERIVATIVES Protein

Mutationsa

CFP

YFP2.1

TABLE I SPECTRAL PROPERTIES

AND THEIR

Excitation maximum

Emission maximum

K26R F64L S65T Y66W N146I M153T V163A N164H

450 nm

480 nm

S65G V68Lb Q69Kb S72A T203Y

514 nm

527 nm

a The mutations that were introduced in GFP to generate CFP and YFP2.1 are listed here. b Extra mutations in YFP2.1 as compared to original YFP.

27

A. Miyawaki, O. Griesbeck, R. Heim, and R. Y. Tsien, Proc. Natl. Acad. Sci. U.S.A. 96, 2135–2140 (1999).

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insensitive to pH changes at physiological pH and that is insensitive to oxidative stress, unlike the original YFP. For the analysis of dimerization, we fused CFP or YFP to the C-terminus of RPTP. Moreover, we mapped the sites that were important for dimerization using deletion mutants, each with CFP or YFP fused to the C-terminus. As a control, we fused CFP or YFP to the C-terminal region of the EGFR. We detected very low levels of FRET in cells coexpressing EGFR-CFP and EGFR-YFP12 (Fig. 3B). In contrast, most of the RPTP constructs we tested showed high levels of sensitized emission (FRET) at 535 nm, the YFP emission optimum, indicating that RPTP and its deletion mutants dimerized. In addition, these results suggest that the C-termini of these constructs are close enough together to allow FRET to occur. The topology of RPTP may facilitate detection of FRET in these transmembrane proteins, since the transmembrane domain reduces the level of freedom of movement to two dimensions. Analysis of FRET between cytoplasmic proteins may require the generation of multiple fusion proteins with CFP/YFP at different sites in the protein.

Single Cell Spectral Microscopy-FRET Analysis

For the SCSM-FRET experiments, cells grown on glass coverslips are transiently transfected with the appropriate CFP- and YFP-fusion constructs. Two days after transfection, the cells are mounted in the sample chamber of the microscope and incubated at 22 C in a HEPES buffered saline buffer (10 mM HEPES, 140 mM NaCl, 5 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM glucose, 0.1% BSA, pH 7.5). Fluorescence intensities are recorded from single living cells and corrected for background, using adjacent nontransfected cells. Routinely, we determine the CFP and FRET spectra upon excitation at 430 (8) nm, and the YFP spectra at 490(8) nm. The spectral data are recorded with an integration time of 0.5 s/nm, slit width 8 nm.

Coimmunoprecipitation

Coimmunoprecipitation of proteins from cell lysates is generally seen as strong evidence for specific in vivo protein–protein interactions. We have used coimmunoprecipitation experiments to investigate dimerization of full length RPTPs.13,28 Moreover, we have used coimmunoprecipitation experiments to map the regions in RPTPs that are involved in 28

C. Blanchetot, L. G. Tertoolen, J. Overvoorde, and J. den Hertog, J. Biol. Chem. 277, 47263– 47269 (2003).

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dimerization.13,29 We and others found that RPTP-D2s bind to RPTP-D1s in an intra- and intermolecular fashion. RPTP-D2 binds to the juxtamembrane region of RPTP-D1.30 RPTP binds to various RPTPD2s, suggesting that cross-talk between RPTPs may be a shared mechanism of regulation.29 Inter- and intramolecular interactions between purified CD45-D1 and CD45-D2 indicate that these interactions are direct.31 Recently, the juxtamembrane region of RPTP was shown to bind in an intramolecular fashion with both PTP domains of RPTP.32,33 The function of RPTP-D2 binding to RPTP-D1 remains to be determined definitively. RPTP-D2 binding to RPTP-D1 leads to inactivation of RPTP-D1 catalytic activity,30 presumably by blocking the catalytic site. In contrast, CD45-D2 binding to CD45-D1 increases CD45-D1 activity, which may be mediated by CD45-D2 induced disruption of CD45-D1/CD45-D1 homodimers.31 Interestingly, interactions between RPTP domains may be regulated by external stimuli. We found recently that RPTP-D2 binding to RPTP is regulated by oxidative stress. The conformation of RPTP-D2 changes from a ‘‘closed’’ to an ‘‘open’’ conformation in response to oxidative stress, as demonstrated by intramolecular FRET (Fig. 3C),13 leading to intermolecular binding of RPTP-D2 to other RPTP domains. Similar experiments with LAR-D2 show that oxidative stress induces a conformational change in LAR-D2 as well. Moreover, heterodimerization of full length RPTP with full length LAR is induced by oxidative stress as demonstrated by coimmunoprecipitation.28

Coimmunoprecipitation Experiments

Cells were lysed after cotransfection of the appropriate constructs in CLB for 20 min on ice. Cells were scraped, lysates were collected and cell debris was removed by centrifugation at 4 C. The lysates were incubated with antibody and Protein A-sepharose beads (2 hr, 4 C). The beads were washed 4
with HNTG buffer. Finally, the beads were resuspended in Laemmli sample buffer, heated at 95 C and run on SDS-PAGE gels. Routinely, whole cell lysates were run in parallel to monitor expression of the proteins. 29

C. Blanchetot and J. den Hertog, J. Biol. Chem. 275, 12446–12452 (2000). M. J. Wallace, C. Fladd, J. Batt, and D. Rotin, Mol. Cell Biol. 18, 2608–2616 (1998). 31 J. Felberg and P. Johnson, J. Biol. Chem. 273, 17839–17845 (1998). 32 A. R. Aricescu, T. A. Fulga, V. Cismasiu, R. S. Goody, and S. E. Szedlacsek, Biochem. Biophys. Res. Commun. 280, 319–327 (2001). 33 E. Feiken, I. van Etten, M. F. Gebbink, W. H. Moolenaar, and G. C. Zondag, J. Biol. Chem. 275, 15350–15356 (2000). 30

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Estimating Relative Binding Affinities

We used coimmunoprecipitation experiments to investigate binding between a large panel of RPTP-D1s and RPTP-D2s. To facilitate comparison between different RPTP-D1s and RPTP-D2s, constructs were generated with different epitope tags, the HA-tag or the Myc-tag, fused to the N-terminus of the RPTP-D1s and RPTP-D2s. Routinely, HA-tagged proteins were immunoprecipitated from transfected HEK 293 cells and coimmunoprecipitated Myc-tagged proteins were detected by immunoblotting. Parallel coimmunoprecipitations from cells cotransfected with a certain HA-tagged RPTP-D1 and a panel of distinct RPTP-D2s allowed assessment of relative binding affinities. Ideally, the amount of immunoprecipitated HA-RPTP-D1 was equal in all samples, as well as the amount of MT-RPTP-D2 in the cell lysate. To allow corrections for differences in expression levels, expression of the different constructs was monitored by immunoblotting. The relative amounts of coimmunoprecipitating MT-RPTP-D2 were an indication for the binding affinity. If the molecular weight of two RPTP-D2s is different, the two MT-RPTP-D2s may be cotransfected with a given HA-RPTP-D1 to determine the relative amount of each coimmunoprecipitating MT-RPTP-D2 from the same lysate. The example in Fig. 4 clearly shows that more RPTPD2 bound to RPTP-
D2 than RPTP-D2, even though equal amounts of the RPTP-D2s were expressed in these cells. From this experiment, it can be deduced that the binding affinity of RPTP-
D2 for RPTP-D2 is higher than for RPTP-D2.

Concluding Remarks

In this chapter we described four different techniques that we have used to analyze RPTP dimerization in living cells. The use of FRET to analyze RPTP dimerization is appealing because it is a noninvasive technique that can be done on living cells and it may eventually be used to detect changes in dimerization in real time in response to stimuli. However, low levels of FRET may not be detected and subtle changes in FRET may also be undetectable. Chemical cross-linking is fundamentally different from FRET analysis. FRET is only dependent on the distance between the two fluorophores. BS3-mediated chemical cross-linking is a two-step chemical reaction, leading to a covalent bond. Therefore, chemical BS3-mediated cross-linking will never result in 100% cross-linked proteins. The requirements for detection of dimerization are more stringent for crosslinkers than for FRET, since the linker between the two reactive groups in BS3 is only  12 A˚, while FRET allows distances up to  60 A˚ between the two fluorophores. Yet, chemical cross-linkers, but not FRET, may allow

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FIG. 4. Assessment of relative binding affinities by coimmunoprecipitation. HEK 293 cells were cotransfected with SV40 promoter driven expression vectors for HA-tagged RPTP lacking D2 (HA-RPTP-
D2) and varying amounts of CMV promoter driven expression vectors for Myc-tagged RPTP-D2 and RPTP-D2, as indicated. The cells were lysed and HARPTP-
D2 was immunoprecipitated using anti-HA MAb 12CA5. After extensive washing, the immunoprecipitates were resuspended in Laemmli sample buffer, heated at 95 C for 5 min and loaded onto a SDS-PAGE gel. After electrophoresis the material on the gel was transferred to PVDF membrane by semidry blotting. The bottom of the blot was probed with anti-Myc MAb 9E10, allowing detection of coimmunoprecipitating MT-RPTP-D2 and MT-RPTP-D2 (top panel). The top of the blot was probed with anti-HA MAb 12CA5 to monitor HA-RPTP
D2 levels (middle panel). In parallel, whole cell lysates (WCL) were run on a gel, blotted and probed with anti-Myc MAb 9E10 to monitor MT-RPTP-D2 and MT-RPTP-D2 expression levels (bottom panel). Note that much more RPTP-D2 than RPTP-D2 coimmunoprecipitated with RPTP-
D2, indicating that the binding affinity is much higher for RPTP-D2.

detection of subtle changes in dimerization. For instance, changes in RPTP dimerization as a result of mutations in the wedge of RPTP were detected by chemical cross-linking, but not FRET.11,12 Genetic cross-linking by introduction of disulfide bonds in the extracellular domain of transmembrane proteins is a useful tool that may indicate that proteins homodimerize. Perhaps more importantly, these enforced dimers may be used as positive controls for chemical cross-linking, FRET experiments and/or functional assays. Finally, coimmunoprecipitation is widely accepted evidence that

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proteins interact with each other. However, the affinity of the interaction has to be relatively high to allow detection of coimmunoprecipitating proteins. For instance, while both cross-linking and FRET readily allow detection of full length RPTP dimers, coimmunoprecipitation of full length RPTP was rarely observed under normal conditions.13 Taken together, our results suggest it is advisable to use multiple approaches to assay dimerization of transmembrane proteins, like RPTPs.

Section III Inhibition, Stimulation, and Modulation of Activity

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[19] Phosphoprotein Inhibitors of Protein Phosphatase-1 By MASUMI ETO, CRAIG LEACH, NIKOLAOS A. TOUNTAS, and DAVID L. BRAUTIGAN Phosphatase Inhibitory Phosphoproteins

Over 25 years ago, a search for proteins that could regulate phosphorylase phosphatase yielded heat-stable proteins that were called inhibitor-1 (I-1) and inhibitor-2 (I-2).1 I-1 potency as an inhibitor increased several hundred-fold after phosphorylation of Thr35 by cAMP-dependent protein kinase (PKA),2 whereas inhibitor-2 was effective without prior phosphorylation. Inhibitor-1 became the prototype for other phosphoprotein inhibitors of PP1. Subsequently these inhibitor proteins were used to distinguish protein phosphatases into type-1 (PP1, sensitive to inhibitors) from type-2 (insensitive).3 Over the years studies revealed that I-1 is involved in regulation of the phosphorylation of cAMP-response element binding factor (CREB), calmodulin-dependent kinase-II, and GluR1 subunit of AMPA receptor.4,5 An analogue of I-1 was discovered in brain and called DARPP-32, for dopamine and cAMP regulated phosphoprotein of 32 kDa.6 DARPP-32 inhibition of PP1 also requires phosphorylation of a Thr residue by PKA. DARPP-32 plays an essential role in the signaling pathway for dopamine in brain, highlighted in the announcement of the 2000 Nobel Prize.7 In the last decade several new PP1 inhibitor phosphoproteins, such as NIPP-1, inhibitor-4, CPI-17, PHI-1 and KEPI have been discovered in mammalian cells (Table I). NIPP-1 was discovered as a heat-stable inhibitor protein that proved to be a fragment of a larger protein with multiple domains.8,9 NIPP-1 localizes in nuclei and functions in RNA processing. NIPP-1 is phosphorylated by PKA or casein kinase II (CK-II) but this 1

F. L. Huang and W. H. Glinsmann, Eur. J. Biochem. 70, 419–426 (1976). G. A. Nimmo and P. Cohen, Eur. J. Biochem. 87, 353–365 (1978). 3 T. S. Ingebritsen and P. Cohen, Eur. J. Biochem. 132, 255–261 (1983). 4 A. S. Alberts, M. Montminy, S. Shenolikar, and J. R. Feramisco, Mol. Cell. Biol. 14, 4398–4407 (1994). 5 D. Genoux, U. Haditsch, M. Knobloch, A. Michalon, D. Storm, and I. M. Mansuy, Nature 418, 970–975 (2002). 6 H. C. Hemmings, P. Greengard, H. Y. Lim, and P. Cohen, Nature 310, 503–505 (1984). 7 P. Greengard, P. B. Allen, and A. C. Nairn, Neuron 23, 435–447 (1999). 8 M. Beullens, A. Van Eynde, W. Stalmans, and M. Bollen, J. Biol. Chem. 267, 16538–16544 (1992). 9 M. Bollen, Trends Biochem. Sci. 26, 426–431 (2001). 2

METHODS IN ENZYMOLOGY, VOL. 366

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PHOSPHORYLATION Inhibitor Inhibitor-1 (I-1) Inhibitor-2 (I-2)

DARPP32 NIPP-1

CPI-17

PHI-1 Inhibitor-4 KEPI

[19]

TABLE I OF PP1 INHIBITOR PROTEINS

Phosphorylation site

Kinase

Function

Thr35 Thr72

PKA GSK3, cdc2, MAPK

Ser86, 120, 121

CK-II

Thr34 Thr75 Ser199

PKA Cdk5 PKA

Ser204

CK-II

Thr38

PKC, ROCK, MYPT1-K, ILK PKC, MYPT1-K PKC, ROCK, ILK PKC, ROCK

Inhibition of PP1 Activation of PP1C–I-2 complex Recognition of Thr72 by GSK3 Inhibition of PP1 Inhibition of PKA Activation of PP1C–NIPP-1 complex Activation of PP1C–NIPP-1 complex Inhibition of myosin phosphatase NDa Inhibition of PP1 ND

Ser12 Thr57 Serb ND Thr73 Serb

PKC, ILK PKC

Inhibition of PP1 ND

a

ND, not determined. Phosphorylation site(s) are not identified.

b

reduces, not enhances, the inhibitory potency towards PP1C.9 Inhibitor-4 was identified as an I-2 analog by searching sequence databases.10 Sequences around two phosphorylation sites of I-2 (Thr72, Ser120/121) are conserved in inhibitor-4, but phosphorylation of inhibitor-4 and the effects on its activity have not yet been studied. In 1995 a phosphoprotein called CPI-17 (for PKC-potentiated inhibitor of 17 kDa) was purified as an inhibitor of myosin phosphatase.11 CPI-17 is similar to I-1 and DARPP-32 in that phosphorylation at a single residue, Thr38, enhances its potency 1000-fold to give an IC50 at nanomolar concentrations and it is specific for PP1 relative to type-2 phosphatases. However, CPI-17 (Thr38) is preferentially phosphorylated by PKC instead of PKA, and CPI-17 was surprisingly different from other PP1 inhibitor proteins because it inhibited with the 10

H. Shirato, H. Shima, G. Sakashita, T. Nakano, M. Ito, E. Y. Lee, and K. Kikuchi, Biochemistry 39, 13848–13855 (2000). 11 M. Eto, T. Ohmori, M. Suzuki, K. Furuya, and F. Morita, J. Biochem. 118, 1104–1107 (1995).

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same potency both PP1C monomer and myosin phosphatase, a PP1 holoenzyme consisting of PP1C bound to regulatory subunits. Phosphorylation of CPI-17(Thr38) is regulated by extracellular stimuli in parallel with myosin phosphorylation and vascular smooth muscle contraction, suggesting that CPI-17 acts to control vascular contractility in response to hormonal signaling.12–14 Proteins related to CPI-17, called PHI-1 and KEPI, were discovered in mammalian cells, and these also can inhibit PP1 holoenzymes, such as myosin phosphatase, but whether they inhibit specific PP1 holoenzymes is not yet known.11,15 These are examples of PP1 inhibitor proteins blocking the activity of PP1 holoenzymes wherein the PP1 catalytic subunit is already complexed to a regulatory subunit. This runs contrary to a popular model that made it into textbooks, showing that PP1 bound at a common site either a targeting subunit or a phosphoinhibitor that both used a VxF motif for recognition. More recent observations support the concept that phosphoprotein inhibitors and regulatory subunits both bind PP1 simultaneously. I-1 was shown to bind to GADD-34 (growth and DNA damage) protein, which also binds PP1 making an I-1–GADD-34–PP1C trimeric complex with I-1 bound to GADD-34 but still able to inhibit the PP1C in a phosphorylationdependent manner.16 Work by our group has discovered that I-2 specifically binds to PP1C on Nek2 kinase to activate kinase activity of the PP1C–Nek2 complex, and I-2 thereby induces centrosome separation.17 I-2 also binds directly to neurabin II/spinophilin18 and to KPI-2, a novel membrane Ser/ Thr kinase,19 both of which also bind PP1. These offer examples of trimeric complexes of a PP1 catalytic subunit plus a regulatory subunit docked to it via a VxF motif plus I-2 that interacts with both of the other subunits. Thus, for CPI-17, I-1 and I-2 there is now evidence that the inhibitor

12

T. Kitazawa, M. Eto, T. P. Woodsome, and D. L. Brautigan, J. Biol. Chem. 275, 9897–9900 (2000). 13 M. Eto, T. Kitazawa, M. Yazawa, H. Mukai, Y. Ono, and D. L. Brautigan, J. Biol. Chem. 276, 29072–29078 (2001). 14 E. F. Etter, M. Eto, R. L. Wardle, D. L. Brautigan, and R. A. Murphy, J. Biol. Chem. 276, 34681–34685 (2001). 15 Q. R. Liu, P. W. Zhang, Q. Zhen, D. Walther, X. B. Wang, and G. R. Uhl, J. Biol. Chem. 277, 13312–13320 (2002). 16 J. H. Connor, D. C. Weiser, S. Li, J. M. Hallenbeck, and S. Shenolikar, Mol. Cell. Biol. 21, 6841–6850 (2001). 17 M. Eto, E. Elliott, T. D. Prickett, and D. L. Brautigan, J. Biol. Chem. 277, 44013–44020 (2002). 18 R. T. Terry-Lorenzo, E. Elliot, D. C. Weiser, T. D. Prickett, D. L. Brautigan, and S. Shenolikar, J. Biol. Chem. 277, 46535–46543 (2002). 19 H. Wang and D. L. Brautigan, J. Biol. Chem. 21, 21 (2002).

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phosphoproteins interact with PP1 that is itself tethered by a targeting subunit. We now imagine that regulation of PP1 in cells incorporates the various inhibitor phosphoproteins to selectively target a subset of PP1 holoenzymes. This way inhibitor proteins work in concert with the PP1 regulatory subunits, not in lieu of them. Kinases can phosphorylate their substrates and simultaneously turn off the opposing PP1 holoenzyme by phosphorylation of these inhibitor subunits. This concept makes for bistable switching circuits, with reciprocal outputs of high kinase/low phosphatase or low kinase/high phosphatase.20 They act as biological on/off switches. Therefore, study of the phosphorylation of these PP1 inhibitors takes on added significance for understanding signaling processes, and here we focus on methods developed for that purpose. Expression and Characterization of PP1 Inhibitor Proteins Preparation of Recombinant Inhibitor Proteins

Heat-stable inhibitor proteins, inhibitor-1, inhibitor-2, CPI-17, PHI-1, and KEPI can be expressed in E. coli as His6, S-tagged fusion protein using pET30 vector system (Novagen, Madison, WI). The S-tagTM sequence peptide has high affinity to a part of ribonuclease S, called S-protein. Various conjugate forms of S-protein, such as peroxidase and agarose-bead conjugates, are available from Novagen, which enable use in detection and purification of recombinant proteins.17 Five inhibitor proteins described above were prepared by the same protocol. The cDNA inserts of inhibitor proteins are obtained by PCR methods from cDNA library or the plasmid of IMAGE clone (Incyte Genomycs, St. Louis, MI). The PCR product is ligated at Bam HI/Eco RI sites of pET30 vector. Buffers Washing buffer [20 mM imidazole–HCl (pH 7.0), 0.5 M NaCl, 20 mM sodium phosphate buffer (pH 7.0)], Elution buffer [0.5 M imidazole–HCl (pH 7.0), 0.5 M NaCl, 20 mM sodium phosphate buffer (pH 7.0)]. Methods 1.

20

Transform E. coli BL21(DE3) with the plasmid vector. Inoculate 50 ml of overnight culture of the bacteria into 450 ml of 2xYT with 30 g/ml kanamycin, and shake for approximately 2 hr at 37
C until OD600 nm > 0.6. Induce the recombinant protein by addition

J. E. Ferrell, Jr., Curr. Opin. Cell. Biol. 14, 140–148 (2002).

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2.

3.

4.

5.

6. 7.

247

of 1.5 ml of 0.4 M isopropylthio- -galactoside and shake for 4–6 hr at 37
C (no longer than 6 hr). Collect the bacteria by centrifugation and wash the pellet with 50 ml of phosphate-buffered saline (PBS). Suspend the bacteria pellet with 20 ml of PBS, including 1 mM EDTA, 0.4 mM Pefabloc SCTM (Roche, Indianapolis, IN, #1429876), and transfer into a 200-ml tall glass beaker. Add 40 mg of lysozyme (Sigma, St Louis, MI, #L-6876), freeze the suspension in 80
C freezer. After thawing out, leave the solution (viscous) for 1 hr on ice, and then incubate for 10 min at 30
C with 10 mM MgCl2 and 10 g/ml DNase I to reduce the viscosity. Add PBS to 50 ml, and add 5 M NaCl and 2-mercaptoethanol to 0.5 M and 0.3% (v/v) respectively. Heat the solution for 10 min in boiling water with gentle shaking. Cool the beaker for 10 min on bench and clarify the solution by centrifugation for 20 min at 10,000g. Transfer the supernatant into a 50-ml conical tube including 2 ml of Ni-NTATM agarose slurry (Qiagen, Valencia, CA, #30210) and incubate for 30 min at room temp with gentle shaking. Collect the resin by centrifugation and transfer into an Econo-PacÕ column (Biorad, Hercules, CA, #7321010). Wash the resin with 50 ml of Washing buffer by gravity flow. Elute the bound protein with Elution buffer and collect each 0.5 ml by 12 tubes. Bring aliquots of fractions to Bradford protein assay. Generally protein is eluted in fractions 2–5. Collect fractions containing the protein, and then dialyze overnight against 200 ml of 50 mM ammonium bicarbonate buffer with 0.1% 2-mercaptoethanol. Lyophilize the protein solution for 24 h. The dried powder could be stored in 20
C for more than 3 years.

This protocol yields over 10 mg of inhibitor proteins with 90% purity from 500 ml of culture. Gel chromatography using an AcA54TM column (1.4  40 cm) can be applied for further purification to obtain nearly homogeneous recombinant proteins. Unlike a N-terminal glutathione Stransferase tag, the His6, S-tag does not interfere with either phosphorylation or inhibitory potency of PP1 inhibitor proteins, so that inhibitor proteins can be used for various assays as fusion proteins.

Preparation of Phospho- and Thiophospho-inhibitor Proteins

Protein kinases for each inhibitor protein are listed in Table I. Excess amount of kinase or prolonged incubation can cause phosphorylations at

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FIG. 1. Inhibition of PP1 by His6, S-CPI-17. PP1 assay was performed in the presence of various concentration of recombinant CPI-17, using 32P-labeled myosin light chain as a substrate. The myosin phosphatase holoenzyme (MLCP) was purified from pig aorta.11 The Mn2 þ -activated PP1C (PP1C0 ) from rabbit skeletal muscle21 was assayed in the presence of 1 mM MnCl2. Phospho (P)-CPI-17 potently inhibits the myosin phosphatase holoenzyme, whereas the Mn2 þ -activated PP1C preparation is less sensitive to P-CPI-17. Because thiophospho-Thr38 is stable, thiophospho (TP)-CPI-17 is capable of inhibiting the PP1C preparation.

nonspecific sites of the inhibitor protein, so conditions have to be optimized by small scale assays. Phosphorylation (20 l) is carried out for 2 hr with 0.2 mg/ml inhibitor protein and various concentration of kinase in 50 mM MOPS-NaOH (pH 7.0) buffer plus 10 mM MgCl2, 1 mM ATP, 1 mM dithiothreitol, 1 M microcystin LR (Calbiochem, San Diego, CA, #475815), and 0.4 mM Pefabloc SC, at 30
C. Reaction is stopped by addition of 10 mg of solid urea and 2 l of 2-mercaptoethanol. Samples are subjected to mobility-shift assay described below (Fig. 3). Thiophosphorylation is done overnight using 1 mM ATP S (Roche, #102342) for thiophosphate donor, instead of ATP. The longer incubation is needed because the rate of thiophosphorylation is lower. For phosphorylation by protein kinase C, 0.1 mM CaCl2, 1 M phorbol-12-myristate-13-acetate and 0.1 mg/ml phosphatidylserine are added to the reaction mixture. To prepare phosphoand thiophospho-inhibitor in large scale, reaction is terminated by heating for 10 min in 100
C bath, and then subjected to dialysis or gel filtration. The preparation can serve in inhibitor assays (Fig. 1), and be used as an affinity ligand for purification of PP1 holoenzymes22 or specific antibodies, described below.

21 22

D. L. Brautigan, C. L. Shriner, and P. A. Gruppuso, J. Biol. Chem. 260, 4295–4302 (1985). S. Senba, M. Eto, and M. Yazawa, J. Biochem. 125, 354–362 (1999).

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PP1 Phosphatase Inhibitor Assay

Protein type-1 phosphatases: Mn2 þ -dependent PP1C preparations are relatively insensitive to phospho-PP1 inhibitor proteins, such as phospho-I123 and phospho-CPI-17 (Fig. 1), in part because phospho-I-1 and phosphoCPI-17 are dephosphorylated by Mn2 þ -dependent PP1C and therefore lose inhibitory potency24 (Fig. 1). On the other hand I-2 inhibits Mn2 þ dependent preparations as potently as native PP1C that does not require added divalent cations.25 These results support our hypothesis that each inhibitor protein recognizes some different conformation of PP1C and these are modulated by allosteric interaction with regulatory subunits. Therefore, it is best to use native PP1C26 or PP1 holoenzymes without added Mn2 þ 11,17,27 for PP1 inhibitor assays. The amount of PP1 that dephosphorylates 30% of substrate in the assay condition should be determined in advance. Assay PP1 activity is assayed at 30
C by measuring dephosphorylation of 32Plabeled phosphorylase a or phospho-myosin light chain as a substrate. Conditions are: 1 M 32P-substrate, 50 mM MOPS-NaOH, pH 7.0, 50 mM NaCl, 0.1 mM EGTA, 10 nM okadaic acid, 1 mM dithiothreitol, 5% glycerol, 0.02% Brij-35TM, 0.4 mM Pefabloc SC, for 10 min at 30
C. Okadaic acid (10 nM) and EGTA (0.1 mM) are added to block any trace of PP2A and PP2B activities in the PP1 preparation. Reaction is initiated by addition of PP1. Data in the absence of PP1 is regarded as blank, set to 0% activity. The PP1 activity minus inhibitor protein is set as 100%, and relative PP1 activities are plotted against various concentrations of inhibitor protein (Fig. 1).

Assays for Phosphorylation of PP1 Inhibitor Proteins

Phosphorylation of inhibitor proteins is analyzed by immunoblotting using phospho-specific antibody or by mobility shift assays on 23

S. Endo, J. H. Connor, B. Forney, L. Zhang, T. S. Ingebritsen, E. Y. Lee, and S. Shenolikar, Biochemistry 36, 6986–6992 (1997). 24 J. G. Foulkes, S. J. Strada, P. J. Henderson, and P. Cohen, Eur. J. Biochem. 132, 309–313 (1983). 25 A. J. Zhang, G. Bai, S. Deans-Zirattu, M. F. Browner, and E. Y. Lee, J. Biol. Chem. 267, 1484–1490 (1992). 26 P. Cohen, S. Alemany, B. A. Hemmings, T. J. Resink, P. Stralfors, and H. Y. Tung, Methods Enzymol. 159, 390–408 (1988). 27 M. Eto, A. Karginov, and D. L. Brautigan, Biochemistry 38, 16952–16957 (1999).

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FIG. 2. Immunoblotting assay using anti-phospho-specific antibody. Recombinant KEPI was phosphorylated for 2 hr at 30
C with various amounts of PKC. Reaction was terminated by addition of solid urea, and a half portion was mixed with 4X SDS sample buffer. The SDS sample was subjected to immunoblotting using anti-(P-Thr73)-KEPI (top panel) and anti-totalKEPI (bottom panel) antibodies.

polyacrylamide gels. The results of two such phosphorylation assays using the same protein samples are shown in Figs. 2 and 3B. Assay using anti phospho-specific antibody is sensitive and can detect multiple phosphorylation sites in a protein independently. Besides, the antibody may be used for immunocytochemistry or immunohistochemistry to analyze phosphorylation of inhibitor protein in cells or tissues. However, anti phospho-specific antibody is not always available and stoichiometry of phosphorylation cannot be determined by the immunoblotting assay (Fig. 2). On the other hand, the mobility shift assay separates phospho from dephospho protein and enables us to measure stoichiometry of phosphorylation. In case no phospho-specific antibodies are available, the mobility shift assay would be a researcher-friendly technique, compared with metabolic labeling methods using 32P-ortho phosphate in an assay. Assay of Phosphorylation by Immunoblotting using Anti-Phospho-specific Antibody (Fig. 2)

Preparation of Phospho-specific Antibodies for Inhibitor Proteins Antigen preparation and antiserum production are available by custom service from several companies. We have used complete commercial services or prepared antibodies by contracting services a la carte. Synthesis of 10 mg of a 13 mer peptide includes a Cys residue at N-terminus and phospho-Ser or -Thr at sixth position from N-terminus. The peptide (2 mg) is conjugated with keyhole limpet hemocyanin (2 mg) via the SH group at Cys residue using m-maleimidobenzoyl-N-hydroxysuccineimide ester. Two rabbits are immunized with the conjugate (0.1 mg/each). At 21 days (not 7 or 14) after first inoculation, 0.05 mg of the conjugate is injected, and this booster injection is repeated every 2 weeks until specific antibody is detected. At 7 days after each booster, a small amount of serum is collected for testing

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antibody production. Production of the specific antibody is tested by dot blot assay or immunoblot assay using unphospho- and phospho-inhibitor proteins, because antibodies that react with phospho-peptide used for antigen may fail to recognize phospho-inhibitor protein. Unphosphoinhibitor protein and phospho-inhibitor protein (2 mg) are conjugated with 2 ml of Affigel 10/15TM (Biorad) resin following manufacturer’s protocol. These recombinant protein affinity resins are critical for purification of the specific antibody. Antiserum (2 ml for protocol 1, or 10–20 ml for protocol 2) from production bleeds (approximately 20 ml every other week) is passed through the unphospho-inhibitor protein resin (called unbound fraction). This removes antibodies against the protein without phosphate. We purify the phospho-specific antibody from this unbound fraction of antiserum. The two different ways for purification depend on the desired quality and intended purpose of the antibody. Protocol 1. The IgG fraction of unbound antiserum is purified using protein A-agarose resin. The antiserum (2 ml) is mixed for 1 hr with 1 ml of Protein A-agarose (Sigma, #P3476) slurry at room temp. The slurry is packed in a Poly-PrepÕ column (Biorad, #731-1550). After washing the resin with 20 ml of PBS, IgG fraction is eluted with 0.1 M Glycine–HCl, pH 1.9 plus 10% 1,4-dioxane. The added organic solvent enhances yield of eluted antibody. Each 0.3 ml of eluant is collected in a tube containing 30 l of 1.5 M Tris–HCl, pH 8.8, so the eluant is drop-by-drop immediately neutralized. Aliquots are subjected to Bradford protein assay. Fractions (2– 3 tubes) containing protein are collected and dialysed against 100 ml of PBS. Protocol 2. To obtain phospho-specific antibody for immunocytochemistry or immunohistochemistry, or to improve quality of the antibody preparation described above, we use phospho-inhibitor protein resin for the purification. The unbound antiserum (10–20 ml) from unphospho-inhibitor protein is mixed overnight with phospho-inhibitor protein resin (1 ml) at 4
C. The resin is collected in Poly-prep column and washed with 20 ml of PBS. The resin is further washed with 10 ml of 0.3 M sodium acetate buffer, pH 5.0. This step often improves specificity of antibody by removing low affinity or non-specific antibodies. The bound antibody is eluted with 0.1 M Glycine–HCl, pH 1.9 plus 10% 1,4-dioxane, and brought to dialysis, as described above. Preparation of Antibody Recognizing both Phospho- and Unphospho-inhibitor (Anti Total-inhibitor Protein)

Both as independent reagents and for important controls showing equal recovery or expression of protein one needs antibodies against phosphoprotein inhibitors that react regardless of the phosphorylation status. Anti

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total-inhibitor protein antibody is produced in rabbits using recombinant protein as an antigen. Alternatively, antigen is inoculated into 2 hens and the specific IgY is prepared from eggs (Aves Lab, Tiggard, OR). Anti totalinhibitor protein IgY is used for immunoblotting with peroxidase-conjugated anti-chicken IgY antibody as a secondary. Antibody is purified as described for protocol 1 above. Preparation of Samples from Tissue Culture Cells for Immunoblotting with Anti-phospho-inhibitor Antibody

Approximately 3–5 mg of total protein are obtained from confluent cells (HEK293) on a 10 cm dish. Reagents SDS lysis buffer (50 mM Tris–HCl, pH 7.4, 1 mM sodium orthovanadate, 1% SDS, 1 mM EDTA), 4x SDS sample buffer (125 mM Tris– HCl, pH 6.8, 8% SDS, 20% glycerol, 0.012% bromophenol blue, 40% 2-mercaptoethanol). Methods 1. 2. 3.

4. 5. 6.

7. 8.

Prepare SDS buffer: Freshly add 1000th vol of 0.4 M Pefabloc SC in SDS lysis buffer. After desired treatment of cells, remove media from culture dish by aspiration. Immediately add 0.5 ml of the SDS buffer per 10 cm dish. Swirl the solution over the cells to ensure rapid lysis and promote denaturation of cellular proteins. Scrape lysate with a cell scraper. The lysate is usually viscous due to cellular DNA. Transfer the lysate to a 1.5-ml tube using wide-tip pipette. Shear the cellular DNA by sonication of the lysate for 2 sec, three times. Clarify the solution by centrifugation for 5 min at room temperature. Determine protein concentration of the samples: Remove an aliquot, dilute with PBS to reduce SDS to 0.1%. Measure total protein concentration using the BCA reagent from Pierce (#23225). * Make calibration curve of protein standards in a 0.1% SDS solution. Add 1/3rd vol of 4x SDS sample buffer. Load 20–100 g of protein for immunoblotting. The sample can be stored at 20
C.

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Measurement of Inhibitor Protein Phosphorylation using Anti-phospho-inhibitor Antibody (Fig. 2)

Samples are subjected to SDS-PAGE and immunoblotting using the phospho-specific antibody. Bovine serum albumin (3%) plus 0.1% Tween 20 in Tris-buffered saline (TBS) gives lower background in anti phosphoinhibitor blot, compared with 5% nonfat milk. Enhanced chemiluminescent method with peroxidase-conjugated secondary antibody (Pierce, Rockford, IL, SuperSignalTM, #34080) is used for sensitive detection. After exposure to X-ray film, the blot is rinsed three times with TBS-T and antibodies on the membrane are stripped for 1 hr at room temp with 0.1 M Glycine–HCl, pH 1.9 plus 1% SDS. The blot is re-stained with the antibody recognizing both unphospho- and phospho-inhibitor protein (anti total-protein antibody). Signals on both X-ray films are quantified using a densitometer, and relative intensity (phospho-inhibitor protein/total-inhibitor protein) represents relative phosphorylation level of inhibitor protein in arbitrary units. Linearity of the relationship between relative intensity and amount of phospho-protein is preexamined on a blot loaded with various amount of phospho-inhibitor protein. Alternatively, colorimetric detection with alkaline phosphataseconjugated secondary antibody (Biorad, #170-6432) can be used for the assay. This method gives linearity over a wider range of relative intensity and amount phospho-protein, but requires duplicate blots for anti phospho- and anti total-protein staining.12 Protein samples from cultured cells stimulated for 10 min with 10 nM calyculin A, prepared as described above, can be subjected to the assay to gauge extent of phosphorylation. Assay of Phosphorylation by Mobility Shift in SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) (Fig. 3A)

Not all phosphoinhibitors show changed mobility in SDS-PAGE, but wherever it applies this method becomes handy. Untagged I-2 but not His6, S-tagged I-2 shows mobility shift on SDS-gel, depending on the phosphorylation. Phospho-I-2 (P-I-2) migrates slower than unphosphoform (U-I-2). Figure 2 shows the results of a SDS-PAGE/immunoblot for P-I-2 at Ser86, 120, and 121 (right lane) and U-I-2 after reaction with calf intestine alkaline phosphatase.28 The protocol for determination of the phosphorylation sites of I-2 is described in the last section.

28

C. Leach and D. L. Brautigan, unpublished results.

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FIG. 3. Mobility shift assay for phospho-inhibitor proteins. (A) SDS-PAGE for phospho-I2. HeLa cells were lysed with SDS lysis buffer and subjected to immunoblotting (left lane). Cells were extracted with the buffer without SDS and incubated for 2 hr with calf intestine alkaline phosphatase (1 unit) and analysed by immunoblotting (right lane). (B) Urea-PAGE for phospho-KEPI. The urea sample after phosphorylation (Fig. 2) was subjected to urea-PAGE and proteins were stained with GelCode Blue solution. Because the gel contains no SDS, the negative charge of the phosphate is sufficient to increase the migration. (C) Lutidine-PAGE for phospho-CPI-17. Purified unphospho- and phospho-CPI-17 were subjected to lutidine-PAGE, and detected using Coomassie blue staining. Because the polarity of electrophoresis was reversed with usual running configuration, such as SDS-PAGE or urea-PAGE, phospho-CPI17 migrates slower than unphospho-form.

Methods 1. 2. 3.

4.

Samples of cultured cells are prepared as described above. The samples are subjected to the SDS-PAGE Laemmli system [12% gel (29 : 1), 0.75 mm thick, and 11 cm long]. A 120 V constant voltage is applied on the gel until the 50 kDa and 35 kDa markers of BioRad Broad Range prestained markers are separated by 2 cm. Detect I-2 on the gel by Coomassie staining or immunoblotting using anti total-I-2 antibody.

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Assay of Phosphorylation by Mobility Shift in Glycerol PAGE

Glycerol PAGE method was established by J. T. Stull for measurement of myosin light chain phosphorylation.29 The technique can be applied for measurement of PHI-1 phosphorylation. In this gel, phospho-PHI-1 migrates faster than unphospho-form. Reagents Urea sample buffer (9 M urea, 10 mM -glycerophosphate, 1 mM EDTA, 10% 2-mercaptoethanol, 0.4 mM Pefabloc SC, 0.002% bromophenol blue), 10x Glycerol PAGE (GP) buffer (0.2 M Tris base, 0.23 M Glycine), 30% Acrylamide/Bis solution, 29 : 1 (Bio-Rad, #161-0156), 10% (w/v) ammonium persulfate solution. Methods 1. Preparation of protein samples using trichloroacetic acid (TCA) for non-SDS gel electrophoresis: a. Cultured cells in 60 mm dishes are grown to  80% confluency. b. After experimental treatment, remove medium, and then add 500 l of ice-cold 10% (w/v) TCA solution, immediately. c. Leave the dish for 10 min on ice and scrape the cells with a cell lifter and transfer the suspension into a 1.5 ml tube. d. Collect the insoluble precipitate of total cell proteins by centrifugation for 10 min at 20,000g. e. Homogenize the protein pellet with 500 l of diethyl ether by a brief sonication pulse to disrupt the pellet. Collect the pellet by centrifugation for 5 min at 20,000g. Repeat the ether washing step two more times. This removes the TCA from precipitated proteins. f. Leave the tube for 10 min on bench to evaporate residual ether. g. Suspend the protein pellet with 50 l of urea sample buffer using a glass stick and add 10 crystals of urea to make the solution saturated with urea. h. Incubate for 30 min at 30
C. i. Sonicate the solution for 2 sec and clarify the solution by centrifugation for 5 min at 20,000g. 2. Gel preparation: Prepare 7 ml of polyacrylamide solution for a 70  80  1 mm gel. Glycerol H 2O 29

3.7 g 0.6 ml

D. Taylor and J. T. Stull, J. Biol. Chem. 263, 14456–14462 (1988).

256

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30% Acrylamide/Bis 10x GP buffer Degas the solution for 5 min. 10% ammonium persulfate N,N,N0 ,N0 -tetramethylethylenediamine (TEMED)

2.8 ml 0.7 ml 80 l 7 l

Immediately pour the solution between glass plates, and then let the gel polymerize for 2 h. The gel can be stored overnight at 4
C wrapped in wetted paper towels and plastic film. 3. Reservoir buffer (200 ml): 10x GP buffer H2O Mercaptoacetic acid Dithiothreitol

20 180 228 140

ml ml mg mg

4. Prerun: Set the gel to the apparatus and apply 300 V of constant voltage for 1 hr in advance to loading samples. 5. Electrophoresis: Load 5 l of urea-samples on the gel and then apply 300 V for 2 h. 6. Immunoblotting: Soak the gel for 30 min in 50 ml of electrotransfer buffer (0.02 M Tris, 0.125 M Glycine, 5% methanol) at room temp. Transfer onto nitrocellulose membrane and perform immunoblotting using anti total-PHI-1 antibody. 7. Quantification: Quantify intensity of the X-ray film derived from monophospho-, diphospho- and unphospho-PHI1 bands using a densitometer. Calculate stoichiometry of the phosphorylation using an equation: Phosphorylation of PHI-1 (mol/mol) ¼ Intensity (mono þ di)/Intensity (mono þ di þ un). Assay of Phosphorylation by Mobility Shift in Urea PAGE (Fig. 3B)

Phosphorylation can be detected by urea-PAGE method. This method is capable of measuring phosphorylation of I-1, I-2, PHI-1,27 and KEPI (Fig. 3B). Reagents 10x urea PAGE (UP) buffer (0.2 M Tris, 1.25 M Glycine). Methods 1. Sample preparation: Add 10 mg of crystalline urea, 2 l of 2mercaptoethanol, and 1 l of 0.1% bromophenol blue to 10 l of protein solution. Dissolve urea by agitation, but some urea crystals will remain in

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solution (showing saturation). Do not heat the sample. Samples from cultured cells described in glycerol PAGE section can be used for urea-PAGE. 2. Gel preparation: Prepare 7 ml of polyacrylamide solution for a 70  80  1 mm gel: solid urea 30% Acrylamide/Bis 29 : 1 10x UP buffer H 2O * dissolve urea completely 10% ammonium persulfate TEMED

3.36 g 2.8 ml 1.4 ml up to 7 ml 100 l 8 l

Immediately pour the solution between the glass plates, and then let the gel polymerize for 0.5 h. The gel can be stored for a night on bench wrapped in wetted paper towels and plastic film. (Do not refrigerate because urea will crystallize at 4
C.) 3. Reservoir buffer: 15 ml of 10x UP buffer plus 135 ml of H2O. 4. Electrophoresis: Apply constant voltage for 2 hr at 150 V. Tracking marker (bromophenol blue) will be run-out from gel. 5. Detection: After electrophoresis the gel is soaked for 30 min in 40% ethanol to remove urea. The washed gel can be subjected to protein staining with GelCodeÕ Blue Stain solution (Pierce, #94590). For immunoblotting, gel is washed for 30 min in electrotransfer buffer (0.02 M Tris, 0.125 M Glycine, 5% methanol) prior to blotting. Protein on membrane is analyzed as described in Glycerol PAGE method. 6. Quantification: Quantify intensity on the Coomassie-stained gel or the X-ray film derived from monophospho-, diphospho- and unphosphoinhibitor bands using a densitometer. Calculate stoichiometry of the phosphorylation using an equation: Phosphorylation (mol/mol) ¼ Intensity (mono þ di)/Intensity (mono þ di þ un). Assay of Phosphorylation by Mobility Shift in Lutidine PAGE (Fig. 3c)

Basic proteins, such as CPI-17, migrate upward on the glycerol- and urea-PAGE systems described above. To detect phosphorylation of CPI-17 one needs to use a reversed polarity gel system, the Lutidine PAGE method.30 This method is also capable of detecting phosphorylation of other basic proteins, such as histones and myelin basic protein.

30

T. Kitazawa, N. Takizawa, M. Ikebe, and M. Eto, J. Physiology 520, 139–152 (1999).

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Reagents 1.0 M MOPS-NaOH (pH 8.0) buffer, 2,6-lutidine (ACROS ORGANICS #12519, NJ, USA), 0.2% (w/v) pyronin Y (Fisher, #BP105) in H2O. Methods 1. Sample preparation: Add 10 mg of crystal urea, 2 l of 2mercaptoethanol, and 1 l of 0.2% pyronin Y to 10 l of protein solution. Dissolve urea by agitation. Some urea crystals should remain in solution (showing saturation). Do not heat the sample. 2. Gel preparation: Prepare 7 ml of polyacrylamide solution for a 70  80  1 mm gel: Urea 30% Acrylamide/Bis 29 : 1 1.0 M MOPS–NaOH, pH 8.0 H 2O * dissolve completely 10% APS TEMED

3.36 g 2.0 ml 0.7 ml up to 7 ml 100 l 8 l

Immediately pour the solution between the glass plates, and then let the gel polymerize for 0.5 h. The gel can be stored overnight on bench at room temperature wrapped in wetted paper towels and plastic film. (Urea will crystallize at 4
C.) 3. Upper reservoir buffer: 5.25 g of MOPS, 2.95 ml of 2,6-lutidine plus H2O to 500 ml. 4. Bottom reservoir buffer: 5 ml of 1.0 M MOPS-NaOH (pH 8.0) plus 45 ml of H2O. 5. Electrophoresis: Reverse the polarity of electrophoresis relative to usual running configuration. Connect the positive electrode (red line) to the upper (not lower) reservoir and the negative electrode (black line) to the lower (not upper) reservoir. Apply constant voltage for 3 hr at 150 V. Tracking marker (pyronin Y) will be run-out from gel. 6. Detection and quantification: After electrophoresis, gel is subjected to Coomassie staining or immunoblotting as described in urea PAGE method. Phospho-CPI-17 migrates slower than unphospho-CPI-17 (Fig. 3c). Determination of in vivo Phosphorylation Sites of Endogenous Inhibitor Proteins

Many proteins, especially phosphatase inhibitors, undergo exceedingly rapid dephosphorylation in cells during lysis, even with phosphatase

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inhibitor compounds. We adopted an acid fixation method to preserve and determine phosphorylation of endogenous proteins. Analysis of phosphorylation shows that Ser86, 120 and 121 of inhibitor-2 in HeLa cells are phosphorylated in HeLa cells.31

Buffers

Urea Resolubilization Buffer (9 M urea, 20 mM Tris, 23 mM glycine, 1 mM EDTA, and 10 mM -glycerophosphate); Immunoprecipitation buffer [50 mM Tris–HCl (pH 8.0) with 1% IGEPAL CA630 (Sigma), 0.5% deoxycholate, 200 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 M microcystin-LR, 20 mM -glycerophosphate, 1 mM sodium orthovanadate, 0.1% 2-mercaptoethanol, and 0.4 mM Pefabloc SC]; Wash buffer [50 mM MOPS-NaOH (pH 7.4) with 0.1% IGEPAL CA630, 150 mM NaCl, 5 mM EGTA, 5 mM EDTA, 1 M microcystin-LR, 20 mM -glycerophosphate, 1 mM sodium orthovanadate, and 0.1% 2-mercaptoethanol].

Methods

1. Fixation and immunoprecipitation of I-2 from HeLa cell lysate: Culture medium of HeLa cells is removed by aspiration and the dish is placed on a mixture of acetone and dry ice, then 1 ml of ice-cold 5% trichloroacetic acid is added for 10 min. Fixed cells are scraped from the dish using a cell lifter. The slurry is transferred to a microcentrifuge tube and a pellet is collected by centrifugation for 10 min at 20,000g. The supernatant is discarded and the pellet is washed twice with cold acetone to remove the TCA. The I-2 is redissolved with 1 ml of urea resolubilization buffer. After incubation for 1 hr at 30
C, the suspension is diluted 10-fold with immunoprecipitation buffer in a 15-ml conical tube. Following centrifugation for 10 min at 10,000g, I-2 is purified by immunoprecipitation technique. The extract is incubated for 2 hr at 4
C with 5 g of affinity purified sheep anti-I-2 antibody32 coupled to protein G beads in a 15-ml conical tube. The beads are collected by centrifugation and washed with Wash buffer three times. The beads are resuspended in SDS-containing sample buffer and the eluted proteins are subjected to SDS-PAGE and stained with Coomassie Blue. 2. Mass spectrometric analysis: The I-2 band is excised from the gel, washed and destained with 0.5 ml of 50% methanol/5% acetic acid overnight at room temperature before dehydration in 200 l of acetonitrile 31 32

C. A. Leach, S. Shenolikar, and D. L. Brautigan, J. Biol. Chem. 278, 26015–26020 (2003). C. Leach, M. Eto, and D. L. Brautigan, J. Cell. Sci. 115, 3739–3745 (2002).

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and complete drying in a vacuum centrifuge. Protein SH groups are reduced by addition of 50 l of 10 mM dithiothreitol and alkylated by addition of 50 l of 100 mM iodoacetamide (both 30 min at room temperature). The gel pieces are dehydrated again and rehydrated in 50 l of 20 ng/l sequencinggrade modified porcine trypsin (Promega) for 5 min on ice. Any excess trypsin solution is removed and the digestion carried out overnight at 37
C. The peptides produced in the digest are extracted with 50 l of 50 mM ammonium bicarbonate plus 50 l of 50% acetonitrile/5% formic acid (2X), and the total extract is concentrated in a vacuum centrifuge to 20 l for analysis. The LC–MS system consisted of a Finnigan LCQ (ThermoQuest) iontrap mass spectrometer with a Protana nanospray ion source interfaced to a self-packed 8 cm  75 m i.d. Phenomenex Jupiter 10 m C18 reverse-phase capillary column. Peptides including phospho-peptides are identified by comparison of their molecular masses with theoretical number. Second dimension mass spectra of product ion from first mass spectra (MS/MS) are used to determine amino acid sequence and phospho-Ser or phospho-Thr in a peptide.33 Acknowledgments This work was supported by the AHA National Center Scientist Development Grant (to M. E.), and USPHS grants GM56362 and CA40042 (to D. L. B.). Partial support for C. L. was provided by the University of Virginia Cell and Molecular Biology Program training grant GF-10461.

33

The mass spectrometric analyses of phospho-I-2 in SDS-gel were done in The W. M. Keck Biomedical Mass Spectrometry Lab at the University of Virginia.

[19] Combinatorial Chemistry and Peptide Library Methods to Characterize Protein Phosphatases By STEFAN W. VETTER and ZHONG-YIN ZHANG

The methods of combinatorial chemistry have developed tremendously over the last decade and have altered many aspects of medicinal chemistry, drug discovery, and the way biological problems can be approached with the

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and complete drying in a vacuum centrifuge. Protein SH groups are reduced by addition of 50 l of 10 mM dithiothreitol and alkylated by addition of 50 l of 100 mM iodoacetamide (both 30 min at room temperature). The gel pieces are dehydrated again and rehydrated in 50 l of 20 ng/l sequencinggrade modified porcine trypsin (Promega) for 5 min on ice. Any excess trypsin solution is removed and the digestion carried out overnight at 37
C. The peptides produced in the digest are extracted with 50 l of 50 mM ammonium bicarbonate plus 50 l of 50% acetonitrile/5% formic acid (2X), and the total extract is concentrated in a vacuum centrifuge to 20 l for analysis. The LC–MS system consisted of a Finnigan LCQ (ThermoQuest) iontrap mass spectrometer with a Protana nanospray ion source interfaced to a self-packed 8 cm  75 m i.d. Phenomenex Jupiter 10 m C18 reverse-phase capillary column. Peptides including phospho-peptides are identified by comparison of their molecular masses with theoretical number. Second dimension mass spectra of product ion from first mass spectra (MS/MS) are used to determine amino acid sequence and phospho-Ser or phospho-Thr in a peptide.33 Acknowledgments This work was supported by the AHA National Center Scientist Development Grant (to M. E.), and USPHS grants GM56362 and CA40042 (to D. L. B.). Partial support for C. L. was provided by the University of Virginia Cell and Molecular Biology Program training grant GF-10461.

33

The mass spectrometric analyses of phospho-I-2 in SDS-gel were done in The W. M. Keck Biomedical Mass Spectrometry Lab at the University of Virginia.

[20] Combinatorial Chemistry and Peptide Library Methods to Characterize Protein Phosphatases By STEFAN W. VETTER and ZHONG-YIN ZHANG

The methods of combinatorial chemistry have developed tremendously over the last decade and have altered many aspects of medicinal chemistry, drug discovery, and the way biological problems can be approached with the

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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help of chemistry.1,2 Combinatorial methods provide tools and strategies to quickly generate large numbers of chemically related compounds (libraries). These libraries can then be screened for activity against a biological target and structure activity relations (SARs) can subsequently be established. The ability to synthesize and to screen thousands of compounds very rapidly accelerates the process of lead structure identification greatly. In the context of characterizing protein phosphatases two aspects are of particular interest: (1) the assessment of the preferred substrates; and (2) the identification of compounds inhibiting the catalytic activity. The process of characterizing protein phosphatases using combinatorial library methods can be conceptually divided into three steps: (1) synthesis of the library; (2) screening of the library; and (3) deducing structure activity relations and identifying consensus or lead structures. It is not possible to give a ‘‘one-fits-all’’ protocol for the use of combinatorial library methods in general, nor for the characterization of protein phosphatases or PTPs in particular. There are many possible experimental approaches for each strategic step and the selection of the right library, the right assay and the right deconvolution strategy depend on many parameters, not at least on the intended goal of the project and the experimental capabilities of the laboratory where the experiments are actually performed. Therefore, only general considerations for each step will be presented here, and a number of examples will be discussed briefly. It also should be noted that the examples presented in this chapter will focus on PTPs. However, the strategies discussed here can be applied to related proteins such as serine/threonine protein phosphatases, dual specific protein phosphatases, and also to SH2 or PTB domain containing proteins.3

Synthesis of Combinatorial Libraries

‘‘Combinatorial libraries’’ are collections of chemical compounds, which have been synthesized in a combinatorial manner. All compounds share a common history of chemical synthesis steps and are structurally related to each other. Each compound can be considered a derivative from any other compounds in the library. A major element in the success of combinatorial chemistry is the ability to chemically synthesize compounds not in solution, but covalently attached to a solid support. Solid phase 1

M. A. Gallop, R. W. Barrett, W. J. Dower, S. P. A. Fodor, and E. M. Gordon, J. Med. Chem. 37, 1384–1401 (1994). 2 M. J. Plunkett and J. A. Ellman, Sci. Amer., 54–59 (1997). 3 S. W. Vetter and Z.-Y. Zhang, Curr. Prot. Pept. Sci. 3, 365–397 (2002).

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synthesis (SPS) eliminates the need for purification of intermediates in multistep chemical transformations because the desired reaction product remains attached to the solid support, while by-products, reagents, and solvents can be removed by simple filtration. The merger of solid phase synthesis and combinatorial strategies allows rapid access to large collections of synthetically prepared compounds (libraries). These libraries can consist of any sort of molecule accessible through chemical synthesis on a solid support. Solid phase synthesis had been pioneered 50 years ago for the synthesis of peptides4 and oligonucleotides and, not surprisingly, the first combinatorial libraries were peptide libraries.5,6 There are two fundamentally different approaches to combinatorial library synthesis:7,8 ‘‘mixture coupling’’ and ‘‘split and recombine.’’ The latter is also known as ‘‘split and mix’’ or ‘‘one-bead one-compound.’’ The first method uses mixtures of reagents in a single reaction vessel, and can be performed on various solid supports, such as beaded resins, plastic pins, or spotted onto membrane sheets. This results in multiple reaction products in a single synthesis step. Libraries obtained by this method can be very large, because the amount of an individual compound decreases proportionally with increasing library size. For instance, a library of random hexapeptides can be synthesized by coupling six times a mixture of 20 amino acids (diversity elements), resulting in 206 ¼ 6.4  107 different peptide sequences. In contrast, the ‘‘split and recombine’’ (one-bead one-compound) method uses one reagent per reaction and subsequently produces only one reaction product per reaction step.9,10 To obtain libraries it is necessary to have one reaction compartment for each diversity element incorporated into the library. For instance, to synthesize a randomized hexapeptide containing all 20 amino acids one needs 20 reaction vessels to couple each amino acid individually onto a 1/20 fraction of the solid support (e.g., beaded resin). The 20 fractions are then recombined into a single reaction vessel after the coupling has been completed, mixed, deprotected and split again into 20 fractions prior to the coupling of the next amino acid. This process is repeated six times until the hexapeptide is completed. This synthesis mode 4

B. Merrifield, Methods Enzymol. 289, 3–13 (1997). H. M. Geysen, S. J. Rodda, and T. J. Mason, Mol. Immunol. 23, 709–715 (1986). 6 K. S. Lam, S. E. Salmon, E. M. Hersh, V. J. Hruby, W. M. Kazmierski, and R. J. Knapp, Nature 354, 82–84 (1991). 7 G. Jung (ed.), ‘‘Peptide and Non-peptide Libraries: A Handbook for the Search of Lead Structures.’’ VCH, Weinheim, 1996. 8 M. Lebl and V. Krachnak, Methods Enzymol. 289, 336–392 (1997). 9 A. Furka, F. Sevenstyen, M. Asgedom, and G. Dibo, Int. J. Prot. Pept. Res. 37, 487–493 (1991). 10 K. S. Lam, M. Lebl, and V. Krchnak, Chem. Rev. 97, 411–448 (1997). 5

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results in libraries where each resin bead carries one peptide sequence. Consequently one needs at least one bead for each peptide in the library. This limits the possible library size to the amount of resin beads one can work with. Practically, one has to use 5–10 resin beads per peptide sequence in the library to ensure a completeness of > 99%. Typical peptide resin contains about 106–108 beads per gram of resin (depending on the size of the beads), therefore limiting the possible size of a ‘‘one-bead one peptide’’ library to about 105–107 members per gram of resin. ‘‘One-bead one-compound’’ libraries are most frequently synthesized on beaded resins, but can also be synthesized on plastic pins, crowns or capsules when larger amounts of material are desired. An alternative to the two combinatorial strategies for library synthesis outlined above is the parallel synthesis of individual compounds. Parallel synthesis requires one defined reaction compartment for each compound to be synthesized and is done on beaded resins distributed into multiwell plates, tagged plastic pins, capsules or on sheet like solid supports (e.g., paper sheets).11–13 Each method has intrinsic advantages and disadvantages. Parallel synthesis is limited to small libraries (hundreds or thousands of compounds), but has the advantage that the identity of each compound is known at all times during the entire synthesis and assaying procedure. Library synthesis by mixture coupling allows generation of very large libraries, but the verification of the presence of individual compounds or general quality assessment of the library is difficult due to the complexity of the mixtures. In addition, the procedure assumes that it is possible to create mixtures of reactants which lead to equimolar product mixtures, but this may be difficult to achieve in many cases.14,15 ‘‘One-bead one-compound’’ libraries offer several advantages. The quality of the library can be tested by analytically characterizing individual beads/compounds and near equimolar representation of all library members can be ensured. The screening can be done with the compounds still bound to the solid support or after their release into solution. The screening can be done either on the level of single compounds or with mixtures of compounds. A minor disadvantage might be that certain compounds e.g., peptide sequences, which are known to be difficult to synthesize (due to 11

E. J. Nicholas, J. Immunol. Methods 267, 3–11 (2002). R. Frank, J. Immunol. Methods 267, 13–26 (2002). 13 N. A. Boyle and K. D. Janda, Curr. Opin. Chem. Biol. 6, 339–346 (2002). 14 K. M. Ivanetich and D. V. Santi, Methods Enzymol. 267, 247–260 (1996). 15 J. A. Boutin, I. Gesson, J. M. Henlin, S. Bertin, P. H. Lambert, J. P. Volland, and J. L. Fauchere, Mol. Diversity 3, 43–60 (1997). 12

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intermolecular aggregation of the peptide on the resin), might be slightly underrepresented in the library.16

Synthesis of (Phospho)peptide Libraries

Chemical libraries can be based on any chemical scaffold, but the most easily accessible scaffold are linear oligoamides, in particular peptides. In addition, peptides are reasonably good substitutes/models for phosphoproteins, the in vivo targets of protein phosphatases. For these reasons, the synthesis of peptide-, phosphopeptide-, and modified peptide libraries will be discussed here, but not the synthesis of nonpeptide derived chemical libraries (e.g., substituted aromatic compounds, substituted polycyclic compounds, linear and branched polyamines, etc.).17–19 Solid phase peptide synthesis (SPPS) has been refined and automated to a high degree over the last 50 years and today peptides can be synthesized fully automated in high quality and yield.20 At the same time, all reagents necessary for the synthesis of peptides are commercially available, easy and safe to handle. The coupling reaction to form an amide bond between two amino aids is not particularly sensitive to oxygen or moisture, and can be performed in open reaction vessels. This makes it possible to synthesize peptides without the need for expensive or specialized equipment, enabling every biochemist to synthesize peptides in a standard laboratory setting. This point is worth mentioning because many biological or biochemically oriented laboratories would like to take advantage of synthetic peptides and peptide libraries, but cannot afford expensive custom synthesis or automated peptide synthesizers. The synthesis of phosphorylated peptides is not different from the synthesis of nonphosphorylated peptides if one uses preformed phosphorylated building blocks, which are commercially available for tyrosine, serine and threonine. Phosphorylated peptides are very suitable to assess the substrate specificity of protein phosphatases, because they can function as actual substrates. To address questions of phosphopeptide binding or PTP 16

R. C. d. L. Milton, S. C. F. Milton, and P. A. Adams, J. Am. Chem. Soc. 112, 6039–6046 (1990). G. M. Figliozzi, R. Goldsmith, S. C. Ng, S. C. Banville, and R. N. Zuckermann, Methods Enzymol. 267, 437–447 (1996). 18 B. A. Bunin, M. J. Plunkett, and J. A. Ellmann, Methods Enzymol. 267, 448–465 (1996). 19 A. Nefzi, J. Ostresh, and R. A. Houghten, Chem. Rev. 97, 449–472 (1997). 20 G. B. Fields, J. Lauer-Fields, R.-Q. Liu, and G. Barany, Principles and practice of solid phase peptide synthesis, in ‘‘Synthetic Peptides’’ (G. A. Grant, ed.), pp. 92–219. Oxford University Press, New York, 2002. 17

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inhibition it is necessary to form stable PTP-peptide complexes, which is not feasible with phosphopeptides because they can be dephosphorylated by the phosphatase. For this purpose, a number of phosphotyrosine mimics or nonhydrolysable phosphotyrosine building blocks have been developed and some of them are commercially available.21–23 Although peptides typically show poor bioavailability and are easily degraded by peptidases, they often provide the easily accessible lead structures for further optimization. To improve bioavailability and in vivo stability, peptides can be modified in many ways, for example, by incorporating nonnative amino acids (dozens are commercially available) into the peptide, N-terminal modification by coupling various carboxylic acids, and alterations of side chains or the peptide backbone (e.g., by alkylation of the amide bond).

Synthesis of Peptides, Phosphopeptides and N-terminally Modified Peptides

The literature on peptide synthesis is very extensive and many excellent review and books are available.24–26 However, the volume of available information makes it sometimes difficult to extract the essential information to synthesize a first peptide (library). Therefore, a few tips regarding certain aspects of peptide synthesis are given here.

Fmoc- vs Boc-chemistry27

Amino acids need temporary protection of their amino groups during amide bond formation and either the Fmoc- or Boc-group are routinely used for that. Fmoc-chemistry is preferable, because Fmoc removal is achieved using piperidine, whereas Boc-group removal requires trifluoroacetic acid (TFA). In addition, the final cleavage of Fmoc-chemistry based

21

T. R. Burke, B. Ye, M. Akamatsu, H. Ford, X. Yan, H. K. Kole, G. Wolf, S. E. Shoelson, and P. P. Roller, J. Med. Chem. 39, 1021–1027 (1996). 22 T. R. Burke, M. S. Smyth, A. Otaka, and P. P. Roller, Tetrahedron Lett. 34, 4125–4128 (1993). 23 B. Ye and T. J. Burke, Tetrahedron Lett. 37, 4733–4736 (1995). 24 W. Chang and P.D. White (eds.), ‘‘Fmoc Solid Phase Peptide Synthesis: A Practical Approach.’’ Oxford University Press, Oxford, 2000. 25 G. B. Fields (ed.), ‘‘Solid-Phase Peptide Synthesis.’’ Academic Press, San Diego, 1997. 26 S. Aimoto, Curr. Org. Chem. 5, 45087 (2001). 27 D. A. Wellings and E. Atherton, Methods Enzymol. 289, 44–67 (1997).

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peptides is done with TFA, whereas Boc-based peptide require the use of liquified HF, which is difficult and hazardous to handle and requires expensive, HF resistant labware.

Reaction Vessel

Peptides can be conveniently synthesized in open cartridges or tubes usually used for solid phase extraction. These tubes (e.g., Sigma-AldrichSuppelco, prod. #57242 or #57179) are made from polypropylene, can be equipped with filter frits, closed with Luer plugs, and are resistant to all reagents and solvents used for peptide synthesis, including TFA during cleavage and deprotection. An inverted powder funnel taped onto an orbital shaker can function as primitive holder for these cartridges. The amount of peptide synthesis resin used in these tubes should be small enough to allow the resin beads to swirl freely once the shaker is set into motion. Typically, 50 mg of resin can be used in 6 ml cartridge and 200 mg in a 20 ml cartridge. Solvents and reagents are removed easily by suction through the filter frit at the bottom of the cartridge into a suction flask connected to a vacuum source.

Solid Support

The choice of the solid support depends on the assay strategy used later and whether the assay will be done in solution or with the peptide still bound to the solid support. If the assay will be done on the solid support, then a material has to be chosen, which will swell in the assay buffer. Hydrophilic peptide resins based on polyethylene glycol (e.g., TentaGelÕ ), or polyamides (e.g., Clear ResinÕ ) should be used.28,29 If the peptide will be assayed in solution, a standard, polystyrene based resin can be employed. Synthesis on paper sheet is possible too.12,30–32 Peptides can be released from the solid support either as C-terminal acids or amides by choice of the appropriate linker (Wang linker resin for acids,33 Rink linker resin for amides34). Typically C-terminal amides are preferred. 28

M. Kempe and G. Barany, J. Am. Chem. Soc. 118, 7083–7093 (1996). C. Blackburn, Biopolymers 47, 311–351 (1999). 30 R. Frank and R. Doering, Tetrahedron 44, 6031–6040 (1988). 31 R. Frank, Tetrahedron 48, 9217–9232 (1992). 32 R. Frank and H. Overwin, Meth. Mol. Biol. 66, 149–169 (1996). 33 S. S. Wang, J. Am. Chem. Soc. 95, 1328–1333 (1973). 34 H. Rink, Tetrahedron Lett. 28, 3787–3790 (1987). 29

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Solvents

Amino acid couplings, Fmoc removal and washing can be done using DMF as the sole solvent. The solvent quality should be ‘‘HPLC grade’’ or ‘‘ACS spectrophotometric grade’’ quality, but does not necessarily need to be of ‘‘peptide synthesis’’ quality.

Activation Reagents

Dozens of activation reagents have been described. Uronium salt based reagents, in particular HBTU ([2-(1H-benzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate] and TBTU [2-(1H-benzotriazole-1yl)-1,1,3,3-tetramethyluronium tetrafluoroborate]) are very efficient in DMF, work generally well, are easy and safe to handle, and should be the first choice.

Amino Acid Derivatives

Fmoc-protected amino acids can be purchased from many vendors and standard side chain protection groups can be used for phosphopeptide synthesis. Preformed phosphotyrosine, phosphothreonine, and phosphoserine building blocks are available with various side chain protection groups.35–37 Mono-O-benzyl protected derivatives (Fmoc-Tyr/Ser/ Thr(PO(OBzl)OH)-OH) are recommended, because they and the amino acid immediately following them can be relatively easily incorporated into the peptide.38–40 35

J. W. Perich, R. M. Valerio, and R. B. Johns, Tetrahedron Lett. 27, 1377–1380 (1986). J. W. Perich, Methods Enzymol. 289, 245–266 (1997). 37 E. A. Kitas, J. W. Perich, J. D. Wade, R. B. Johns, and G. W. Tregear, Tetrahedron Lett. 30, 6229–6232 (1989). 38 H. Schmid, S. Vetter, W. Bannwarth, and E. Kitas, Diester building blocks of pSer and pThr suitable for Fmoc solid phase peptide synthesis, in ‘‘Innovation and Perspectives in Solid Phase Synthesis and Combinatorial Libraries’’ (R. Epton, ed.), pp. 525–528. Mayflower Scientific Ltd, Birmingham, 1996. 39 H. Schmid, S. Hoving, S. Vetter, T. Vorherr, and E. Carafoli, Synthesis and purification of unphosphorylated and phosphorylated phospholamban, in ‘‘Pept. Proc. Am. Pept. Symp.’’ (J. P. Tam and P. P. Kaumaya, eds.), Vol. 15, pp. 709–710. Dordrecht, NL, Kluwer, 1999. 40 P. White and J. Beythien, Preparation of phosphoserine, threonine and tyrosine containing peptides by the Fmoc methodology using pre-formed phosphoamino acid building blocks, in ‘‘Innovation and Perspectives in Solid Phase Synthesis’’ (R. Epton, ed.), pp. 553–560. Mayflower Scientific Ltd, Birmingham, 1996. 36

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Amino Acid Coupling

Fmoc-amino acid derivatives should be dissolved in DMF and activated with 0.9 equivalent of HBTU, 0.9 equivalents of HOBt and 1.8 equivalents of sym.-collidine (2,4,6-trimethylpyridine) as base. A three-fold excess of amino acid derivative relative to the resin incubated for 20 min to 1 hr usually gives satisfactory coupling efficiency. Completeness of the coupling should always be controlled by the Kaiser test.41 If the coupling was incomplete, or the amino acid derivative is in short supply or expensive, then multiple couplings can be performed, for instance by coupling first 1.2 equivalents of the derivative for several hours, followed by a second (or third) coupling with 0.3 equivalents overnight. After removal of the coupling mixture the resin needs to be washed several (5–8) times with DMF. Fmoc Removal

The temporary Fmoc-protection group can be removed by incubating the resin in 20% piperidine in DMF for 2  3 min and an additional 5 min reaction.42 After removal of the Fmoc group the resin should be washed with DMF several (8–12) times to remove completely the remaining piperidine. The resin should be shaken thoroughly during amino acid couplings and Fmoc removal. Deprotection/Cleavage

Deprotection protocols for Fmoc-based peptides vary mainly in the amount and sort of added scavengers, reagents which quench reactive side products produced during deprotection. A cleavage mixture of 92% trifluoroacetic acid, 5% triisopropylsilane, and 3% water works well in many cases and minimizes contaminations of the library with remaining scavengers. Other popular cleavage cocktails are reagent K or reagent B.43 If benzylation of tryptophan residues is a concern (release of benzyl cation from O-benzyl protected phosphotyrosine building blocks), then a small amount of Fmoc-Trp-OH can be added to the cleavage cocktail as scavenger. Fifty milligrams of peptide resin can be deprotected with 4–5 ml cleavage cocktail. 41

E. Kaiser, R. L. Colescott, C. D. Bossinger, and P. I. Cook, Anal. Biochem. 34, 595–598 (1970). 42 T. Vuljanic, K.-E. Bergquist, H. Clausen, S. Roy, and J. Kihlberg, Tetrahedron 52, 7983–8000 (1996). 43 C. A. Guy and G. B. Fields, Methods Enzymol. 289, 67–83 (1997).

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Postcleavage Work Up

After the cleavage has been completed, the resin should be washed two or three times with 2–3 ml portions of neat TFA to extract all peptide from the resin. The combined TFA cleavage solutions are concentrated under a stream of nitrogen and the peptide precipitated with cold diethyl ether. The peptide is collected by centrifugation, redissolved in a small amount of acetic acid and again precipitated with diethyl ether. Several washes are necessary to remove scavengers and cleaved off protecting groups. Finally, the peptide is lyophilized from acetic acid or acetic acid/water mixtures and should be obtained as white powder. Phosphopeptides

Phosphopeptides can be synthesized like nonphosphorylated peptides, except that a phosphorylated tyrosine, threonine, or serine building block is incorporated into the peptide. Phosphorylated building blocks incorporate usually without major difficulties in the peptide, but sometimes slow down the coupling of the subsequent amino acid. It is therefore necessary to control the completeness of the coupling reactions carefully. N-terminally Modified (Phospho)peptides

An N-terminal modification can be easily achieved by coupling any carboxylic acid to the free amino terminus of the last amino acid. The most popular N-terminal modification is an N-terminal acetyl group, but any compound with reactivity towards primary amino groups, such as sulfonyl chlorides, aldehydes, or isothiocyanates can be conjugated as well. When choosing an N-terminal modification one has to consider its stability towards TFA during deprotection/cleavage of the peptide. Phage Display Libraries Since in vivo substrates for protein phosphatases are proteins, it would be desirable to assay combinatorial libraries of phosphorylated proteins. However, chemical synthesis of peptides is often restricted by practical considerations to fewer than 50 residues. Synthetic peptide libraries typically contain peptide shorter than 20 residues. Phage display technology on the other hand allows presentation of libraries of recombinant proteins on the surface of phages.44,45 The unique advantage of phage display protein 44

A. Pini, F. Viti, A. Santucci, B. Carnemolla, L. Zardi, P. Neri, and D. Neri, J. Biol. Chem. 34, 21769–21776 (1998). 45 G. P. Smith and V. A. Petrenko, Chem. Rev. 97, 391–410 (1997).

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libraries is that randomized residues are displayed within the framework of a large protein, which is particularly relevant when protein–protein interactions are addressed. The diversity of phage display libraries is restricted, due to limitations in transformation efficiency of bacterial host cells, to typically less than seven randomized amino acids.46,47 This means that chemically synthesized peptide libraries can be significantly larger than phage display libraries. In addition, the display of phosphorylated proteins on phage particles presents a further challenge and has serious limitations. Examples for the use of phage displayed phospho- and cyclic peptides for the characterization of SH2 domains can be found in Gram et al.,48 Dente et al.,49 and Olingo et al.50 Library Assays and Screening Strategies

Once a library of compounds is obtained one has to screen the library for its activity towards its intended target. The assay strategies used to probe protein phosphatases with a library of compounds should be chosen carefully in order to get results as reliable and useful as possible. Generally, the screening conditions influence the selection process and the results of the screening (you get what you screen for). Three basic features can be easily assayed: phosphatase activity, the inhibition of phosphatase activity, and the binding of compounds to a phosphatase. In general, one should screen under conditions as close as possible to the question under investigation. For example, if one wishes to assess the substrate preference of a PTP, then one should try to develop an assay where indeed catalytic activity is measured, rather than just binding of a potential substrate to the PTP. The following paragraphs present examples for methods that have been used to assay the activity, inhibition, and binding properties of PTPs. Phosphatase Activity

Assays measuring the catalytic activity of a phosphatase towards a library of substrates allow the assessment of the substrate preference of the phosphatase. Although results obtained from analysis of phosphorylated 46

E. R. Zabarovsky and G. Winberg, Nucleic Acid Res. 18, 5912 (1990). D. M. Heery and L. K. Dunican, Nucleic Acid Res. 17, 8006 (1989). 48 H. Gram, R. Schmitz, J. F. Zuber, and G. Baumann, Eur. J. Biochem. 246, 633–637 (1997). 49 L. Dente, C. Vetrani, A. Zucconi, G. Pelicci, L. Lanfrancone, P. G. Pelicci, and G. Cesareni, J. Mol. Biol. 269, 694–703 (1997). 50 L. Oligino, F.-D. Lung, L. Sastry, J. Bigelow, T. Cao, M. Curran, T. R. Burke, S. Wang, D. Krag, P. P. Roller, and C. R. King, J. Biol. Chem. 272, 29046–29052 (1997). 47

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peptides (e.g., optimal or consensus motifs) may not always have a direct physiological relevance, they are valuable for initial characterization of phosphatase active site and development of potent and specific inhibitors. PTP activity can be followed by either substrate consumption or product formation and both can be measured by either continuous or quench methods.51 Quenched methods require the termination of the assay at a certain time point during the course of the reaction. The dephosphorylation of the substrate is stopped and the relative concentration of substrate and/or product is determined. The concentrations or ratios between phosphorylated/de-phosphorylated peptides can be determined by methods such as HPLC, mass spectrometric analysis,52 amino acid analysis,53 gel electrophoresis54 or immunological methods using phosphotyrosine specific antibodies.55 The release of phosphate ions as the result of the phosphatase activity can be followed using chromogenic assays.56,57 Quench methods are relatively laborious because they require several pipetting steps for addition of reagents, yield relatively few data points and do not allow real time analysis of the dephosphorylation reaction. Continuous methods are clearly preferable over quench methods whenever possible. For the analysis of PTP activity a very convenient assay takes advantage of the change in fluorescence of (phospho)tyrosine residues upon dephosphorylation.58 The assay is noninvasive, requires only small amounts of enzyme and substrate. Most importantly it allows real time analysis and yields many data points from a single experiment. Whenever possible, a noninvasive, continuous, real time assay should be used.

Protocol for Continuous Analysis of Phosphotyrosine Peptide Dephosphorylation by Fluorescence Spectrometry51,58

PTP-catalyzed hydrolysis of phosphotyrosine containing peptides is monitored by following the increase in tyrosine fluorescence at 305 nm with excitation at 280 nm. When the substrate concentration is low [S] < < Km, the 51

D. F. McCain and Z.-Y. Zhang, Methods Enzymol. 345, 507–518 (2002). P. Wang, H. Fu, D. F. Snavley, M. Freitas, and D. Pei, Biochemistry 41, 6202–6210 (2002). 53 J. A. Cooper, B. M. Sefton, and T. Hunter, Methods Enzymol. 99, 387–402 (1983). 54 M. L. Sohaskey and J. E. Ferrell, Mol. Biol. Cell 10, 3729–3743 (1999). 55 X. Espanel, M. Huguenin-Reggiani, and R. Hooft van Huijsduijnen, Protein Sci. 11, 2326–2334 (2002). 56 P. A. Lanzetta, L. J. Alvarez, P. S. Reinach, and O. A. Candia, Anal. Biochem. 100, 95–97 (1979). 57 M. J. Black and M. E. Jones, Anal. Biochem. 135, 233–238 (1983). 58 Z.-Y. Zhang, D. Maclean, A. M. Thieme-Sefler, R. W. Roeske, and J. E. Dixon, Anal. Biochem. 211, 7–15 (1993). 52

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Michaelis-Menten equation reduces to a first order reaction with respect to [S]: v ¼ kcat =Km ½S½E:

ð1Þ

For PTP1B, enzyme assays are performed at the following conditions: Buffer: 50 mM 3,3-dimethylglutarate, pH 7.0, 1 mM EDTA, 1 mM DTT, NaCl to an ionic strength of 150 mM; concentration of peptide substrate: 1–10 M; assay temperature: 25–37
C; final PTP1B concentration: 5–100 nM. Protocol for Quenched Assay Analysis of Phosphate Release from Ser/Thr Phosphorylated Peptides57

Measuring the initial rate of phosphatase activity at different substrate concentrations under steady state conditions allows acquisition of kinetic parameters using the Michaelis-Menten equation. This requires that the substrate concentration is much higher than the enzyme concentration and that the substrate concentration decreases only by about 10% during the course of the assay. The substrate peptide is dissolved in a buffer of choice at several concentrations. If kcat/Km is desired, then substrate concentrations should be < 0.1Km. If both Kcat and Km are desired, then substrate concentrations between 0.2Km and 5Km are appropriate. The reaction is initiated by adding the phosphatase to the substrate and the reaction is stopped by adding the quenching solution after a specific amount of time has elapsed. Reagents for the determination of phosphate are added and the amount of inorganic phosphate determined. Solution A: 16 mM ammonium molybdate in water; Solution B: 0.8 M ascorbic acid in 50% (w/v) trichloroacetic acid, made fresh and kept between 0 and 4
C; Solution C: 70 mM trisodium citrate, 0.1 M sodium arsenite in 2% (v/v) acetic acid; Solution D: 10% (w/v) trichloroacetic acid in water. Assay

The phosphatase assay reaction is done in 200 l volumes and the reaction is quenched by addition of 100 l of solution D. Solutions A and B are mixed 2 : 3 and 250 l of the A/B mixture is added to the quenched assay solution. Solution C (500 l) is added after 2 min. The final color develops within 5 min and is stable for several hours. Absorbance is

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measured at 700 nm and the amount of released phosphate determined by comparison to a standard calibration curve.

Inhibition of PTP Activity

In principle, the same methods suitable for measuring PTP activity can be used to measure the inhibition of phosphatase activity. However, inhibition of PTP activity is actually easier to measure than the activity towards a substrate library, because one has the freedom to choose a reference substrate. This reference substrate can be designed such that it has properties which are compatible with a continuous assay. Popular and commercially available chromogenic substrates (e.g., Molecular Probes) for continuous phosphatase assays are p-nitrophenol phosphate (pNPP), 5-bromo-4-chloro-3-indolyl phosphate (BCIP) and the fluorogenic substrates fluorescein diphosphate (FDP), dimethylacridinone phosphate (DDAO) and, 4-methylumbelliferyl phosphate (MUP). In addition, phosphotyrosine or phosphotyrosine containing peptides can be used as reference substrates. A detailed description of assays for PTP activity can be found in a recent review.51

Binding of Library Compounds to Protein Phosphatases

Binding of library compounds to protein phosphatases is often used as a substitute assay for the direct measurement of substrate turnover or inhibition of activity. The assumption behind screening for binding rather than catalytic activity is that the binding of a compound into the active site is prerequisite for catalytic processing of a substrate, as well as for the action of competitive inhibitors. Binding of a ligand into the active site of a protein phosphatase without catalytic processing (and subsequent dissociation of the reaction products) can be achieved by either disabling the phosphatase by mutagenesis of residues crucial for catalysis, or by employing mimics of phosphotyrosine/-threonine/-serine which are stable towards hydrolysis by the enzyme. Both approaches are frequently used, inactivation of PTPs can be achieved by mutation of the catalytically important residues (e.g., Asp181, Cys215, or Gln262, in PTP1B)59–61 and several hydrolytically stable 59

A. J. Flint, T. Tiganis, D. Barford, and N. K. Tonks, Proc. Natl. Acad. Sci. U.S.A. 94, 1680–1685 (1997). 60 K. L. Milarski, G. Zhu, C. G. Pearl, D. J. McNamara, E. M. Dobrusin, D. MacLean, A. Thieme-Sefler, Z.-Y. Zhang, T. Sawyer, S. J. Decker, J. E. Dixon, and A. R. Saltiel, J. Biol. Chem. 268, 23634–23639 (1993). 61 L. Xie, Y.-L. Zhang, and Z.-Y. Zhang, Biochemistry 41, 4032–4039 (2002).

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phosphotyrosine analogues have been developed and are commercially available. Assays based on detecting ligand binding into the catalytic active site select ligands which show tight binding (low Kd) to the protein. Since substrate specificity is determined by kcat/Km, not by affinity alone, the binding assays are therefore not well suited to characterize substrate properties of PTPs. However, tight binding into an enzyme’s active site is a main feature of many inhibitors and it is reasonable to assume that a tightly binding compound would inhibit PTP activity through competition with the substrate for access to the active site. Peptides containing nonhydrolyzable phosphotyrosine derivatives or other phosphotyrosine mimics are structurally targeted into the active site and function often as pseudosubstrates through competitive inhibition. Binding of library compounds to a PTP can be assayed in several ways, either with both partners in solution or with one of them immobilized on a solid support. For instance, a library immobilized on a membrane sheet can be used directly in a Western-blot like assay. The PTP can be detected in several ways, either by labeling the PTP (radioactivity, fluorescence label) or through immunological methods (antibodies, peptide tag, fusion protein, etc.). In addition, it is also possible to screen peptide libraries for ligands which do not bind into the active site of the enzyme and therefore do not exhibit inhibitory activity or substrate properties.62

Deconvolution of Library Screenings

The deconvolution of data from combinatorial library screenings is most important and the deconvolution strategy has to be considered prior to library synthesis. When it comes to the deconvolution of library screening results, one is confronted with two unrelated problems. The first concerns the identity of the compounds selected during the screening assay, and the second is how to deduce a consensus substrate sequence or optimal structure of an inhibitor from those screening results. The identity of a compound is easily determined when its entire synthetic history is known. This history can be recorded either directly by readable tags or labels, or by encoded tags or labels. Examples for easily

62

S. Vetter, High-affinity non-inhibitory ligands for alkaline phosphatase revealed from a restricted heptapeptide library, in ‘‘Solid Phase Synth. Comb. Libr.’’ (R. Epton, ed.), pp. 407–410. Mayflower Science Ltd, Kingswinford, UK, 1999.

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readable tags or labels are radio frequency tags63–65 or, color encoded tags for synthesis performed on pins, direct lab journal entries for small libraries synthesized in cartridges or tea bags, or simply the position (i.e., X–Y coordinates) of a library synthesized as a two-dimensional array on a membrane sheet or a multiwell plate. Directly readable tags are only applicable for relatively small libraries, because they obviously require individual tracking of single compounds during each synthesis step, which becomes unpractical when very large libraries (> 105 members) are synthesized. Encoded tags or labels are cosynthesized during the synthesis of the library and can later be read by specialized techniques. Examples for encoded tags are cosynthesized peptides,66 oligonucleotides,67 mass-spec tag,68 inert gas chromatographic tag69 or fluorescence labels.70 These methods are typically not necessary for peptide libraries, because their amino acid sequences can be identified by direct peptide sequencing, however, they are very useful for N-terminally modified peptides or small organic molecules. Libraries, which cannot be tagged or labeled, such as libraries synthesized by mixture coupling, can be deconvoluted through positional scanning. Deconvolution by positional scanning requires the creation of one sub-library per diversity element (e.g., amino acid) and position. In each sub-library one position is structurally defined, while all other variable positions are randomized. For example, to scan all positions in a hexapeptide library built from 20 amino acids one has to synthesize 6  20 ¼ 120 individual sub-libraries. Positional scanning limits the screening effort, because each sub-library has to be assayed only once to establish a ranking of the most favorable substitution(s) in each position. In theory, the optimal substrate/inhibitor can then be deduced by combining the best substitution for each position into a single structure/sequence. However, positional scanning does not allow the identification of individual structures or sequences and neglects synergistic and allosteric effects of neighboring diversity elements (amino acid side chains). 63

R. W. Armstrong, P. A. Tempest, and J. F. Cargill, Chimia 50, 258–260 (1996). K. C. Nicolaou, X. Y. Xiao, Z. Parandoosh, A. Senyei, G. J. Parke, and M. P. Nova, Angew. Chem. Int. Ed. Engl. 34, 2289–2291 (1995). 65 E. J. Moran, S. Sarshar, J. F. Cargill, M. M. Shahbaz, A. Lio, A. M. M. Mjalli, and R. W. Armstrong, J. Am. Chem. Soc. 117, 10787–10788 (1995). 66 Y. W. Cheung, C. Abell, and S. Balasubramanian, J. Am. Chem. Soc. 119, 9568–9569 (1997). 67 J. Nielsen, S. Brenner, and K. D. Janda, J. Am. Chem. Soc. 115, 9812–9813 (1993). 68 H. M. Geysen, C. D. Wagner, W. M. Bodnar, C. J. Markworth, G. J. Parke, F. J. Schoenen, D. S. Wagner, and D. S. Kinder, Chem. Biol. 3, 679–688 (1996). 69 M. H. J. Ohlmeyer, R. N. Swanson, L. W. Dillard, J. C. Reader, G. Asouline, R. Kobayashi, M. Wigler, and W. C. Still, Proc. Natl. Acad. Sci. U.S.A. 90, 10922–10926 (1993). 70 X. Lui, L. H. Takahashi, W. L. Fitch, G. Rozing, C. Bayle, and F. Couderc, J. Chromatography 924, 323–329 (2001). 64

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Positional scanning can also be applied to libraries which have been synthesized in the ‘‘one-bead one-compound’’ format. For instance, pools of beads or soluble peptides can be sequenced and the relative amino acid frequencies in each position determined.71 Positional scanning is generally a valuable tool to identify substitution patterns and trends. If a library has been synthesized such that it is possible to identify individual compounds, for example in a ‘‘one-bead one-compound’’ library, then it would be most rigorous to assay each compound individually. However, for large libraries this becomes a major effort and it will be necessary to assay pools of compounds or the entire library at once. In such cases one can either choose to deconvolute pools of compounds by positional scanning or to pick a representative number of individual compounds (beads) and to extract consensus motifs from these individual sequences. The analysis of individual compounds and subsequent deduction of consensus sequences is preferable compared to pool sequencing, because all chemical and physical features of each compound are accessible and cooperativity effects between side chains may be observable. To obtain reliable data, it is often necessary to sequence a relatively large number of individual compounds. A conceptual road map through the many possible avenues for the characterization of protein phosphatase using combinatorial chemistry and library methods is shown in Fig. 1. Obviously, a multitude of possible combinations of library design/synthesis, assay strategies and deconvolution methods can be utilized. A number of explicit examples are indicated in the figure by subscribed numbers, which refer to the examples briefly explained and commented on below. Example 155

A library of phosphotyrosine containing peptides was synthesized by SPOT technique on paper sheets as solid support in a 20  20 spot array. Each spot contained a mixture of 400 peptide sequences of the format AABX1(pY)X2BAAA. The positions X1 and X2 represent one of 20 amino acids and correspond to the two dimension of the 20  20 spot array. The positions B were randomized by coupling a mixture of 20 amino acids. The paper bound library was assayed by incubation with PTP1B in order to identify preferred substrate peptides (1a in Fig. 1). Dephosphorylation of peptides was measured by detecting the amount of phosphotyrosine 71

Z. Songyang, S. E. Shoelson, M. Chandhuri, G. Gish, T. Pawson, D. M. Haser, F. King, T. Roberts, S. Ratnofsky, R. J. Lechleider, B. G. Neel, R. B. Birge, J. E. Fajardo, M. M. Chou, B. Hanafusa, L. C. Schaffhausen, and L. C. Cantley, Cell 72, 767–778 (1993).

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FIG. 1. Road Map through the possibilities to characterize protein phosphatases with combinatorial and library methods. The number under individual decision points refer to the number of the examples given in the text.

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remaining in each peptide spot using a mixture of anti-pTyr antibodies. In an unrelated experiment (1b in Fig. 1) the binding of the catalytically inactive mutant PTP1B-D181A to the peptides was measured by using the radiolabeled phosphatase for radiometric imaging. Deconvolution was simple in this case, because the position of each spot on the membrane encodes the amino acid substitutions in positions X1 and X2. Interestingly, results from the PTP1B-D181A binding assay and catalytic dephosphorylation assay did not identify the same peptides as optimal binder and substrate, respectively. In addition, the anti-pTyr antibody mixture used to detect the extent of dephosphorylation failed completely to bind to a subset of 20 peptides containing a cysteine residue in X2 position. This underlines the difficulty to reliably quantify the extent of de-/phosphorylation in diverse peptides using antibodies.

Example 272

In this example a library of 153 nine-residue peptides were synthesized which contained a central pTyr residue, only one defined position, and alanine residues in all other seven positions. Each peptide was assayed in solution for dephosphorylation by PTP1B using a continuous fluorescence based assay and kcat/Km values were determined for each peptide. The deconvolution was done by identifying the most favorable substitution in each position and combining them into a single consensus sequence. This consensus peptide was found to be a highly active substrate (kcat/ Km ¼ 2.2  107 M1 s1). More importantly, since this strategy uses single peptide substrates of defined purity it was possible to obtain reliable kinetic data for each substitution and to reveal several theretofore unknown features of PTP1B substrate specificity.

Example 362

This is an example for the identification of peptides binding to a phosphatase but not targeting the catalytic active site. A library of nonphosphorylated peptides was synthesized in the ‘‘one-bead onecompound’’ format. The library was assayed for peptides with binding activity to alkaline phosphatase and phosphatase binding beads/peptides were identified based on the catalytic activity of the bound enzyme. BCIP (5-bromo-4-chloro-3-indolyl phosphatase), which formed a colored, 72

S. W. Vetter, Y.-F. Keng, D. S. Lawrence, and Z.-Y. Zhang, J. Biol. Chem. 275, 2265–2268 (2000).

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insoluble product upon dephosphorylation was used as a substrate. Colored beads were isolated from the library and deconvolution was achieved by peptide sequencing directly from single beads. Comparing the individual sequences suggested a consensus peptide sequence for recognition of alkaline phosphatase. Example 473

A specific PTP1B inhibitor was identified from a library of modified peptides. The peptides were synthesized in parallel and were modified at their N-termini by coupling of an unnatural amino acid followed by a carboxylic acid. In addition, to target the peptides into the active site of PTP1B they contained a pTyr residue or nonhydrolyzable pTyr mimic. The screening of the library was done after release of the compounds into solution, using a competition binding ELISA assay. A reference peptide with high binding affinity for PTP1B was immobilized on multi-well plates and a catalytically inactive GST-PTP1B-C215S mutant fusion protein was bound to the reference peptide. Addition of the soluble library compounds led to partial release of the GST-PTP1B-C215C fusion protein from the immobilized reference ligand due to competition binding. The amount of retained PTP1B was measured by ELISA using anti-GST antibodies. Deconvolution was trivial, because each tested library compound was known. This method identified a novel PTP1B inhibitor with high potency (Ki ¼ 2.4 nM) and selectivity. Example 565

Radio frequency tagging is another method to record the synthetic history and subsequently structure of library compounds. This method was used to tag a library of 125 tripeptide-substituted cinnamic acids. The library was synthesized on capsules containing the radio frequency transponder and then each transponder was scanned, distributed into an individual well of a multi-well plate and the compound was cleaved from the solid support. Thus each well contained one chemical compound with known chemical structure. The library was then screened for inhibition of PTP1B activity using p-nitrophenol phosphate as a reference substrate.

73

K. Shen, Y.-F. Keng, L. Wu, D. S. Lawrence, and Z.-Y. Zhang, J. Biol. Chem. 276, 47311– 47319 (2001).

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Example 674

Affinity selection of peptides with PTP1B binding affinity from a mixture of peptides, followed by deconvolution by mass spectrometry has also been demonstrated. A peptide library based on the known PTP1B substrate peptide (DADEpYL) derived from the EGF receptor was synthesized using a nonhydrolyzable pTyr mimic (F2Pmp) and contained one randomized position (X). The library was incubated with PTP1B and PTP1B-peptide complexes separated from nonbinding peptides by gel filtration. Mass spectrometry was then used to identify the relative frequency of individual amino acids in the randomized position X. All five positions of the parent peptide were scanned in this fashion and a consensus sequence obtained by combining the most frequent substitution in each position. Example 775

A library screening based on inhibition of PTP1B activity used a peptide library containing a nonhydrolyzable pTyr mimic (mY) and a quench assay to measure the amount of released phosphate from a reference peptide. The library DXXX(mY)LIP was synthesized in the ‘‘one-bead one-compound’’ format and three sets of 20 sub-libraries were created for each defined position. Each of the 3  20 ¼ 60 sub-libraries was incubated with a reference phosphotyrosine peptide substrate and PTP1B was added to initiate substrate dephosphorylation. The degree of PTP1B inhibition was measured using quenched point assay. Deconvolution and assembly of a consensus sequence was done by combining the substitution giving rise to the strongest inhibition of PTP1B activity in each position. Example 866

The substrate preferences of the PTP LAR were assayed by dephosphorylation of substrate peptides bound to resin beads. The phosphotyrosine containing library was synthesized in the ‘‘one-bead onecompound’’ format and an encoding tag was created by cosynthesizing a second set of peptides which had identical amino acid sequence, but

74

G. Huyer, J. Kelly, J. Moffat, R. Zamboni, Z. Jia, M. J. Gresser, and C. Ramachandra, Anal. Biochem. 258, 19–30 (1998). 75 M. C. Pellegrini, H. Liang, S. Mandiyan, K. Wang, A. Yuryev, I. Vlattas, T. Sytwu, Y.-C. Li, and L. P. Wennogle, Biochemistry 37, 15598–15606 (1998).

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contained a glycine residue, instead of a pTyr residue in this position. This was achieved by coupling a mixture of 70% pTyr and 30% glycine. In addition, the N-terminal amino group of the peptides remained protected with the Fmoc-group during the assay. The assay itself was based on preferential proteolytic cleavage of peptides containing tyrosine residues compared to peptides containing phosphotyrosine residues by the protease -chemotrypsin. The bead bound peptide library was first incubated with PTP LAR and then with -chemotrypsin. Dephosphorylated peptides were cleaved by -chemotrypsin C-terminal to the Tyr residue resulting from the dephosphorylation of the pTyr residue in substrate peptides. The newly created free amino termini were detected using an amino group specific fluorescent label. Beads showing the strongest fluorescence were isolated and the Fmoc-group was removed from the N-terminus of the encoding peptide strands, which were then sequenced by Edmann degradation. The part of the substrate peptide remaining on the bead did not interfere with the sequencing of the encoding strand, because the N-terminus was blocked by the fluorescence label. The encoding tag allowed to obtain the sequence of the original substrate peptide, because the glycine residue had prevented proteolytic cleavage. This method, while conceptually interesting, has serious practical limitations because aromatic residues and lysine residues cannot be used for library construction.

General Advice

1.

2.

3.

4.

Chemical synthesis is the most useful route to peptide libraries in general and phosphopeptide libraries in particular. Chemical synthesis is the only suitable way to libraries of peptide derivatives and peptide analogues. Phage display libraries are great for the display of proteins, but not suitable for work with phosphopeptides. ‘‘One-bead one-compound’’ libraries are often easier to synthesize and can be better characterized analytically than libraries obtained through reaction of reagent mixtures. The assay used should be designed such that it addresses the question under investigation. If the interest is substrate specificity, then the assay should be based on substrate turnover, not on pseudo-substrate binding or binding to a catalytically inactive mutant (remember: you get what you screen for). If individual compounds are identified, then as many as possible should be looked at in order to recognize shared properties or families of compounds with common features. If positional

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scanning is used, then one has to keep in mind that cooperativity effects between individual positions can be missed. Results from library screenings always have to be confirmed by resynthesizing a series of possible consensus compounds and reassaying them.

Acknowledgments Z.-Y. Z is supported in part by National Institutes of Health Grant AI48506, the G. Harold and Leila Y. Mathers Charitable Foundation, and the Irma T. Hirschl Foundation.

[20] Activity of PP2C is Increased by Divalent Cations and Lipophilic Compounds Depending on the Substrate By JOSEF KRIEGLSTEIN, DAGMAR SELKE, ALEXANDER MAAßEN, and SUSANNE KLUMPP Introduction

The - and -isozymes of serine/threonine protein phosphatase type-2C (PP2C) belong to the phosphatases of the first generation, described side by side with PP1, PP2A and PP2B in the 1980s.1 In the following text we will refer to PP2C , fully aware that in the meantime more isozymes than  and contributed to build up a larger PP2C-family. From the very beginning, PP2C was described as an enzyme requiring Mg2 þ - or Mn2 þ -ions for activity.2 Later on molecular biology revealed that PP2C belongs to a gene family distinct from PP1, PP2A and PP2B.3 Since then the PP2C enzymes continue to be strangers in the field: (i) activity of PP2C is not inhibited by okadaic acid and related compounds; (ii) neither targeting nor regulatory subunits are known for PP2C; and (iii) an inhibitor for PP2C is still not available. Furthermore, there are only a few articles discussing possible regulatory mechanisms for PP2C. Long ago dimerization was suggested;4 1

T. S. Ingebritsen and P. Cohen, Protein Phosphatases: Properties and Role in Cellular Regulation, Science 221, 331–338 (1983). 2 C. H. McGowan and P. Cohen, Protein phosphatase-2C from rabbit skeletal muscle and liver: an Mg2 þ -dependent enzyme, Methods Enzymol. 159, 416–426 (1988). 3 P. T. W. Cohen, Nomenclature and Chromosomal Localization of Human Protein Serine/ Threonine Phosphatase, Adv. Prot. Phosphatases 8, 371–376 (1994). 4 G. Mieskes and H.-D. So¨ling, On the 6-phosphofructo-1-kinase phosphatase activity of protein phosphatase 2C and its dimeric nature, FEBS Lett. 181(1), 7–11 (1985).

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5.

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scanning is used, then one has to keep in mind that cooperativity effects between individual positions can be missed. Results from library screenings always have to be confirmed by resynthesizing a series of possible consensus compounds and reassaying them.

Acknowledgments Z.-Y. Z is supported in part by National Institutes of Health Grant AI48506, the G. Harold and Leila Y. Mathers Charitable Foundation, and the Irma T. Hirschl Foundation.

[21] Activity of PP2Cb is Increased by Divalent Cations and Lipophilic Compounds Depending on the Substrate By JOSEF KRIEGLSTEIN, DAGMAR SELKE, ALEXANDER MAAßEN, and SUSANNE KLUMPP Introduction

The - and b-isozymes of serine/threonine protein phosphatase type-2C (PP2C) belong to the phosphatases of the first generation, described side by side with PP1, PP2A and PP2B in the 1980s.1 In the following text we will refer to PP2Cb, fully aware that in the meantime more isozymes than  and b contributed to build up a larger PP2C-family. From the very beginning, PP2C was described as an enzyme requiring Mg2 þ - or Mn2 þ -ions for activity.2 Later on molecular biology revealed that PP2C belongs to a gene family distinct from PP1, PP2A and PP2B.3 Since then the PP2C enzymes continue to be strangers in the field: (i) activity of PP2C is not inhibited by okadaic acid and related compounds; (ii) neither targeting nor regulatory subunits are known for PP2C; and (iii) an inhibitor for PP2C is still not available. Furthermore, there are only a few articles discussing possible regulatory mechanisms for PP2C. Long ago dimerization was suggested;4 1

T. S. Ingebritsen and P. Cohen, Protein Phosphatases: Properties and Role in Cellular Regulation, Science 221, 331–338 (1983). 2 C. H. McGowan and P. Cohen, Protein phosphatase-2C from rabbit skeletal muscle and liver: an Mg2 þ -dependent enzyme, Methods Enzymol. 159, 416–426 (1988). 3 P. T. W. Cohen, Nomenclature and Chromosomal Localization of Human Protein Serine/ Threonine Phosphatase, Adv. Prot. Phosphatases 8, 371–376 (1994). 4 G. Mieskes and H.-D. So¨ling, On the 6-phosphofructo-1-kinase phosphatase activity of protein phosphatase 2C and its dimeric nature, FEBS Lett. 181(1), 7–11 (1985).

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then phosphorylation of PP2C by casein kinase II was reported;5 and recently inactivation by hydrogen peroxide for the enzyme from plants has been described.6 Inhibition by calcium ions also was observed in several cases, but the concentration was too high to account for physiological relevance.7–9 The requirement of Mg2 þ - or Mn2 þ -ions for PP2C activity still holds true. No other mono- or divalent cation was found capable of supporting PP2C activity. Therefore, PP2C assays are routinely performed in the presence of 10–20 mM Mg2 þ . This is applied for peptide substrates as well as for phosphoradiolabeled protein substrates. The physiological concentration of free Mg2 þ , however, is considered to be 0.5–1.5 mM only. Under these conditions PP2C activity is approximately 10% of the activity measured at 20 mM Mg2 þ . In 1998, we discovered that PP2C activity is highly active at physiological Mg2 þ concentrations provided certain unsaturated lipophilic compounds were present.10 The concentration of the lipids necessary so far exceeds the concentration of free fatty acids under physiological conditions. However, those assays were performed using casein as a substrate, and casein itself is known to bind free fatty acids, thus reducing the effective concentration to a much lower value. Phosphopeptides are often used for handy and sensitive assays of protein phosphatase activities. A relatively low sensitivity of the subsequent colorimetric determination of inorganic phosphate is disadvantageous and even detrimental for measuring PP2C activity in crude tissue extracts. Phosphopeptides are very useful, however, for quickly detecting and quantifying at least partially purified phosphatases. But there are clear limitations. Using phosphopeptides in the context of regulatory mechanisms or searching for activators and inhibitors (e.g., via high throughput screenings) may be misleading.

5

T. Kobayashi et al., Phosphorylation of Mg2 þ -dependent protein phosphatase  (Type 2C) by casein kinase II, Biochem. Biophys. Res. Commun. 195(1), 484–489 (1993). 6 M. Meinhard and E. Grill, Hydrogen peroxide is a regulator of ABI1, protein phosphatase 2C from Arabidopsis, FEBS Lett. 508, 443–446 (2001). 7 M. D. Pato and E. Kerc, Regulation of smooth muscle phosphatase-II by divalent cations, Mol. Cell. Biochem. 101, 31–41 (1991). 8 Y. Wang et al., A Mg2 þ -dependent, Ca2 þ -inhibitable Serine/Threonine Protein Phosphatase from Bovine Brain, J. Biol. Chem. 270(43), 25607–25612 (1995). 9 S. Klumpp et al., Protein phosphatase type-2C isozyme present in vertebrate retinae: purification, characterization, and localisation in photoreceptors, J. Neurosci. Res. 51, 328–338 (1998). 10 S. Klumpp, D. Selke, and J. Hermesmeier, Protein phosphatase type 2C active at physiological Mg2 þ : stimulation by unsaturated fatty acids, FEBS Lett. 437, 229–232 (1998).

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Determination of PP2C-activity [32P]Casein as a Substrate

Phosphorylation of casein and dephosphorylation by PP2C were performed as previously described.11 In brief, the substrate is prepared by phosphorylation using PKA (Sigma, P-4890) and [ -32P]ATP. Gel filtration on Sephadex G-50 is used to separate the labeled protein from unincorporated nucleotides. Activity of 5–15 ng purified recombinant PP2C is measured (30 l, 30
) in 33 mM Tris–HCl, pH 7.0, 1.3 mg/ml bovine serum albumin, 0.1% 2-mercaptoethanol, 1 or 10 mM MgCl2 as indicated, and 1 M [32P]casein (1  106 cpm/ml). Reactions are terminated by trichloroacetic acid, centrifuged, and the supernatant analyzed for [32P] content. BAD Phosphorylated at Ser-155 as a Substrate

Purified GST-BAD (12 g) is incubated in 25 mM Tris–HCl, pH 7.5, 10 mM MgCl2, 1 M ATP including 5 Ci [ -32P]ATP and 100 ng PKA in a total volume of 12 l at 37
for 30 min. Phospho-BAD is then separated from unincorporated ATP by the use of centri-SEP columns (750g for 2 min at room temperature). Dephosphorylation reactions are carried out with 1.2 g GST-phospho-BAD, 0.3 g PP2Cb, 50 mM Tris–HCl, pH 7.5, 1 mM Mg2 þ , 500 M oleic acid or 10% DMSO in a total volume of 15 l. Reactions are terminated after incubation for 30 min at 37
by the addition of 5 l sample buffer (15 mM Tris–HCl, pH 6.8, 4% SDS, 2% 2-mercaptoethanol, 10% sucrose, 8 M urea, 10 mM EDTA, 0.06% bromophenol blue). Phosphorylation and dephosphorylation of GST-BAD is analyzed after 12.5% SDS-PAGE by autoradiography. A modified version of the phosphorylation and dephosphorylation of BAD has been reported earlier.12 Phosphopeptides as Substrates

Phosphopeptides are dissolved in 50 mM Tris–HCl, pH 7.0, 0.1 mM EGTA to give a 1 mM stock solution and stored at 20
. Dephosphorylation reactions are performed in microplates (30 l, 30
, 10 min) containing 50 mM Tris–HCl, pH 7.0, 1 or 10 mM Mg2 þ as indicated, 11

S. Klumpp and D. Selke, Purification and Characterization of Protein Phosphatase Type 2C in Photoreceptors, Methods Enzymol. 315, 570–578 (2000). 12 S. Klumpp, D. Selke, and J. Kriegstein, Protein phosphatase type 2C dephosphorylates BAD, Neurochem. Inter. 42(7), 555–560 (2003).

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50 ng purified recombinant PP2Cb, 0.2 mM of the phosphopeptide, and terminated by the addition of 30 l of 1.66 M HClO4 and 65 l H2O. The inorganic phosphate released is subsequently quantified via malachite green by a procedure modified from previous reports.13,14 Per well are added 75 l of a premixed acidic ammonium molybdate solution (43 l of 28 mM (NH4)6Mo7O24 in 2.1 M H2SO4 and 32 l of 0.76 mM malachite green in 0.35% Tween-20). After 20 min at 30
OD630 is measured using a microplate reader. The phosphate release is calculated using a calibration curve established with NaH2PO4 (from 0 to 12 nmol NaH2PO4 per well).

Results and Comments General Remarks

For the studies described here we used recombinant and NiNTApurified PP2Cb as phosphatase enzyme source. Since, therefore, other phosphatases are not present, addition of okadaic acid is not necessary. With regard to substrates of PP2C we focused on phosphoproteins and phosphopeptides. Artificial substrates such as para-nitrophenyl phosphate were not examined, mainly because kinetic studies once demonstrated that phosphopeptides are greatly preferred over completely artificial substrates.15 Two decades ago the phosphoprotein substrate used here, [32P]casein, had been well established in the literature as in vitro substrate for PP2C.2 Recently the other protein studied herein, the proapoptotic protein BAD, also had been identified as a substrate of PP2C.12 Phosphorylated synthetic peptides as tools for studying protein phosphatases—including PP2C—have been known for a long time.16 The phosphopeptide substrates used here are derived from AMP-activated protein kinase (FLRpTSCG) and pyruvate kinase (RRPpTVA; T-kemptide). There is one more chapter on PP2C and phosphothreonyl peptide substrates in this volume. The authors would like to refer to Donella-Deana, Boschetti, and Pinna (this volume, Chapter 1, pp. 3–17). Independent of whether phosphopeptides or phosphoproteins are used for determination of PP2C activity the linear range of phosphate release is 13

E. B. Cogan, G. B. Birrel, and O. H. Griffith, A robotics-based automated assay for inorganic and organic phosphates, Anal. Biochem. 271(1), 29–35 (1999). 14 J. D. Mahuren et al., Microassay of phosphate provides a general method for measuring the activity of phosphatases using physiological, nonchromogenic substrates such as lysophosphatidic acid, Anal. Biochem. 298(2), 241–245 (2001). 15 C. C. Fjeld and J. M. Denu, Kinetic analysis of human serine/threonine protein phosphatase 2Calpha, J. Biol. Chem. 274(29), 20336–20343 (1999). 16 L. A. Pinna and A. Donella-Deana, Phosphorylated synthetic peptides as tools for studying protein phosphatases, Biochim. Biophys. Acta 1222, 415–431 (1994).

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restricted to 25–30% of the phosphate content. Assays should be kept within this limit. This is achieved by diluting the PP2C enzyme source as appropriate. The storage temperature of the PP2C substrates is crucial and differs depending on peptide or protein source. Phosphopeptides should be kept frozen, 20
yielding the same recovery as 80
, yet hydrolysis at 4
may be huge. Phosphate release can be as much as 75% within four days at 4
. This is in sharp contrast to the phosphoproteins: [32P]Casein and [32P]BAD can be kept at 4
as a concentrated stock solution with no significant hydrolysis for weeks. Phosphopeptides dose-dependently inhibit dephosphorylation of [32P]casein by PP2C. For instance, addition of 2 mM RRPpTVA or 2 mM FLRpTSCG reduces the dephosphorylation of [32P]casein by 85% and 95%, respectively. For control, one of the unphosphorylated peptides was added instead. It had no effect on the rate of dephosphorylation for [32P]casein by PP2C (data not shown). Differences in the Dependence of Divalent Cations for Activity

As known for many years PP2C activity does require a high concentration of Mg2 þ ions. This has been observed to be independent of whether a phosphoprotein substrate ([32P]casein) or a phosphopeptide substrate FLRpTSCG is used for dephosphorylation by PP2C (Fig. 1A). The dose response curves are quite similar yielding maximal activity of purified recombinant PP2C at 20–30 mM Mg2 þ , or a half maximal effect at about 2–3 mM Mg2 þ , respectively. The activity of PP2C in the presence of Mn2 þ instead of Mg2 þ differs dramatically (Fig. 1B). Using a protein substrate Mn2 þ is as effective as Mg2 þ , yielding the same specific activity of PP2C at an even somewhat lower concentration, such as 10 mM Mn2 þ . In contrast, phosphopeptides cannot undergo dephosphorylation by PP2C in the presence of Mn2 þ . This discrepancy has been seen in various phosphopeptides, suggesting that this is due to the limited number of amino acids rather than to the individual amino acid composition (data not shown). Differences in the Effect of Lipophilic Oxidizable Compounds

With regard to the chemical structure special requirements have to be fulfilled by lipophilic compounds so that activation of PP2C does occur at low and physiological Mg2 þ concentrations:10  

Chain length of more than 15 C-atoms. Special location of the double bond.

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287

FIG. 1. Cation-dependence of purified recombinant PP2Cb using 1 M of the protein [32P]casein (j–j) or 0.2 mM of the peptide FLRpTSCG (s–s) as a substrate. (A) Activity measured in the presence of 0.1–100 mM Mg2 þ . (B) Determination of PP2C activity with the same concentrations of Mn2 þ instead of Mg2 þ .

 

Acidic group (esterified unsaturated fatty acids are no longer active). Cis-configuration (corresponding unphysiological trans-derivatives are not active).

Here we have used oleic acid (18:1 cis-
9) for activation of PP2C. As reported earlier oleic acid stimulates PP2C activity approximately 10-fold at 0.5–1 mM Mg2 þ using [32P]casein as a substrate (Fig. 2A). This is in sharp contrast to the effect of oleic acid on PP2C activity when a phosphopeptide is used as a substrate instead of the phosphoprotein. At 1 mM Mg2 þ oleic acid potently inhibits PP2C acting on the phosphopeptide FLRpTSCG (Fig. 2B). This discrepancy—activation in one case with a protein substrate, inhibition in the other with a peptide substrate—is neither specific for casein nor for the AMPK-peptide. Dephosphorylation of other phosphopeptides, e.g., RRPpTVA, also is inhibited by oleic acid (Fig. 2C) whereas dephosphorylation of yet another protein, [32P]BAD, is once again activated by oleic acid (Fig. 2D). This suggests that the activity of PP2C increases with unsaturated lipophilic compounds specifically towards protein substrates, whereas the dephosphorylation of peptide substrates by PP2C is reduced upon addition of unsaturated lipophilic compounds. At 10 mM Mg2 þ both the activating effect of oleic acid on the dephosphorylation of a protein substrate (Fig. 3A) as well as the inhibitory effect of oleic acid on the dephosphorylation of a peptide substrate (Fig. 3B) are significantly reduced. It is intriguing to observe that both plant and vertebrate PP2C react in response to lipophilic compounds—although in opposite directions. PP2C

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FIG. 2. Effect of oleic acid on the activity of PP2Cb at 0.5–1 mM Mg2 þ depending on whether a phosphoprotein or a phosphopeptide is used as substrate. Oleic acid is dissolved in DMSO. Final concentrations in the assays are 10% DMSO for control, or 500 M oleic acid in 10% DMSO. (A) Dephosphorylation of [32P]casein activated by oleic acid. (B) Dephosphorylation of FLRpTSCG inhibited by oleic acid. (C) Dephosphorylation of RRPpTVA inhibited by oleic acid. (D) Dephosphorylation of [32P]BAD activated by oleic acid (autoradiogram).

activity from plants is inhibited by long-chain polyunsaturated fatty acids using [32P]casein as a substrate, and this inhibition is not dependent upon the Mg2 þ -concentration.17 Structural diversity may account for the opposite response when challenged with fatty acids: Plant PP2C shares only 30% identity with mammalian PP2Cb and it carries an additional NH2-terminal extension with a Ca2 þ /calmodulin-binding site. In conclusion, the maximal activity of PP2Cb is observed with 15 mM Mg2 þ or when lipophilic compounds are present with 1–2 mM Mg2 þ , i.e., lipophilic compounds shift the maximum PP2C activity into a physiological range of Mg2 þ -concentration. The data presented also give a warning to high-throughput-screenings designed to use phosphopeptides as a substrate of PP2C. The identity, hence chemistry, of amino acids surrounding

17

E. Baudouin, I. Meskiene, and H. Hirt, Unsaturated fatty acids inhibit MP2C, a protein phosphatase 2C involved in the wound-induced MAP kinase pathway regulation, Plant J. 20(3), 343–348 (1999).

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FIG. 3. Dephosphorylation of different substrates by PP2Cb in the presence of 10 mM Mg2 þ and 500 M oleic acid. (A) Dephosphorylation of [32P]casein. (B) Dephosphorylation of FLRpTSCG. Note that in contrast to Fig. 2 (0.5–1 mM Mg2 þ only) both the stimulating effect of oleic acid on protein substrates (3A vs 2A) as well as the inhibitory effect of oleic acid on phosphopeptide substrates (3B vs 2B) are greatly reduced when 10 mM Mg2 þ is present.

potential dephosphorylation sites is indicative of whether or not a phosphopeptide is a substrate of PP2C. Those surrounding amino acids, however, are not sufficient in the regulatory studies. The effect of activators and inhibitors of PP2C depends on more than 10–15 amino acids. Data obtained with phosphopeptides may not necessarily reflect the effects on protein substrates.

[21] Regulation of Calcineurin by Oxidative Stress By MANIK C. GHOSH, XUTONG WANG, SHIPENG LI, and CLAUDE KLEE Introduction

Calcineurin (also called protein phosphatase-2B) is a Ca2 þ -dependent serine/threonine protein phosphatase (for recent reviews see Refs. 1–3). Its identification as the target of the immunosuppressive drugs (FK506 and 1

J. Aramburu, A. Rao, and C. B. Klee, in ‘‘Calcineurin: From Structure to Function. Curr. Topics in Cell. Regulation’’ (E. Stadtman and B. Chock, eds.), pp. 237–295. Academic Press, New York, NY, 2000. 2 F. Rusnak and P. Mertz, Physiol. Rev. 80, 1483 (2000). 3 C. S. Hemenway and J. Heitman, Cell Biochem. Biophys. 30, 115 (1999).

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REDOX REGULATION OF CALCINEURIN

FIG. 3. Dephosphorylation of different substrates by PP2C in the presence of 10 mM Mg2 þ and 500 M oleic acid. (A) Dephosphorylation of [32P]casein. (B) Dephosphorylation of FLRpTSCG. Note that in contrast to Fig. 2 (0.5–1 mM Mg2 þ only) both the stimulating effect of oleic acid on protein substrates (3A vs 2A) as well as the inhibitory effect of oleic acid on phosphopeptide substrates (3B vs 2B) are greatly reduced when 10 mM Mg2 þ is present.

potential dephosphorylation sites is indicative of whether or not a phosphopeptide is a substrate of PP2C. Those surrounding amino acids, however, are not sufficient in the regulatory studies. The effect of activators and inhibitors of PP2C depends on more than 10–15 amino acids. Data obtained with phosphopeptides may not necessarily reflect the effects on protein substrates.

[22] Regulation of Calcineurin by Oxidative Stress By MANIK C. GHOSH, XUTONG WANG, SHIPENG LI, and CLAUDE KLEE Introduction

Calcineurin (also called protein phosphatase-2B) is a Ca2 þ -dependent serine/threonine protein phosphatase (for recent reviews see Refs. 1–3). Its identification as the target of the immunosuppressive drugs (FK506 and 1

J. Aramburu, A. Rao, and C. B. Klee, in ‘‘Calcineurin: From Structure to Function. Curr. Topics in Cell. Regulation’’ (E. Stadtman and B. Chock, eds.), pp. 237–295. Academic Press, New York, NY, 2000. 2 F. Rusnak and P. Mertz, Physiol. Rev. 80, 1483 (2000). 3 C. S. Hemenway and J. Heitman, Cell Biochem. Biophys. 30, 115 (1999).

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cyclosporin A) complexed with their respective binding proteins (FKBP12, cyclophilin A) revealed its key role in the Ca2 þ -dependent steps of T cell activation.4 The demonstration that these compounds are specific inhibitors of calcineurin led to the purification of one of its major substrate, the transcription factor, NFAT, and together with the overexpression of a constitutively active derivative of calcineurin helped to reveal the roles of calcineurin in the regulation of cellular processes as diverse as gene expression, ion homeostasis, muscle differentiation, embryogenesis, secretion, and neurological functions.1–6 The Ca2 þ -dependent activity of calcineurin is under the control of two different Ca2 þ -regulated proteins, calcineurin B, an integral subunit of the enzyme, and calmodulin (CaM). Ca2 þ binding to calcineurin B has only a small stimulatory effect on enzyme activity but results in the exposure of the CaM-binding domain of calcineurin A, a prerequisite for calmodulin binding to calcineurin and full activation of its phosphatase activity.7 It is generally accepted that CaM activation is the result of the displacement of the autoinhibitory domain that results in the exposure of the active site of the enzyme.1 The active site of calcineurin, that contains stoichiometric amounts of iron and zinc, was identified as a binuclear (Fe3 þ –Zn2 þ ) center.8,9 In the crystal structure of calcineurin the assigned coordination of Zn2 þ is the same as in the kidney bean (Fe3 þ –Zn2 þ ) acid purple protein phosphatase but the coordinating ligands of Fe3 þ are different.10–12 This difference may reflect a difference in the oxidation state of iron required for activity, which has been the subject of a controversy.13–17 Because of a partial loss of natural metal cofactors during the affinity chromatography step, the purified enzyme has a low specific activity that is stimulated to different extents by exogenous metal ions such as Mn2 þ or Mg2 þ .18 In contrast, the specific activity of calcineurin 4

J. Liu, J. D. Farmer et al., Cell 66, 807 (1991). E. N. Olson and R. S. Williams, Cell 101, 689 (2000). 6 G. R. Crabtree, J. Biol. Chem. 276, 2313 (2001). 7 S.-A. Yang and C. B. Klee, Biochemistry 39, 16147 (2000). 8 M. M. King and C. Y. Huang, J. Biol. Chem. 259, 8847 (1984). 9 L. Yu, A. Haddy, and F. Rusnak, J. Am. Chem. Soc. 117, 10147 (1995). 10 N. Strater, T. Klabunde, P. Tucker, H. Witzel, and B. Krebs, Science 268, 1489 (1995). 11 J. P. Griffith, J. L. Kim et al., Cell 82, 507 (1995). 12 C. R. Kissinger, H. E. Parge et al., Nature 378, 641 (1995). 13 K. F. Qin, S. V. Khangulov, C. Liu, and C. Y. Huang, The FASEB J. 9, A1347 (1995). 14 L. Yu, J. Golbeck, J. Yao, and F. Rusnak, Biochemistry 36, 10727 (1997). 15 X. Wang, V. C. Culotta, and C. B. Klee, Nature 383, 434 (1996). 16 M. C. Ghosh, A. Samouilov, J. Zweier, and C. B. Klee, manuscript in preparation. 17 D. Namgaladze, H. W. Hofer, and V. Ullrich, J. Biol. Chem. 277, 5962 (2002). 18 C. B. Klee, M. H. Krinks, A. S. Manalan, P. Cohen, and A. A. Stewart, Methods Enzymol. 102, 227 (1983). 5

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in crude brain extracts is 3–10 times that of the purified enzyme, it does not depend on exogenous metal ions but it is rapidly inactivated when exposed to Ca2 þ in the presence of CaM.15,17 The protection of calcineurin against inactivation by superoxide dismutase and its reactivation by ascorbate led Wang et al.15 to propose that inactivation was the result of oxidation of the iron cofactor. The low activity of the purified enzyme could then be due, not only to the partial loss of metal cofactors, but also to a Ca2 þ –CaMdependent oxidation of the active site Fe2 þ . To definitively identify the oxidation state of iron in the native enzyme the purification procedure had to be modified to prevent the loss of the natural cofactors. Different assay conditions were used throughout the purification to monitor both the depletion of metal cofactors (requirement for metal ions in standard assays) and the oxidation state of iron (activation by ascorbate under anaerobic assay conditions).

Calcineurin Assays Principle

The protein phosphatase activity of calcineurin is routinely measured by the Ca2 þ and CaM-dependent release of inorganic phosphate from a 32Plabeled commercially available synthetic peptide (DLDVPIPGRFDRRVSVAAE) corresponding to the phosphorylation site of type II regulatory subunit of cAMP-dependent protein kinase.19 Contribution of proteases to the liberation of radioactivity from the substrate is ruled out by the retention of degraded peptides on the DOWEX columns used to separate the residual phosphorylated peptide from inorganic phosphate.

Reagents

Substrate Phosphorylation of the synthetic peptide is accomplished by mixing 45 l (150 nanomoles) of a peptide solution (3.3 mM in H2O), 300 l (300 nmol) of 1 mM [ -32P]ATP (specific activity 3000–5000 cpm/pmol), 500 l of 2x phosphorylation buffer (40 mM MES buffer, pH 6.5, 0.4 mM EGTA, 0.8 mM EDTA, 4 mM MgCl2) adjusting the volume to 1 ml and adding 4 g (24
103 units) of catalytic subunit of cAMP-dependent protein kinase (Boehringer Mannheim Co., Indianapolis, IN). Almost complete phosphorylation is routinely obtained in 15 min and incubation at 19

D. K. Blumenthal, K. Takio et al., J. Biol. Chem. 261, 8140 (1985).

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30 C for 30 min yields reproducibly 0.8–0.9 mole Pi/mole peptide. The labeled peptide is separated from ATP and Pi by reverse phase chromatography on a disposable C18 cartridge (Sep-Pak from Waters). Prior to application of the radiolabeled peptide solution, the cartridge is first washed with 3 ml 30% acetonitrile in 0.1% trifluoroacetic acid followed by 5 ml 0.1% trifluoroacetic acid. The peptide solution is then applied to the cartridge with a 1 ml syringe and the cartridge washed with 0.1% trifluoroacetic acid (about 20 ml) until the effluent contains less than 1000 cpm/l. The washing procedure insures that blank values (incubation in absence of enzyme) are less than 0.1% of the total counts applied to the DOWEX columns (this is particularly important to quantitate low enzyme levels). The radiolabeled peptide is then eluted with 0.5 ml additions of 30% acetonitrile in 0.1% trifluoroacetic acid. The fractions containing the radiolabeled peptide, identified with a Geiger counter, are pooled (usually 1.5 to 2 ml) and divided in 8 fractions that are flash evaporated overnight in a Speed Vac. To avoid peptide loss over time nonsiliconized polyethylene tubes should be used for storage at 70 C. Stock Buffers stored at 20 C 2x Buffer I: 40 mM Tris–HCl, pH 8, containing 0.2 M KCl, 12 mM MgCl2 and 0.8 mg/ml bovine serum albumin. 2x Buffer II: 80 mM Tris–HCl, pH 7.5, containing 0.2 M KCl, 2 mM MnCl2 and 0.8 mg/ml bovine serum albumin. Inhibitors Okadaic acid (Boehringer Manheim Co.): A stock solution (3 M in dimethylsulfoxide) is diluted to 40 M in buffer I, sonicated for 3 min, and stored at 20 C. CaM antagonist: synthetic peptide M13 (FRRWKKNFIAVSAANRFKKISSSGAL): 1 mM stock solution in buffer I. FK506: 5 mM stock solution in ethanol diluted to 50 mM in buffer I and sonicated for 3 min prior to storage at 20 C. FKBP (Sigma), 2.5 mg/ml in 10 mM Hepes buffer, pH 8.0, 0.5 mM DTT, 0.004% NaN3. Reducing Agents Dithiothreitol (DTT): 0.5 M in deaerated water (stored at 70 C). Ascorbic acid (Sigma): 0.5 M in deaerated H2O adjusted to pH 7.5 with KOH (50 l aliquots are stored at 20 C).

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Metal Ions 0.03 M stock solution of Fe(NH4)2SO4 (AnalaR) from Gallard Schlesinger made under N2 in 10 mM HCl and stored under N2. 0.2 M MgCl2 (AnalaR) from Gallard Schlesinger. 0.2 M MnCl2 (AnalaR) from Gallard Schlesinger. DOWEX AG 50W-X8 DOWEX AG 50W-X8 (200–400 mesh, from BioRad) columns (0.5 ml) converted to the H þ form prior to use by washing with 10 ml H2O, 1 ml 1 M NaOH, 2 ml 1 M HCl and 4 ml H2O. The same columns, stored in 1 M NaOH, can be used for up to 6 months except when used more than five times for assays of calcineurin activity in crude tissue extracts. Stop Solution Mix 10 ml of 1 M potassium phosphate, pH 7.5, with 5 ml 100% trichloroacetic acid (w/v) and add 85 ml H2O. Anticalcineurin Antibodies A polyclonal antibody against bovine brain calcineurin in phosphatebuffered saline (PBS) can be used for the quantification of calcineurin in brain extracts but polyclonal IgGs against the C-terminal peptides of the  and isoforms of calcineurin A mixed with a polyclonal IgG against calcineurin B are preferable in other tissues that contain different relative amounts of calcineurin A isoforms. Standard solutions of recombinant calcineurin A and A and calcineurin B diluted to a concentration of 1 g/ml in denaturing solution prior to electrophoresis. Bovine erythrocyte superoxide dismutase (Worthington Biochemical Co., Freehold, NJ) 2 mg/ml in buffer I.

Procedure

The protein phosphatase activity of calcineurin is usually determined in the presence of EGTA, Ca2 þ , and Ca2 þ with calmodulin. Prior to assays the enzyme should be kept in the presence of 1–2 mM EGTA to avoid exposure of the active center that results in the inactivation of the enzyme. Standard Assays They are carried out in the presence of 6 mM Mg2 þ or 1 mM Mn2 þ . The purified enzyme, partially depleted of its natural metal cofactors

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(iron and zinc), requires exogenous metal ions for activity. The specific activity of calcineurin, measured under these conditions, varies with the extent of depletion of metal cofactors, the oxidation state of residual iron at the active site, and the nature of the metal ion used to activate the enzyme. Three mixes are prepared and kept at 0 C before starting the assays. Mix A: Buffer I is made 0.2 mM EGTA and 3 mM DTT prior to addition of calcineurin (always stored in the presence of  1 mM EGTA) at a final concentration of 0.075 m or 0.75 M to measure CaM-independent activity. Mix B: When present variable components are added to buffer I at the following concentrations: CaM (3 M), Mg2 þ (18 mM) or Mn2 þ (3 mM), M13 (1.8 M), superoxide dismutase (2 g/ml), FK506 (1.5 M), FKBP (3 M), okadaic acid (0.3 M). Mix C: Substrate (2.5–5
105 cpm/ml) is added to buffer I made 1 mM 2þ Ca or 3 mM EGTA when assays are carried out in the presence of EGTA (Mix C). Buffer II is substituted to Buffer I in the three mixes when the activity is measured in the presence of MnCl2 instead of MgCl2. In this case the activity cannot be measured in the presence of EGTA and the concentration of EGTA in Mix A is reduced to 1 mM to reduce EGTA concentration in the assay to 0.33 mM. Twenty l of Mix A is mixed with 20 l of Mix B and the mixture is incubated at 30  C for 3 min. The reaction is started by addition of 20 l of Mix C. After 5–10 min incubation at 30 C the reaction is stopped by addition of 0.5 ml of stop solution. Each reaction mixture is then applied to a Dowex column followed by 0.5 ml of H2O. The combined eluates are collected in scintillation vials and counted with 10 ml Aquasol. With purified calcineurin the reaction follows pseudo first order kinetics up to 30 min. Except in the case of the determination of kinetic constants, the rate constants are used to calculate the specific activity of the enzyme normalized to 1 M substrate as previously described.18 One unit of enzyme catalyzes the release of one micromole Pi per min in the presence of 1 M substrate under the assay conditions described above. Anaerobic Assays Anaerobic assays are used to detect the extent of oxidation and depletion of iron. They are carried out in the absence of Mg2 þ or Mn2 þ , in the absence of ascorbate and in the presence of 10 mM ascorbate with and without addition of 0.2 mM Fe2 þ . To insure detection of the stimulation of calcineurin by Fe2 þ , EGTA concentration in mixes A and B is reduced to 0.1 mM. Tubes containing Mix C and a mixture of 20 l of Mix A and 20 l of Mix B are first equilibrated at 0 C for 2 hr in an anaerobic chamber equilibrated with 95% N2 and 5% H2. When present, ascorbic acid is added to Mix A at a final concentration of 30 mM prior to be placed in

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CALCINEURIN ACTIVITY

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TABLE I a IN CRUDE RAT BRAIN EXTRACTS

Additions

EGTA Ca2 þ Ca2 þ þ M13 Ca2 þ þ FK506 þ FKBP12 Ca2 þ þ Mg2 þ

Specific activityb units/mg 0.005  0.005 0.1  0.005 0.02  0.001 0.010  0.001 0.09  0.002

a

Rat brain extracts were prepared as described in Ref. 20. The enzyme units are based on duplicate determinations under the standard conditions described in the text. To prevent inactivation during the assay the incubation time was limited to one min. b The specific activity is based on calcineurin concentration determined by quantitative analysis of Western blots.

the anaerobic chamber. The enzyme containing tubes are then closed, incubated for 3 min at 30 C and 20 l of Mix C added to start the reaction. Fe(NH4)2SO4 (2 l of the stock solution diluted to 9 mM in 1 mM HCl prior to assays) is added to the enzyme containing tubes 15 sec before addition of Mix C. Incubation time is continued but limited to 1–3 min to avoid loss of activity due to the dissociation of Fe2 þ or oxidation to Fe3 þ . If an anaerobic chamber is not available, all buffers and reagent solutions should be made under N2 with deionized H2O through which N2 has to be bubbled for at least 2 hr before use. Determination of Calcineurin Activity in Crude Tissue Extracts

As illustrated in Table I in crude rat brain extracts, inhibition by the calmodulin-binding peptide M13 and the complex of FK506 with its binding protein, FKBP, can be used to identify the Ca2 þ/CaM-dependent phosphatase activity of calcineurin assayed in the presence of the inhibitor of protein phosphatase-1 and 2A, okadaic acid.18 Cyclosporin A can be used instead of FK506 to inhibit calcineurin activity. In both cases addition of the respective exogenous binding proteins (cyclophilin A or FKBP12) is required to obtain complete inhibition. Addition of exogenous CaM is not required in brain extracts but may be necessary in other tissues. Because of the presence of high levels of okadaic acid-insensitive p-nitrophenylphosphatase and the inability to inhibit the p-nitrophenylphosphatase activity of calcineurin with FK506 or cyclosporin A, p-nitrophenylphosphate cannot 20

P. M. Stemmer, X. Wang et al., FEBS Lett. 374, 237 (1995).

296

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TABLE II MODIFIED PURIFICATION OF CALCINEURIN Specific activity b

a

Cn

Crude extract DE23 CaM-Sepharose

mg 188 80 11

Standard assay CaM þ CaM

0.02 0.003 0.001

0.030 0.009 0.009

ascorbate units/mg 0.03 0.010 0.02

Anaerobic assayc þ ascorbate þ ascorbate þ Fe2 þ

0.04 0.05 0.1

0.04 0.05 0.12

a

Calcineurin concentration in the crude extract and the DE23 fraction from 1 kg bovine brain was determined by quantitative Western blots and spectrophotometrically for the CaMSepharose fraction using an extinction coefficient of "1% 276nm ¼ 7.4 (C. B. Klee and P. M. Stemmer, unpublished observations). b Standard assays were carried out as described in the text in the presence of 6 mM Mg2 þ . c Anaerobic assays were carried out as described in the text in the presence of 1 M CaM and either in the absence of ascorbate, or in the presence of 10 mM ascorbate with and without 2 mM Fe2 þ . The incubation time was 2 min.

be used as a substrate to measure calcineurin activity in crude tissue extracts.21 The lack of stimulation and reproducible small inhibition observed upon addition of exogenous metal ions indicates that calcineurin has retained its endogenous metal cofactors. Provided that calcineurin is not exposed to Ca2 þ and CaM (the incubation time should be limited to 1–2 min), calcineurin activity in crude extracts can be measured under standard assay conditions. However, as illustrated in Table II, some preparations of crude brain extract can be stimulated up to two-fold when assayed under anaerobic conditions in the presence of ascorbate. At pH 7.5 the maximum value observed for the Vmax of calcineurin in crude rat brain extract is 43 mol/min mg with a Km of 210 M.* CaM-Dependent Inactivation of Crude Calcineurin

The high specific activity of the crude enzyme, which can be up to 20 times that of the purified enzymey assayed in the presence of Mg2 þ at pH 8.0 or Mn2 þ at pH 7.5, depends on short incubation times.20 21

X. Wang and C. B. Klee, unpublished observations.

*The km value varies from 40 to 200 M depending on the purity and homogeneity of the peptide substrate.21 y The specific activity of purified calcineurin varies between 0.005–0.05 units/mg (in the presence of Mg2 þ or Mn2 þ and 1 M substrate) depending on the extent of metal depletion during the purification procedure. It is always higher in the presence of Mn2 þ than in the presence of Mg2 þ .

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297

FIG. 1. Ca2 þ dependence of the CaM-mediated inactivation of calcineurin. Calcineurin activity in crude rat brain extract was measured under standard assay conditions. Prior to addition of Mix 3, crude brain extract (0.2 mg protein/ml) were incubated in the presence of Ca2 þ as indicated in the figure. Free Ca2 þ concentrations were calculated with a computer program using the binding constants of EGTA and EDTA for Ca2 þ of Fabiato and Fabiato.29 The concentration of Ca2 þ in Mix 3 was adjusted to insure the same final concentration of free Ca2 þ (0.3 mM ) in the assays after addition of substrate.21

Preincubation in the presence of Ca2 þ , but not EGTA, results in a rapid loss of activity.15 This Ca2 þ -dependent inactivation depends on the presence of Ca2 þ and endogenous CaM.15,17 No inactivation is observed when the enzyme is preincubated in the presence of EGTA or of Ca2 þ and M13. The Ca2 þ dependence of the CaM-dependent inactivation of calcineurin, illustrated in Fig. 1, is similar to that of CaM-dependent activation of the enzyme, suggesting that the inactivation also depends on the displacement of the autoinhibitory domain and exposure of the active site.21 Accordingly, calcineurin is protected against inactivation by addition of a synthetic peptide (ITSFEEKAKGLDRINERMPPRRDAMP) corresponding to the autoinhibitory domain of calcineurin.22 A 50% decrease of the rate in inactivation is observed upon addition of 50 M peptide during the preincubation.21 Protection of Calcineurin against Inactivation by Superoxide Dismutase

The protection of calcineurin against inactivation by superoxide dismutase suggested that the oxidation or reduction of calcineurin by 22

Y. Hashimoto, B. A. Perrino, and T. R. Soderling, J. Biol. Chem. 265, 1924 (1990).

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INHIBITION, STIMULATION, MODULATION OF ACTIVITY

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15 superoxide anion (O Accordingly, 2 ) was responsible for its inactivation. the rate of inactivation is increased in the presence of the superoxide generating agent, paraquat, or of xanthine and xanthine oxidase, and it is decreased under anaerobic conditions.15,17,21 At concentrations lower than 0.1 mM H2O2, the product of the dismutation of O 2 did not increase the rate of inactivation, and the inactivation was not prevented by addition of catalase.21 However, at higher concentrations (  1 mM) H2O2, as well as other oxidants, inhibits the phosphatase activity of crude as well as purified calcineurin which had lost 90% of its activity.23 The sensitivity of calcineurin to oxidants has also been observed in vivo.24 Provided that the mechanisms of O 2 and H2O2-mediated inhibition of calcineurin activity are identical, the precise identification of the oxidizing agent responsible for calcineurin inactivation remains to be determined. It was recently reported that nitric oxide (NO) protects calcineurin against inactivation by O 2 indicating that peroxynitrite resulting from the reaction of NO with O is unable to inhibit calcineurin under these conditions.17 2 Unlike tyrosine phosphatases, which are inactivated following oxidation of their active site cysteine, calcineurin does not contain an active site cysteine. Modification of the five exposed cysteines of calcineurin by Nethylmaleimide did not significantly affect its protein phosphatase activity.25 It was therefore likely that oxidation or reduction of iron at the active center, rather than oxidation of a cysteine residue, was responsible for the inactivation of calcineurin by O 2 . The reducing agents, DTT, cysteine, and -mercaptoethanol were not able to reactivate calcineurin, but a rapid reactivation of calcineurin was achieved by ascorbic acid, a good reducing agent of iron (Fig. 2). It suggested that the inactivation was the result of the conversion of the metal cofactor Fe2 þ to Fe3 þ .15–17

Identification of Calcineurin as an (Fe2 þ –Zn2 þ ) Protein Phosphatase

To definitively demonstrate that inactivation of calcineurin is the result of the conversion of the active site Fe2 þ to Fe3 þ , a purified preparation of calcineurin that has retained its metal cofactors was used to determine the oxidation state of iron by EPR spectroscopy and to correlate enzyme activity with the oxidation state of iron.

23

D. Sommer, K. L. Fakata, S. A. Swanson, and P. M. Stemmer, Eur. J. Biochem. 267, 2312 (2000). 24 T. A. Reiter, R. T. Abraham, M. Choi, and F. Rusnak, J. Biol. Inorg. Chem. 4, 632 (1999). 25 M.-P. Strub and C. B. Klee, unpublished observations.

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299

FIG. 2. Protection and reactivation of calcineurin by ascorbate. Calcineurin activity in crude rat brain extract (0.2 mg protein/ml) was measured under standard assay conditions in the absence of ascorbate (s), in the presence of 5 mM ascorbate added at zero time (m) or at 8 min as shown by the arrow. At the indicated times the percent of substrate remaining was determined in 60 ml aliquots of the incubation mixture.21

Modified Purification Procedure

To minimize the loss of endogenous metal cofactors, the procedure for the purification of calcineurin described in Ref. 18 was modified as recommended by King and Huang.8 Only 1 kg of brain tissue is processed at a time to shorten the time of exposure to Ca2 þ and CaM during the affinity chromatography step responsible for the loss of metal ions. Briefly, fresh brains,z obtained less than 2 hr after slaughtering the animals, are cut in square–inch cubes and homogenized in 2.5 liters of buffer A18 in 1.5 liters batches in a Warring blender for 15 sec once at low and once at medium speed. The temperature should not exceed 10 C. After dilution with 12 liters of buffer B18 the DE-23 cellulose chromatography step is performed as previously18 described with the volumes of buffer, the size of the column, flow rates and fraction sizes adjusted to the reduced size of the preparation. Reducing the size of the calmodulin–Sepharose column from 75 to 50 ml for a preparation of calcineurin from 1 kg of brain tissue decreases the yield to 10–15 mg for 1 kg of brain tissue, but insures the

z The specific activity of crude calcineurin varies from 0.04 to 0.2 units/mg with different preparation of brain extracts. Prolonged exposure to Ca2 þ , caused by delays in obtaining fresh tissues or methods of euthanasia, greatly reduced the activity of crude calcineurin. The activity of calcineurin in homogenates of frozen brains is also greatly reduced and requires Mg2 þ for activity.

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INHIBITION, STIMULATION, MODULATION OF ACTIVITY

[22]

selective binding of calcineurin.} This two-step procedure yields calcineurin that is reproducibly more than 90% pure and allows one to bypass the Sephadex-G200 and the electrofocusing steps. A comparison of the specific activities of calcineurin measured under standard assay conditions to that measured under anaerobic conditions throughout this procedure is shown in Table II. The calcineurin activity in the crude bovine brain extract and the DE23 cellulose eluate is always measured in the presence of okadaic acid. Provided that the incubation time is less than 2 min, the specific activity of calcineurin in the crude extract, measured under standard conditions in the presence of Mg2 þ , is similar to that measured under anaerobic conditions in the absence of added metal. A slight stimulation by ascorbate reveals some degree of inactivation. The low specific activity compared to that of rat brain extracts (Table I) is reproducible and may reflect the fact that the brains were not immediately soaked in a buffer containing EGTA, as was done for the rat brains. Optimum concentrations of ascorbate required to reverse the inactivation in crude extracts have not been determined. It is also possible that inactivation of calcineurin by prolonged exposure to Ca2 þ is not reversible by ascorbate. After the DE-23 cellulose chromatography step calcineurin activity is dependent upon addition of CaM and the removal of superoxide dismutase renders the enzyme more susceptible to inactivation, as indicated by the three-fold increase of the specific activity in the presence of ascorbate. As expected after the affinity chromatography step, carried out in the presence of Ca2 þ , an even greater stimulation by ascorbate is observed.ô The small stimulation by Fe2 þ suggests that calcineurin has retained most of its iron cofactor. Because of the inhibitory effect of Zn2 þ on enzyme activity due to the substitution of Fe2 þ by Zn2 þ , depletion of the zinc site cannot be estimated.26 The large increase of the specific activity of calcineurin upon addition of ascorbate measured under anaerobic conditions in the absence 26

}

M. Ghosh, S. P. Li, and C. B. Klee, unpublished observations.

The size of the CaM-Sepharose column depends on the batch of CaM-Sepharose. Commercially available CaM-Sepharose has a higher capacity than CaM-Sepharose prepared as described in Klee et al.18 It is recommended to test different batches of CaM-Sepharose to determine the optimum size of the column yielding highly purified calcineurin. This modified purification procedure is well suited to the purification of brain calcineurin which is a major component of CaM-binding proteins in this tissue but it may be difficult to achieve the same degree of purity with other tissues or organisms where calcineurin is not as abundant. Anaerobic conditions and addition of superoxide dismutase or ascorbate may be necessary to detect calcineurin activity by using longer incubation times. ô The large increase in the specific activity of calcineurin after the affinity chromatography step suggests that the low activity in the crude extract may be due to the presence of inhibitory proteins removed during the CaM-Sepharose step.

[22]

REDOX REGULATION OF CALCINEURIN

301

of added metal ions supports the conclusion that the low activity of this preparation of calcineurin is not the result of a significant loss of metal cofactors but of a modification of the oxidation state of iron. Determination of Metal Content by Atomic Absorption Spectroscopy

Retention of the metal cofactors is confirmed by determination of iron and zinc content of the affinity-purified calcineurin with a Perkin Elmer Model 5000 spectrometer equipped with a HGA-500 graphite furnace. Calcineurin is first subjected to a gel filtration on a Sephadex G25 column equilibrated with 40 mM Chelex-treated Tris–HCl buffer, pH 7.5 containing 0.1 M KCl to which is added 0.1 mM DTT and 1 g/ml leupeptin, as indicated below. This chelex-treated buffer contains less than 0.3 M iron and 1 M zinc. Standard curves in the corresponding buffers and protein solutions are used to correct for metal content and matrix effects. Zinc and iron Atomic Spectroscopy Standards (1 mg/ml in 2% nitric acid) from Perkin Elmer Co. are diluted with the buffer used to elute the enzyme from G-25 column to get the standard curves. The disodium salt of EDTA (final concentration of 0.1 mM) is added to the iron-containing solutions to prevent precipitation of iron as ferric hydroxide. The preparations of calcineurin purified by this modified protocol, which have a specific activity greater than 0.1 units/mg determined under anaerobic conditions in the absence of Fe2 þ , contain routinely 0.8–0.9 mol of iron and 0.8–1 mol of zinc per mol of enzyme.26 They are suitable for the identification of the redox state of iron by EPR spectroscopy. Identification of the Oxidation State of Iron

After the Sephadex G-25 chromatography, 8–10 mg of calcineurin (10– 15 ml) are concentrated in a Centriprep-30 concentrator to a final volume of 3 ml. If needed, calcineurin is subjected to a second buffer exchange to insure removal of contaminating iron and zinc. Calcineurin, further concentrated to 350 l to a final concentration of 17–20 mg/ml, can be used for EPR measurements in liquid nitrogen or preferably liquid helium. The loss of metal ions, reproducibly observed during this concentration procedure, can be minimized, but not prevented, by rapid handling of the concentration steps. As was reported for the (Fe3 þ –Zn2 þ ) binuclear centers of kidney bean purple acid phosphatase the EPR spectrum revealed a signal at g ¼ 4.3.16,27 A quantitative analysis of this signal indicates that 90% of iron 27

J. L. Beck, J. de Jersey, B. Zerner, M. P. Hendrich, and P. G. Debrunner, J. Am. Chem. Soc. 110, 3317 (1988).

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INHIBITION, STIMULATION, MODULATION OF ACTIVITY

[22]

associated with calcineurin is in the oxidized form. The loss of the EPR signal accompanied with an increase of the specific activity from 0.004–0.01 units/mg to 0.1–0.12 units/mg after treatment with 10 mM sodium ascorbate for 10 min at room temperature is a strong indication that EPR signal is due to Fe3 þ bound to calcineurin rather than contaminating Fe3 þ . Conversely a weak signal at g ¼ 4.3 in a preparation of calcineurin purified in the presence of ascorbate, which has preserved a moderate enzymatic activity, is greatly increased upon treatment with H2O2 while its activity is decreased.17 The signal at g ¼ 4.3, which is not affected by phosphate binding, was previously attributed to contaminating iron as opposed to the signals at g ¼ 5.5 and 8.1 shifted by phosphate to 5.0 and 8.4.14 Since these two signals can only be detected at temperature below 10 K it is advisable to perform the EPR experiments in liquid helium. Signals at g ¼ 6.5, 5.5, and 5.0 are only detectable after oxidation in the reduced form of calcineurin.17 Determination of the pH dependence of calcineurin activity adds support to its identification as an (Fe2 þ –Zn2 þ ) protein phosphatase. As shown in Fig. 3 the pH optimum of the affinity purified ascorbate-reduced calcineurin is 6.1, significantly higher than 4.9 for the (Fe3 þ –Zn2 þ ) acid purple phosphatases.16 The pH-dependence of ascorbate treated calcineurin is almost identical to that of the crude enzyme and its activity is not

FIG. 3. pH dependence of the protein phosphatase activity of calcineurin. The Ca2 þ /CaM stimulated protein phosphatase activity of purified calcineurin containing 0.8 mol of iron and 0.8 mol of zinc was measured under anaerobic conditions in the presence of ascorbate (panel A) and under standard conditions (panel B) in 40 mM MES buffer pH 5.5, 5.8, and 6.1, 40 mM MOPS buffer pH 6.7, and 7.0 and 40 mM Tris–HCl buffer pH 7.5 and 8.0. No metal added (s), and in the presence of 0.1 mM Mn2 þ (
), 6 mM Mg2 þ (j), 0.5 mM Fe2 þ (d). The pH dependence of crude rat brain calcineurin measured under identical conditions but in the ) is shown for comparison. The incubation time absence of ascorbate or added metal ( was 1 min.16,21

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303

significantly affected by Mn2 þ or Mg2 þ indicating that calcineurin has retained its metal cofactors. Accordingly, prior to reduction calcineurin activity is very low and only slightly stimulated by Mn2 þ with a pH optimum at 7.5 and Mg2 þ at pH 8.0. This stimulation is significantly lower than the one observed with standard preparations of calcineurin which have retained less than stoichiometric amounts of their metal cofactors. It is recommended to take advantage of the two-fold increased activity at pH 6.1 in the presence of ascorbate and anaerobic conditions to detect calcineurin in tissues or organisms where low levels of calcineurin are difficult to detect prior to purification.

Concluding Remarks

There is general agreement that calcineurin contains a binuclear iron– zinc active center. The inactivation of the p-nitrophenylphosphatase activity of calcineurin by dithionite and the failure to reactivate it by ascorbate led Yu et al.9 to propose that the (Fe3 þ –Zn2 þ ) is the active form of the enzyme. It was also shown that the p-nitrophenylphosphatase activity of Fe2 þ substituted calcineurin is inhibited not only by reduction with dithionite but also by oxidation with H2O2 suggesting that it requires a mixed valence (Fe2 þ –Fe3 þ ) active center.14 More recent evidence indicates that, unlike the p-nitrophenylphosphatase, the protein phosphatase activity of (Fe3 þ –Zn2 þ ) calcineurin is increased by ascorbate and only slightly inhibited by dithionite and therefore depends on the reduced state of its iron cofactor which is oxidized during the purification procedure. In contrast its p-nitrophenylphosphatase activity is not activated by ascorbate and apparently not dependent on the reduced state of iron.9 The basis for the different requirements for the dephosphorylation of these substrates remains to be elucidated. In contrast to calcineurin, the purple acid phosphatases do not require Fe2 þ for activity. A comparison of the coordination spheres of iron in these two classes of enzymes suggest that the tyrosine residue coordinated to iron in purple acid phosphatase may increase its nucleophilicity and prevent a requirement for Fe2 þ . In calcineurin this tyrosine is replaced by a histidine. The perfect conservation of the iron coordinating amino acid residues in all the known sequences of calcineurin suggests that the requirement for Fe2 þ is equally conserved. Indeed, both human and yeast calcineurin are protected against inactivation by ascorbate and inactivated by O 2 . Recombinant calcineurin A coexpressed in E. coli with calcineurin B also contains iron and zinc as metal cofactors and requires treatment with ascorbate for activity. However, it is difficult to

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obtain calcineurin preparations with stoichiometric contents of iron and zinc because of the relatively low affinity of the enzyme for these two metals. Attempts to reconstitute the (Fe2 þ –Zn2 þ ) enzyme are impaired by the competitive binding of either one of the two metal ions to the two sites. The sensitivity of calcineurin to oxidative stress has also been observed in vivo. Two independent signaling pathways affected by calcineurin in yeast, the recovery from G1 arrest after pheromone treatment and the adaptation to a high salt stress, are dependent on superoxide dismutase when cells are grown under aerobic but not anaerobic conditions.15 In hippocampal neurons, the Ca2 þ - and time-dependent inactivation of calcineurin upon  prolonged exposure to Ca2 þ , which is prevented by O2 scavengers, provides a mechanism to insure a temporal modulation of the Ca2 þ signal.28 Addition of H2O2 to culture media inhibits the calcineurin-mediated activation of the transcription factor, NFAT, in Jurkat cells.24 Thus, the reversible inactivation of calcineurin by oxidation of its binuclear metal center provides a mechanism to couple the Ca2 þ regulation of cellular processes with the redox potential of the cells and revealed a novel regulatory role for iron.

28 29

H. Bito, K. Deisseroth, and R. W. Tsien, Cell 87, 1203 (1997). A. Fabiato and F. Fabiato, J. Physiol. (Paris) 75, 463 (1979).

[22] Analysis of the Regulation of Protein Tyrosine Phosphatases in Vivo by Reversible Oxidation By TZU-CHING MENG and NICHOLAS K. TONKS

The Protein Tyrosine Phosphatases (PTPs) are now established as critical regulators of the signal transduction events that underlie such fundamental cellular functions as growth, proliferation, differentiation, metabolism, and motility. The PTPs are defined by the presence of a signature sequence motif, [I/V]HCXXGXXR[S/T]. They can be subdivided into two broad categories, those enzymes that are specific for phosphotyrosyl residues in proteins, termed the classical PTPs, and the dual specificity

METHODS IN ENZYMOLOGY, VOL. 366

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obtain calcineurin preparations with stoichiometric contents of iron and zinc because of the relatively low affinity of the enzyme for these two metals. Attempts to reconstitute the (Fe2 þ –Zn2 þ ) enzyme are impaired by the competitive binding of either one of the two metal ions to the two sites. The sensitivity of calcineurin to oxidative stress has also been observed in vivo. Two independent signaling pathways affected by calcineurin in yeast, the recovery from G1 arrest after pheromone treatment and the adaptation to a high salt stress, are dependent on superoxide dismutase when cells are grown under aerobic but not anaerobic conditions.15 In hippocampal neurons, the Ca2 þ - and time-dependent inactivation of calcineurin upon
prolonged exposure to Ca2 þ , which is prevented by O2 scavengers, provides a mechanism to insure a temporal modulation of the Ca2 þ signal.28 Addition of H2O2 to culture media inhibits the calcineurin-mediated activation of the transcription factor, NFAT, in Jurkat cells.24 Thus, the reversible inactivation of calcineurin by oxidation of its binuclear metal center provides a mechanism to couple the Ca2 þ regulation of cellular processes with the redox potential of the cells and revealed a novel regulatory role for iron.

28 29

H. Bito, K. Deisseroth, and R. W. Tsien, Cell 87, 1203 (1997). A. Fabiato and F. Fabiato, J. Physiol. (Paris) 75, 463 (1979).

[23] Analysis of the Regulation of Protein Tyrosine Phosphatases in Vivo by Reversible Oxidation By TZU-CHING MENG and NICHOLAS K. TONKS

The Protein Tyrosine Phosphatases (PTPs) are now established as critical regulators of the signal transduction events that underlie such fundamental cellular functions as growth, proliferation, differentiation, metabolism, and motility. The PTPs are defined by the presence of a signature sequence motif, [I/V]HCXXGXXR[S/T]. They can be subdivided into two broad categories, those enzymes that are specific for phosphotyrosyl residues in proteins, termed the classical PTPs, and the dual specificity

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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phosphatases (DSPs), which have the ability to recognize Ser/Thr, as well as Tyr residues. In the classical PTPs the signature motif is contained within a conserved catalytic domain of 280 residues, which is flanked on either the Nor C-terminal side by noncatalytic sequences that serve a regulatory function. The classical PTPs exist as either receptor-like, transmembrane proteins, consistent with a potential role in the direct regulation of signal transduction by ligand controlled dephosphorylation of tyrosyl residues in proteins, or as nontransmembrane, cytoplasmic species. There are 38 classical PTPs, which can be grouped into 17 subtypes, depending upon structural similarities in their catalytic and regulatory domains.1 The DSPs display greater variation in structure than the classical PTPs and, although there is conservation in the fold of the catalytic domain, sequence similarity between the two groups is largely restricted to the signature motif. The DSPs can be divided into three groups, the largest of which comprises the VH1like enzymes and which includes those DSPs that have been implicated in the regulation of MAP kinases.2 In addition, there are the myotubularins (MTMs)3,4 and the cell cycle regulatory proteins cdc25A, B and C.5 Current estimates suggest that the family of PTPs in humans will comprise a total of  100 enzymes.6 Now that the sequencing of human genome is completed and the composition of the PTP family can be defined, it will be important to develop mechanisms to explore their function. Consistent with an important function as regulators of signal transduction, the activity of the PTPs is tightly controlled in vivo. Various mechanisms of regulation have been described, from ligand binding to the receptor-like PTPs and targeting of nontransmembrane PTPs to defined subcellular locations, to regulation of PTP function by covalent modification, such as phosphorylation and proteolysis. Most recently a novel tier of control of PTP function has become apparent. It is well established that phagocytic cells, such as neutrophils, produce Reactive Oxygen Species (ROS) to support their microbicidal function. More recently it has become apparent that the production of ROS in nonphagocytic cells is important for propagating an optimal response to stimulation by growth factors, cytokines, and hormones. Thus, in a manner analogous to reversible protein phosphorylation, the 1

J. N. Andersen, O. H. Mortensen, G. H. Peters, P. G. Drake, L. F. Iversen, O. H. Olsen, P. G. Jansen, H. S. Andersen, N. K. Tonks, and N. P. Moller, Mol. Cell. Biol. 21, 7117–7136 (2001). 2 A. Theodosiou and A. Ashworth, Genome Biol. 3, 3009.1–3009.10 (2002). 3 J. Laporte, F. Blondeau, A. Buj-Bello, and J. L. Mandel, Trends Genet. 17, 221–228 (2001). 4 M. J. Wishart, G. S. Taylor, J. T. Slama, and J. E. Dixon, Curr. Opin. Cell. Biol. 13, 172–181 (2001). 5 I. Nilsson and I. Hoffmann, Prog. Cell. Cycle. Res. 4, 107–114 (2000). 6 N. K. Tonks (2003) Handbook of Cell Signaling 1, in press.

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reversible oxidation of target proteins in a cell may regulate the function of those proteins in response to various agonists and thus elicit a cellular response to stimulation.7,8 The important question then becomes what are the targets of ROS that mediate these effects on signaling. The unique features of the active site of the PTPs and their mechanism of catalysis has drawn attention to their potential for regulation by reversible oxidation.

Mechanism of PTP-Mediated Catalysis

The PTP active site is present as a pronounced cleft on the surface of the enzyme. The base of this cleft, which is formed by the PTP loop, contains the signature motif that defines the PTP family, [I/V]HCXXGXXR[S/T], within which the Cys and Arg residues are invariant and essential for catalysis. The PTP loop adopts a cradle conformation to bind the phosphate moiety of the substrate and to position the essential Cys residue for nucleophilic attack on the substrate. PTPs catalyze dephosphorylation of their substrates via a 2-step mechanism involving a thiophosphate intermediate of the active site Cys. In the classical PTPs the sides of the cleft are formed by three motifs. The ‘‘pTyr loop’’ contains a Tyr residue (Tyr 46 in PTP1B) that is highly conserved in the classical PTPs and defines the depth of the active site cleft, thereby conferring specificity for pTyr residues as substrates. This residue is not present in the DSPs, which are characterized by a more open active site cleft. The ‘‘Q loop’’ contains a Gln residue, equivalent to Gln 262 in PTP1B, which mediates hydrolysis of the cysteinyl phosphate catalytic intermediate. Finally, there is a conformationally flexible loop (the WPD loop), which moves to close the active site following substrate binding and contains the essential, invariant Asp residue (D181 in PTP1B). In the first step of catalysis this Asp functions as a general acid to protonate the phenolate leaving group of the substrate whereas in the second step it functions as a general base, in combination with the Gln residue of the Q loop, to activate a water molecule and promote hydrolysis of the phospho-enzyme intermediate.9 Due to the environment of the active site, in particular the presence of the invariant Arg residue in the signature motif, the catalytic Cys displays an unusually low pKa and is present predominantly as the thiolate anion at neutral pH. This not only enhances its ability to execute a nucleophilic attack on the phosphate group of the substrate in the first step of catalysis, but also renders it susceptible to oxidation. In fact, we noted in the original characterization of PTP1B that the presence of reducing agents was essential to maintain 7

T. Finkel, Curr. Opin. Cell. Biol. 10, 248–253 (1998). T. Finkel, FEBS Lett. 476, 52–54 (2000). 9 D. Barford, Z. Jia, and N. K. Tonks, Nat. Struct. Biol. 2, 1043–1053 (1995). 8

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FIG. 1. Schematic representation of the regulation of PTP activity by oxidation. In the unique environment of the PTP active site, the invariant Cys residue of the signature motif displays an unusually low pKa and is present predominantly as the thiolate anion. This enhances its nucleophilic properties but renders the PTPs susceptible to oxidation. Oxidation can yield a single-oxidized sulfenic acid modification of the Cys (Cys-SOH), which inhibits activity because the oxidized Cys can no longer function as a nucleophile. The sulfenic acid is rapidly converted to a sulfenamide (isothiazolidine) by formation of a covalent bond between the sulfur atom of the Cys and the main chain nitrogen atom of the adjacent residue, Ser 216 (represented as ‘‘SN’’ in the figure), resulting in a novel 5-atom ring structure at the active site.10a,10b This novel covalent modification induces a profound conformational change at the active site in which the oxidized Cys adopts a solvent-exposed position that would facilitate reduction to the active form of the enzyme. This modification also helps to prevent further oxidation that would result from the addition of 2 (sulfinic acid) or 3 (sulfonic acid) oxygens to the active site Cys, which would lead to irreversible oxidation of the enzyme. Therefore, oxidation has the potential to form the basis for a mechanism of reversible regulation of PTP activity. (See Color Insert.)

catalytic activity.10c It has now been shown that treatment of various PTPs,11 dual specificity phosphatases12 and low molecular weight PTPs13 with H2O2 in vitro leads to oxidation of the active site Cys with concomitant inhibition of activity, because the modified Cys can no longer function as a phosphate acceptor. 10a

A. Salmeen, J. A. Anderson, M. P. Myers, T.-C. Meng, J. A. Hinks, N. K. Tonks, and D. Barford, Nature 423, 769–773 (2003). 10b R. L. M. van Montfort, M. Congreve, D. Tisi, R. Carr, and H. Jhoti, Nature 423, 773–777 (2003). 10c N. K. Tonks, C. D. Diltz, and E. H. Fischer, J. Biol. Chem. 263, 6731–6737 (1988). 11 S. R. Lee, K. S. Kwon, S. R. Kim, and S. G. Rhee, J. Biol. Chem. 273, 15366–15372 (1998). 12 J. M. Denu and K. G. Tanner, Biochemistry 37, 5633–5642 (1998). 13 A. Caselli, R. Marzocchini, G. Camici, G. Manao, G. Moneti, G. Pieraccini, and G. Ramponi, J. Biol. Chem. 273, 32554–32560 (1998).

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Although oxidation of Cys by the addition of 2 (sulfinic acid) or 3 (sulfonic acid) oxygens would be irreversible, oxidation to sulfenic acid (Cys-SOH) is reversible, and, therefore, has the potential to form the basis of a mechanism for regulation of PTP activity in vivo (Fig. 1). In fact, the oxidation of PTPs in response to EGF,14 insulin,15 and PDGF16 has now been demonstrated. A major obstacle to such studies is the requirement for a method by which the oxidized/inactivated PTPs could be distinguished from reduced/activated PTPs in a cellular context. Here we describe a modified ‘‘in-gel’’ PTP activity assay to allow visualization of a profile of PTPs that become oxidized following a particular stimulus. General Principles of the Assay

Triggering of the cells with an appropriate stimulus that generates intracellular ROS leads to the production of two pools of PTPs, those that have become oxidized in response to the stimulus and those that remain unaffected. These pools can be distinguished by harvesting the cells under anaerobic conditions and lysing in a buffer containing the alkylating agent iodoacetic acid (IAA). Anaerobic conditions will minimize the possibility of postlysis oxidation. The majority of the PTPs in the lysate, which have not encountered ROS in the cell, will be irreversibly inactivated by alkylation of their active site Cys by the IAA. In contrast, those PTPs in which the active site Cys has been oxidized to sulfenic acid in response to the stimulus will be protected from alkylation. The activity of this latter pool of PTPs can be regenerated and visualized in an ‘‘in-gel’’ phosphatase assay, applying the procedure originally described in Ref. 17. An aliquot of cell lysate is subjected to SDS-PAGE in a gel that has been cast containing a radioactively-labeled substrate. Proteins in the gel are sequentially denatured, then renatured in the presence of reducing reagents. Under these conditions, the activity of the PTPs in which the active site Cys had been subjected to stimulus-dependent oxidation to sulfenic acid was recovered, whereas those that were not oxidized in response to the initial stimulus, and were irreversibly alkylated in the lysis step, remain inactive. The gel is then left to incubate in the renaturation buffer, allowing dephosphorylation of substrate to occur in the region of the gel immediately surrounding the PTP proteins. The reaction is then terminated by staining and destaining of the gel, which also provides a record 14

Y. S. Bae, S. W. Kang, M. S. Seo, I. C. Baines, E. Tekle, P. B. Chock, and S. G. Rhee, J. Biol. Chem. 272, 217–221 (1997). 15 K. Mahadev, A. Zilbering, L. Zhu, and B. J. Goldstein, J. Biol. Chem. 276, 21938–21942 (2001). 16 T. C. Meng, T. Fukada, and N. K. Tonks, Mol. Cell. 9, 387–399 (2002). 17 K. Burridge and A. Nelson, Anal. Biochem. 232, 56–64 (1995).

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FIG. 2. Modified ‘‘in-gel’’ PTP assay to measure oxidation of PTPs induced by cellular stimuli. After triggering with an appropriate stimulus two pools of PTPs are created within the cell, the majority of the PTPs that remain unaffected and those in which the active site Cys has been oxidized. Following lysis under anaerobic conditions in buffer containing IAA, those PTPs that have not encountered ROS in the cell become irreversibly inactivated by alkylation of their active site Cys. In contrast, any PTPs that were oxidized in response to the stimulus are resistant to alkylation. The mixture is then subjected to an ‘‘in-gel’’ phosphatase assay in a SDS-PAGE gel that is cast containing a radioactively-labeled substrate. An aliquot of cell lysate is subjected to SDS-PAGE and proteins in the gel are sequentially denatured, then renatured in the presence of reducing reagents. Under these conditions, the activity of the PTPs in which the active site Cys had been subjected to stimulus-dependent oxidation to sulfenic acid is recovered, whereas those that were not oxidized in response to the initial stimulus, and were irreversibly alkylated in the lysis step, remain inactive. When the gel is left to incubate in the renaturation buffer, dephosphorylation of substrate occurs in the region of the gel immediately surrounding the PTP proteins. After staining and destaining, the gel is dried and exposed to film. The presence of a PTP can be visualized by substrate dephosphosphorylation, with the appearance of a clear, white area on the black background of labeled substrate. (See Color Insert.)

of protein loading. The gel is finally dried and exposed to film. The presence of a PTP is visualized by substrate dephosphorylation and the appearance of a clear, white area on the black background of labeled substrate (Fig. 2).

Buffers

Buffer A: 25 mM sodium acetate pH 5.5, 1% NP-40, 150 mM NaCl, 10% Glycerol. Buffer B: 20 mM Tris–HCl pH 8, 150 mM NaCl, 2.5 mM EDTA, 1 mg/ml lysozyme.

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Buffer C: 50 mM imidazole pH 7.2, 10 mM DTT. Buffer D: 50 mM imidazole pH 7.2, 10 mM DTT, 5mM EDTA, 50% glycerol. Buffer E: 50 mM imidazole pH 7.2, 10 mM DTT, 0.1% 2-mercaptoethanol. Buffer F: 50 mM imidazole pH 7.2, 30 mM MgCl2, 10 mM MnCl2, 10 mM DTT, 1 mM Na3VO4, 1% Triton X-100, 0.2 mM ATP, 0.1% 2-mercaptoethanol. Buffer G: 50 mM Tris–HCl pH 8, 20% isopropanol. Buffer H: 50 mM Tris–HCl pH 8, 0.2% 2-mercaptoethanol. Buffer I: 50 mM Tris–HCl pH 8, 6 M Guanidine-HCl, 0.3% 2mercaptoethanol. Buffer J: 50 mM Tris–HCl pH 8, 0.04% Tween 40, 1 mM EDTA, 0.03% 2-mercaptoethanol Construction of a Customized Anaerobic Work Station

In light of the susceptibility to oxidation displayed by the active site Cys of members of the PTP family, it is important to perform these studies in an anaerobic environment so as to minimize any postlysis oxidation. A customized anaerobic work station (overall dimensions: 20 inches length  36 inches width  26 inches height, shown in Fig. 3) was produced by the

FIG. 3. The work station used to generate cell lysates in an anaerobic environment. (See Color Insert.)

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Machine Shop of Cold Spring Harbor Laboratory, to provide an oxygenfree environment to prepare cell lysates. Two side entries were made with removable, airtight sealed doors ( 12 inches in diameter) that provide easy transfer of materials and equipment in and out of the work station. Two bare hand entry ports (Fisher Scientific, #11-391-103) were mounted in holes (7.75 inches in diameter) of the front of the work station. These ports were made of 10 layers of soft rubber, which provide easy access and withdrawal of hands while maintaining an airtight seal. An entry port to introduce a flow of argon is included in the top of the left side panel of the chamber with the exit port placed at the bottom of the right side panel. Prior to commencing the experiment, the required accessories (including cell scrapers, pipettes, tips, buffers, samples, ice tray for processing lysates, vacuum line to aspirate media, and a vortex mixer) should be placed inside the work station, as shown in Fig. 3. High purity argon gas is used to purge the work station initially and a consistent argon flow maintains an oxygen-free environment while cells are lysed inside the chamber.

Preparation of Cell Lysates

In order to promote the anaerobic environment and ensure that the oxidation status of the PTPs in the lysate reflects accurately that of the intact cell, degassed buffers should be used for these studies. Degas the Buffer A at 30 in-Hg vacuum for at least 1 hr and, immediately prior to use, supplement with iodoacetic acid (IAA, 10 mM), catalase (100 g/ml), superoxide dismutase (100 g/ml), aprotinin (25 g/ml) and leupeptin (25 g/ml), then protect the buffer from light and place it inside the anaerobic work station. Following stimulation of cells with the agonist under investigation (such as growth factors, cytokines, hormones, or stress stimuli), move the cell plates inside the anaerobic working station with constant flow of argon and close the side entry doors. Rinse the cells with degassed 1x PBS, then lyse by addition of degassed IAA-containing Buffer A to the plate. The amount of lysis buffer varies between cell lines (e.g., 0.7 ml for Rat 1 fibroblasts, 1.0 ml for HepG2 cells) and optimal conditions should be determined empirically. Scrape the cells into lysis buffer to facilitate lysis, then transfer the lysate to brown-color tubes. After a brief vortex, remove tubes from the anaerobic work station and incubate the lysates on a rocker at room temperature for 10 min. Precipitate cell debris by centrifugation at 12,000g for 1 min, then aspirate the supernatant and mix it with 2x SDS-PAGE loading buffer immediately and boil for 4 min.

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Measurement of PTP Activity Using the In-Gel Assay Preparation of a Radioactively-Labeled, Phosphorylated Substrate

Preparation of the Protein Tyrosine Kinase GST-FER Transform competent E. coli cells (strain BL-21) with plasmid encoding GST-FER fusion protein.18 Pick a single colony from the LB-AMP plate for a small-scale culture (25 ml LB-AMP) in a shaker at 37 C for overnight. Inoculate 10 ml of overnight culture in 240 ml LB-AMP. After 2 hr of culture at 37 C, add IPTG to a final concentration of 0.4 mM. Transfer the culture to a shaker at 30 C for additional 2 hr of incubation. Harvest the bacterial cells by centrifugation at 6000g at 4 C for 30 min, then resuspend the pellet in 7 ml of ice-cold Buffer B, and incubate at 4 C on a rocker for 20 min. Add the following reagents to the bacterial lysate at the indicated concentrations (Triton X-100, 1%, DNase, 2.5 g/ml, RNase, 10 g/ml, PMSF, 1 mM and DTT 10 mM), then incubate at 4 C for additional 15 min. Briefly sonicate the lysate for 15 sec to shear DNA and reduce viscosity, then precipitate the cell debris by centrifugation at 20,000g for 30 min at 4 C. Recover the supernatant and incubate it with 250 l of GSHSepharose beads (Amersham-Pharmacia) at 4 C for 30 min. Wash the beads twice with 7 ml Buffer C and, after the final wash, aspirate the buffer completely then add 250 l of Buffer D. Resuspend the beads and divide the preparation into 50 l aliquots in screw top vials for storage at
70 C. Labeling of the Substrate Thaw a 50 l aliquot of the GST-FER beads on ice, then wash twice with 1 ml of Buffer E. After the final wash, resuspend the beads in 25 l of Buffer E and add the following reagents to the tube sequentially: 800 l of kinase reaction buffer (Buffer F), 3.5 l of [ ] 32P-labeled ATP (ICN crude grade product #35020, equivalent to 0.4 mCi) and unlabeled substrate (50 l of a 20 mg/ml stock of Reduced Carboxamidomethylated and Maleylated Lysozyme (RCML) or poly Glu : Tyr [4 : 1] (Sigma)). Other substrates could also be tested according to requirements. Place the tube on a rocker behind a plexiglass shield and incubate at room temperature for 6 hr. Terminate the kinase reaction by addition of 100 l of 100% TCA and allow to incubate on ice for at least 30 min, to precipitate the 32P-labeled substrate. Harvest the labeled substrate by centrifugation at 12,000g for 15 min at room temperature. Discard the supernatant, and resuspend the precipitate in 200 l of Tris-base (pH 11). Separate the dissolved substrate from 18

Y. Shen, G. Schneider, J. F. Cloutier, A. Veillette, and M. D. Schaller, J. Biol. Chem. 273, 6474–6481 (1998).

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GST-FER beads by centrifugation at 12,000g at room temperature for 5 min. Collect the supernatant and discard the beads that form the pellet. Equilibrate a prepacked Sephadex G-25 column (Amersham-Pharmacia PD-10 columns, #17-0851-01) with 10 ml of 50 mM imidazole pH 7.2 containing 0.1% fish gelatin as a carrier protein to reduce nonspecific binding to the column. Subsequently wash the column with 10 ml of 50 mM imidazole pH 7.2 alone. Apply the 32P-labeled substrate dissolved in Trisbase to the Sephadex G-25 column, then add 8 ml of 50 mM imidazole buffer pH 7.2 to elute the substrate. Collect the eluate in 8 fractions of 1 ml volume each. Count 2 l of each fraction in scintillant to identify the fractions enriched in the labeled substrate (usually fraction 4). For RCML, fraction 4 should contain more than 1  105 cpm/l, whereas for poly (Glu : Tyr) 4 : 1, there should be 2.5  105 cpm/l.

In-gel PTP Assay

Prepare SDS-PAGE Gel Containing

32

P-Labeled Substrate

Mix buffers and reagents (except TEMED and ammonium persulfate) required for a 10% SDS-gel. Different percentage gels can be used if required. Add the radioactively labeled substrate to the gel solution at a level of 1.5  106 cpm/20 ml, equivalent to approximately 2 M pTyr. Mix by pipetting several times, then add TEMED and ammonium persulfate and mix again. Pour the gel behind a plexiglass shield, then allow it to polymerize for at least 4 hr at room temperature. Gel Running and Processing After solubilization in sample buffer, apply the IAA-treated cell lysates to the SDS-PAGE gel containing a radioactively labeled substrate and resolve the proteins by electrophoresis. After electrophoresis, place the gel in a glass tray containing fixation buffer (300 ml of Buffer G) and incubate on a rocker at room temperature overnight to remove SDS. Remove the isopropanol by washing the gel twice, each time with 250 ml of Buffer H at room temperature for 25 min, then denature the proteins by incubating the gel in a buffer containing guanidine–HCl (300 ml of Buffer I) at room temperature for 90 min. Wash the gel twice with 250 ml of Buffer J at room temperature for 1 hr each time to remove the guanidine–HCl, then initiate renaturation of the proteins by incubating the gel in 250 ml of Buffer J containing 3 mM DTT. After 1 hr place the gel in a fresh 250 ml aliquot of Buffer J containing 3 mM DTT and leave overnight at room temperature. During this time those PTPs that were initially oxidized in response to the cellular stimulus renature and, due to the presence of DTT in the buffer, are

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reduced back to an active state in which they are capable of dephosphorylating the 32P-labeled substrate that surrounds the protein band in the gel. This dephosphorylation reaction is terminated by Coomassie Blue staining and destaining of the gel, which should then be dried and exposed to film.

Stimulus-Induced Oxidation of PTPs

The production of intracellular ROS, such as H2O2, has been shown to be important for optimal tyrosine phosphorylation-dependent signaling.16,19 Interestingly, a variety of early studies highlighted the fact that addition of exogenous H2O2 is sufficient to augment tyrosine phosphorylation, for example, inducing cell proliferation or serving as an insulin-mimetic.20,21 This assay can be used to examine the effects of H2O2 on the oxidation of PTPs. As shown in Fig. 4, addition of extracellular H2O2 induces the

FIG. 4. Treatment of Rat-1 fibroblasts with H2O2 induced oxidation of multiple PTPs. Serum-starved Rat-1 cells were exposed to 200 M H2O2 for the indicated times. Cells were harvested in lysis buffer (Buffer A) containing 10 mM IAA under anaerobic conditions. Aliquots of lysates (25 g) were subjected to the in-gel PTP activity assay using poly Glu : Tyr, 4 : 1 as substrate.

19

M. Sundaresan, Z. X. Yu, V. J. Ferrans, K. Irani, and T. Finkel, Science 270, 296–299 (1995). D. Heffetz, I. Bushkin, R. Dror, and Y. Zick, J. Biol. Chem. 265, 2896–2902 (1990). 21 D. Heffetz, W. J. Rutter, and Y. Zick, Biochem. J. 288, 631–635 (1992). 20

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oxidation of multiple intracellular PTPs. Interestingly, this oxidation is required for the mitogenic effects of H2O2.16 The observation that multiple PTPs are oxidized in response to H2O2 is consistent with the fact that the signature motif, containing the unique active site Cys residue, is found in all PTPs, and that the members of this family share the same mechanism of catalysis. Therefore, it is possible that the regulation of PTP function by reversible oxidation may apply broadly across the whole family. In contrast to the oxidation of multiple PTPs in response to extracellular H2O2, the response to a physiological stimulus is more specific. Treatment of Rat 1 fibroblasts with PDGF led primarily to the transient oxidation of one PTP of apparent Mr  70 k (Fig. 5 and in

FIG. 5. Stimulation of Rat 1 fibroblasts with PDGF induced transient oxidation of SHP-2. (A) Serum-starved Rat-1 cells were exposed to 50 ng/ml PDGF-BB (R&D System, #220-BB) for the indicated times. Lysates were prepared in the absence (lane 1) or presence of 10 mM IAA (lanes 2–6) under anaerobic conditions, and subjected to in-gel PTP activity assay (25 g lysate protein per lane). The arrowhead indicates a PTP of apparent Mr  70 k that was transiently oxidized in response to PDGF stimulation. (B) The PDGF-stimulated Rat-1 cells were harvested in lysis buffer containing 10 mM IAA. Lysates (500 g) were incubated with various amounts of antibody to either SHP2 or SHP1, as indicated. Immunodepleted lysates were recovered and subjected to in-gel PTP activity assay (25 g lysate protein per lane). The arrowhead indicates the position of the 70 k PTP that was oxidized in response to PDGF (lane 1) and was immunodepleted from lysates with the anti-SHP-2 antibody (lanes 2–4), but not with antibody to SHP-1 (lane 5).

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Meng et al.16). Interestingly, the time course of oxidation of this PTP precisely matched the time course of autophosphorylation of the PDGF receptor following growth factor stimulation.16 With the availability of the human genome sequence and the definition of the complement of PTPs in humans, it is now possible to use the apparent molecular weight of the PTP in the ‘‘in-gel’’ assay to predict its identity. This prediction can then be tested by immunodepletion using antibodies to PTPs of that molecular weight. For example, in the case of PDGF stimulation, the 70 k PTP is immunodepleted from cell lysates with antibodies to the SH2 domain-containing PTP SHP2 (Fig. 5B). It is known that ligand-induced autophosphorylation of the PDGF receptor (PDGFR) generates docking sites for various signaling proteins, including SHP2. We have also shown that mutant forms of the PDGFR that were unable to bind to SHP2 displayed enhanced autophosphorylation and enhanced activation of MAP kinase.16 Thus, SHP2 appears to recognize the PDGFR as a substrate and functions as an inhibitor of PDGFR signaling. Interestingly, it was only the population of SHP2 that was bound to the PDGFR that was susceptible to reversible oxidation and inhibition. These observations led us to propose that PDGF stimulation induces localized production of ROS, leading to the rapid oxidation of the pool of SHP2 that has been recruited into a complex with the PDGFR. This augments autophosphorylation of the receptor and the initiation of the signaling response. The transient nature of the oxidation ensures reduction and reactivation of the pool of SHP2, which promotes dephosphorylation of the PDGFR and termination of the signal. Summary and Perspectives

The data derived from this assay illustrate how ligand-induced production of ROS may augment tyrosine phosphorylation-dependent signaling in general through inactivation of PTPs (Fig. 6). The production of ROS is observed in response to a wide variety of stimuli, including growth factors, hormones, cytokines, and activators of Gprotein coupled receptors, which lead to PTK activation. For example, in a more recent study we observed that insulin induced the transient oxidation of 2 PTPs, PTP1B, and the 45 kDa isoform of TC-PTP, both of which function as antagonists of insulin receptor signaling (T. C. Meng, D. A. Buckley and N. K. Tonks, unpublished observations). The general operating principle is that the stimulus enhances tyrosine phosphorylation directly, by activation of a PTK, and/or indirectly, by inactivation of a PTP. Thus, one function of ROS produced following agonist stimulation is to inactivate transiently the critical PTP that provides the inhibitory

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FIG. 6. Regulation of Protein Tyrosine Phosphatase (PTP) activity by reversible oxidation. A wide variety of physiological stimuli trigger the production of Reactive Oxygen Species (ROS), which may serve to fine-tune tyrosine phosphorylation dependent signaling pathways. For example, ligand-dependent activation of a receptor protein tyrosine kinase (RTK) triggers the activity of a Rac-dependent NADPH oxidase, leading to production of ROS. ROS oxidize the active site Cys residue of members of the PTP family, converting it from a thiolate ion (S
, the active form) to sulfinic acid (S-OH) and then a sulfenamide (-S-N-).10a,10b Oxidation results in inhibition of PTP activity, thereby promoting tyrosine phosphorylation. However, oxidation of the PTPs is transient. Restoration of PTP activity following reduction back to the thiolate form of the active site Cys residue terminates the tyrosine phosphorylation dependent signal. A variety of growth factors, hormones, and cytokines induce ROS production and stimulate tyrosine phosphorylation. We are developing methods to identify the PTPs that become oxidized in response to a physiological stimulus as a way of establishing links between particular PTPs and the regulation of defined signaling pathways. (See Color Insert.)

constraint upon the system, thus facilitating the initiation of the signaling response to that stimulus. We reasoned that by identifying which PTPs are oxidized in response to a particular stimulus, we would reveal those PTPs that are critical for downregulating the signaling response to that stimulus. Therefore, stimulus-induced oxidation may be used as a means of ‘‘tagging’’ and identifying those PTPs that are integral to the regulation of the signaling events triggered by a defined stimulus. It is important to note that some PTPs, in particular the receptor-like enzymes, do not renature efficiently in SDS-PAGE gels and, therefore, would not be easily detected in the ‘‘in-gel’’ assay. It will be important to develop immobilized, broad specificity, active site-directed inhibitors (such as phenyl arsine oxide coupled to a support) as affinity matrices to substitute for the SDS-PAGE gels as a means of isolating the oxidized PTPs in this strategy. This would also facilitate the application of mass spectrometry-based sequencing methods to the identification of such

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PTPs. The application of state-of-the-art technologies, such as RNA interference and the use of substrate-trapping mutant PTPs, would allow us then to define the function of the PTPs that were identified in this way. Hopefully this will provide further insights into the physiological function of members of the PTP family and potentially lead to the identification of novel targets for therapeutic intervention in diseases associated with dysfunctional tyrosine phosphorylation-dependent signaling.

Section IV Expression Systems

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[24] Preparation and Characterization of Recombinant Protein Phosphatase 1 By TAKUO WATANABE, EDGAR F. DA CRUZ E SILVA, HSIEN-BIN HUANG, NATALIA STARKOVA, YOUNG-GUEN KWON, ATSUKO HORIUCHI, PAUL GREENGARD, and ANGUS C. NAIRN

Protein phosphatase-1 (PP1) is a major eukaryotic serine/threonine protein phosphatase that regulates such diverse cellular processes as cell cycle progression, protein synthesis, muscle contraction, carbohydrate metabolism, transcription, and neuronal signaling.1–4 It appears that little of the free PP1 catalytic subunit (
37 kDa, PP1C) exists in cells. Rather, the precise role played by PP1 in its diverse functions is attributed to its interaction with a large variety of regulatory subunits that include both inhibitor and targeting proteins. Inhibitors include protein inhibitor-1, its homologue DARPP-32 (dopamine- and cAMP-regulated phosphoprotein, Mr 32,000), and inhibitor2. Phosphorylation of inhibitor-1 at Thr35, or of DARPP-32 at Thr34, by protein kinase A (PKA), is required for PP1 inhibition. In contrast, unphosphorylated inhibitor-2 interacts with PP1C. A growing number of targeting subunits have also been identified that localize PP1C to specific subcellular compartments, thereby influencing its substrate specificity and local function. These include glycogen-targeting subunits, myofibrillartargeting subunits, several nuclear-targeting proteins, and proteins such as spinophilin and neurabin that target PP1C to dendritic spines of neurons. Many of the proteins that interact with PP1C share a common binding, or docking, site, that comprises a docking motif containing one or more basic amino acid followed by two hydrophobic residues separated by a variable amino acid (the so-called RVXF motif).5 X-ray crystallographic analysis has indicated that the docking motif interacts in an extended manner with a hydrophobic channel in PP1C situated on the side opposite that of the active site. Thus, while more complex oligomeric structures may exist, PP1C interacts with its regulatory subunits in a mutually exclusive manner. 1

S. Shenolikar, Annu. Rev. Cell Biol. 10, 55–86 (1994). P. Greengard, P. B. Allen, and A. C. Nairn, Neuron 23, 435–447 (1999). 3 J. B. Aggen, A. C. Nairn, and R. Chamberlin, Chem. Biol. 7, R13–R23 (2000). 4 M. Bollen, Trends Biochem. Sci. 26, 426–431 (2001). 5 M. P. Egloff et al., EMBO J. 16, 1876–1887 (1997). 2

METHODS IN ENZYMOLOGY, VOL. 366

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Despite the recent advances that have been made in the biochemical characterization of PP1, these studies have been hampered by a lack of easily prepared recombinant PP1C. It has been impossible to overexpress PP1C in mammalian cells, presumably due to the key role it plays in cellular function. We have therefore analyzed various methods for preparation of PP1C in bacteria, and have developed methods for preparation of PP1C in Sf9 cells using the baculovirus expression system. The preparation and characterization of these different recombinant PP1C preparations is discussed.

Materials

Oligonucleotides were synthesized by Operon Technologies, Inc. (Berkley, CA). The QuickChange site-directed mutagenesis kit was from Stratagene. The Bac-to-Bac Baculovirus Expression System including the vector plasmid pFastBac-HT, DH10BAC competent cells, and recombinant TEV protease were from Life Technologies (Gaitherburg, MD). Sf9 cells were from Novagen (Madison, WI). CompleteTM protease inhibitor cocktail tablets and protease inhibitor E-64 were from Boehringer-Mannheim (Indianapolis, IN). NHS-Hi Trap and glutathione-Sepharose were from Pharmacia Biotech (Uppsala, Sweden). Ni-nitrilotriacetic acid (NTA) agarose was from Qiagen (Valencia, CA). Immobilon-P was from Millipore (Bedford, MA). PNUTS peptide was synthesized and purified by reversedphase column HPLC at W. M. Keck Foundation Biotechnology Resource Laboratory, Yale University. The Protein Tyrosine Phosphatase Assay System was from New England Biolabs (NEB). Monoclonal antibody for PP1 (E-9) was from Santa Cruz Biotechnology Inc. (Santa Cruz, CA). Native PP1C was purified from rabbit skeletal muscle, using sequential chromatography on DEAE-cellulose, heparin-Sepharose, phenyl-Sepharose, Sephacryl S-200 and Mono-Q resins (largely as described below for recombinant PP1C, see also Ref. 6). DARPP-32 and inhibitor-2 were prepared from E. coli as described.7 GST-tagged spinophilin and GSTPNUTS were prepared from E. coli as described.8,9 6

H. B. Huang, A. Horiuchi, J. Goldberg, P. Greengard, and A. C. Nairn, Proc. Natl. Acad. Sci. U.S.A. 94, 3530–3535 (1997). 7 H.-B. Huang et al., J. Biol. Chem. 274, 7870–7878 (1999). 8 P. B. Allen, Y. G. Kwon, A. C. Nairn, and P. Greengard, J. Biol. Chem. 273, 4089–4095 (1998). 9 L. C. Hsieh-Wilson, P. B. Allen, T. Watanabe, A. C. Nairn, and P. Greengard, Biochemistry 38, 4365–4373 (1999).

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Preparation of Phosphorylated and Thiophosphorylated DARPP-32

Phosphorylated recombinant DARPP-32 was prepared essentially as described.7 DARPP-32 (2 mg) was dissolved in 1 ml of 50 mM Hepes (pH 7.4), 1 mM EGTA, 10 mM magnesium acetate, and 1 mM ATP. The phosphorylation reaction was started by addition of 2 g of PKA catalytic subunit and the incubation was carried out for 60 min. For preparation of [32P]DARPP-32, [32P]ATP replaced ATP. For preparation of thiophosphorylated DARPP-32, 1 mM thiophospho-ATP (Boehringer Mannheim) replaced ATP, and the reaction was carried out at 30 C for 5 days, with fresh PKA (2 g) and thiophospho-ATP (1 mM final concentration) added every 24 hr. Phosphorylated DARPP-32 and thiophospho-DARPP-32 were purified by HPLC using a C-18 column. Preparation of Phosphorylase a

Phosphorylase b (93 mg of lyophilized powder; Sigma) was phosphorylated with phosphorylase kinase (2 mg, Sigma) in 3 ml of reaction buffer containing 0.2 mM ATP, [ -32P]ATP (3000 Ci/mmol, Amersham), 100 mM Tris–HCl, pH 8.2, 100 mM sodium glycerol-1phosphate, 0.1 mM CaCl2 and 10 mM magnesium acetate. Microcystin (final concentration, 100 nM) was added to inhibit endogenous protein phosphatases. The reaction mixture was incubated for 2 hr at 30 C, then 5.45 ml of 70% ammonium sulfate was added (to 45% of saturation). The reaction mixture was put on ice for 30 min and then centrifuged at 19,000g for 15 min. The pellet was resuspended in 240 l of 10 mM Tris–HCl, pH 7.5, 0.1 mM EGTA, 10% glycerol and 436 l of 70% ammonium sulfate was added (to 45% of saturation). The mixture was incubated on ice for another 30 min and centrifuged at 19,000g for 15 min. The pellet was resuspended in 600 l of 10 mM Tris–HCl, pH 7.5, 0.1 mM EGTA, 10% glycerol, transferred to a dialysis tube and dialyzed at 4 C against 10 mM Tris–HCl, pH 7.5, 0.1 mM EGTA (at least four changes of 1 liter each). Precipitated phosphorylase a and solution from the dialysis tube were transferred to a microcentrifuge tube and incubated on ice for 2 hr. After centrifugation at 19,000g for 15 min, the pellet was resuspended in 600 l of cold 10 mM Tris–HCl, pH 7.5, 0.1 mM EGTA, 15 mM 2-mercaptoethanol and stored at 4 C. Dephosphorylation of 32

32

P-phosphorylase a or

32

P-DARPP-32

P-labeled phosphorylase a (10 M) or phospho-Thr34-DARPP-32 (0.5–20 M) was incubated with PP1 in a reaction mixture (40 l)

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containing 50 mM Tris–HCl, pH 7.0, 300 mM NaCl, and 0.3 mg/ml bovine serum albumin (BSA). The detergent Brij 35 (0.01%, w/v) was often added to the incubation mixture. Reactions were performed at 30 C for 10 min and stopped by adding 200 l of 20% (phosphorylase) or 50% (DARPP-32) trichloroacetic acid (TCA). Samples were centrifuged at 19,000g for 10 min (for DARPP-32, samples were incubated on ice for 1 h), and 32P in the supernatant was analyzed by Cerenkov counting in a scintillation counter (Beckman). Unless indicated, assays of E. coli PP1 contained 1 mM MnCl2. For assay of inhibitors, PP1 and inhibitors were preincubated for 15–30 min on ice.

Dephosphorylation of p-nitrophenyl Phosphate ( pNPP) and Tyrosine-phosphorylated Myelin Basic Protein

Phosphatase activity was measured in a reaction mixture (0.1 ml) containing 13 mM pNPP, 10 mM HEPES, pH 7.0, and 300 mM NaCl. Reactions were performed at 30 C for 20 min and stopped by adding 1 ml of 0.2 N NaOH. Phosphate release from the reaction product ( p-nitrophenol) was analyzed by measurement of absorbance at 405 nm using a molar extinction coefficient of 18,000 M1 cm1. The rate of nonenzymatic hydrolysis of the substrate was corrected by measuring the optical density in the absence of PP1. [32P]myelin basic protein was phosphorylated by Abl protein tyrosine kinase and tyrosine phosphatase activity was assayed using the Protein Tyrosine Phosphatase Assay System as described by the manufacturer (NEB).

Expression of PP1 in E. coli

Rabbit PP1 cDNA was a generous gift from N. Berndt. PP1 was expressed in E. coli and purified using two different methods. Method 1

The rabbit PP1C cDNA was contained within a pDR540 plasmid containing a trp-lac fusion promoter that preceded the PP1 DNA. E. coli (DH5), harboring the wild-type rabbit PP1 cDNA, were grown in 5 liters of LB medium containing 1 mM MnCl2 and ampicillin (0.1 mg /ml) at 37 C with shaking at 250 rpm. After 4 hr incubation (an OD600nm of
0.6), 0.3 mM isopropyl- -D-thiogalactopyranoside (IPTG) was added to induce the expression of PP1C. Cells were incubated for up to an additional 13 hr,

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and then centrifuged at 4000g for 30 min. The pellet was resuspended in 300 ml of 20 mM Tris–HCl buffer (pH 7.5) containing 1 mM MnCl2, 0.2 mM PMSF, 4 mM benzamidine, 15 mM 2-mercaptoethanol, 0.1 mM EGTA, pepstatin A (10 g/ml), leupeptin (10 g/ml) and chymostatin (10 g/ml). Cells were lysed using a French press (1000–1500 psi). The lysate was centrifuged at 20,000g for 20 min, and the supernatant was loaded onto a heparin-Sepharose column (1.5  20 cm), followed by washing with one column volume of buffer A (20 mM Tris–HCl, 1.0 mM MnCl2, 0.1 mM EGTA, 4.0 mM benzamidine, 0.2 mM PMSF and 15 mM 2-mercaptoethanol, pH 7.5) plus 0.1 M NaCl. Proteins were eluted with a linear gradient (500 ml total volume) from 0.1 to 0.6 M NaCl (in buffer A). Fractions containing PP1 activity were pooled and concentrated to 15 ml by ultrafiltration using an Amicon YM-10 membrane. One volume of buffer A containing 3.4 M NaCl was added to the pooled sample to give a final concentration of 1.7 M NaCl. The sample was loaded onto a phenyl-Sepharose column (1.5  18 cm), the column washed with 40 ml of buffer A containing 1.7 M NaCl, and proteins eluted using a linear gradient (500 ml total volume) from 1.7 to 0.5 M NaCl (in buffer A containing 10% glycerol). Fractions containing PP1 activity were pooled and concentrated to 5 ml by ultrafiltration. The concentrated sample was loaded on a Sephacryl S-200 column (2.5  120 cm) equilibrated with 20 mM triethanolamine, pH 7.0, 0.1 mM EGTA, 1.0 mM MnCl2, 4.0 mM benzamidine, 0.2 mM PMSF, 10% glycerol, 0.3 M NaCl, 15 mM 2mercaptoethanol. Fractions containing PP1 activity were pooled and concentrated to 4 ml by ultrafiltration. The sample was diluted by addition of 10 ml of buffer B (20 mM triethanolamine, 0.1 mM EGTA, 10% glycerol, 1 mM MnCl2, 0.1% 2-mercaptoethanol, pH 7.0) to give a final concentration of NaCl of less than 0.1 M. The diluted sample was passed through a 0.45 m filter, and loaded onto a Mono-Q anion exchange column equilibrated in buffer B. The column was washed and proteins eluted using a linear gradient of 0.1–0.4 M NaCl (in buffer B) (1 ml/min over 50 min). PP1C was eluted at
0.22 M NaCl and was stored in 50% glycerol at 20 C. Method 2

PP1C DNA was subcloned from pDR540 (Pharmacia) into pET28a (Novagen) to produce a recombinant enzyme with a His6-tag at the Nterminus. However, the T7 promoter from pET28 proved to be too strong: large amounts of PP1 were synthesized but most of the protein was found to be insoluble. The His6-PP1 was therefore cloned back into pDR540 under control of the tac promoter. BL21 cells were then transformed with the

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pDR540 plasmid harboring His6-PP1C. Transformed cells were grown at room temperature (
22–23 C) in LB medium containing 100 g/ml ampicillin and either 1 mM MnCl2 or 100 M CoCl2. At OD600, protein expression was induced with 0.3 mM IPTG and cells were further incubated overnight at room temperature. The cells were harvested by centrifugation, resuspended in buffer C (10 mM Tris–HCl, pH 8.0, 30 mM imidazole, 300 mM NaCl, 10% glycerol, CompleteTM protease inhibitor cocktail, and either 1 mM MnCl2 or 1 mM CoCl2) and lysed using a French press. Cell debris was removed by centrifugation at 35,000g for 15 min, and the supernatant was clarified by centrifugation at 35,000g for 45 min. The resulting supernatant was loaded on a Ni-NTA-agarose column (volume, 3 ml) at a rate of 1 ml/min; the column was washed with buffer C, and His6-PP1C was eluted with an increasing linear gradient of imidazole (30–500 mM) in buffer C (total gradient volume, 30 ml). The fractions were collected and analyzed by SDS-PAGE. Fractions containing His6-PP1 were pooled and stored in small aliquots at 80 C.

Preparation of PP1 in Sf9 Cells

Two different procedures were used to prepare recombinant PP1 using the baculovirus method.

Method 1

Rat PP1C or human PP1C cDNAs was subcloned into pBlueBac (Invitrogen) and recombinant transfer plasmids were transfected into Sf9 cells with linearized AcMNPV DNA (Invitrogen) using lipofectin (PP1C) or into Sf9 cells with BaculoGold (Pharmingen) viral DNA (PP1C ). The medium containing the recombinant virus was collected after 5–7 days. Individual recombinant viral clones were isolated after plaque purification in the presence of Bluo-Gal. High-titre stocks of virus were generated in spinner cultures of Sf9 cells at a low multiplicity of infection (MOI
0.1 pfu/cell). For large scale purification, Sf9 cells (107 cells/ml) were infected with recombinant virus at a MOI of 1–2 for 1 hr at room temperature, followed by dilution with Grace’s medium (without or with 1 mM MnCl2) to give a final cell density of 106/ml. Infected cells were grown in spinner flasks at 27 C with constant stirring at 50–60 rpm. The cells (
1.5 liter) were incubated for 48 hr, and then were harvested by centrifugation at 1000g for 3 min. Cells were washed with 20 mM Tris–HCl, pH 7.5, 0.1 mM EGTA,

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15 mM 2-mercaptoethanol, 0.25 M sucrose, resuspended in 20 mM Tris– HCl, pH 7.5, 2 mM EGTA, 2 mM EDTA, 15 mM 2-mercaptoethanol, 0.2 mM PMSF, and lysed by ultrasonication. The lysate was centrifuged at 20,000g for 20 min, and the supernatant was purified by chromatography using DEAE-cellulose (3  25 cm column, linear gradient of 0.08–0.5 M NaCl), heparin-Sepharose and Sephacryl S-200 (as described above for E. coli, Method 1). In some cases, chromatography on Mono-Q resin was carried out as described above. Sf9 PP1C prepared by Method 1 was stored in 50% glycerol at 20 C. If PP1C was expressed with MnCl2 in the incubation medium, 1 mM MnCl2 was also added to all buffers during purification.

Method 2

EcoRI and XbaI restriction sites were added in the 50 end and 30 end of the PP1C DNA, and the DNA was inserted in-frame into pFastBac-HT. For some constructs, cDNA encoding glutathione S-transferase (GST), followed by a TEV protease cleavage site and a FLAG tag, was introduced in-frame between the His6 tag and PP1C sequences. Thus, proteins had either a His6 tag, with a TEV protease cleavage site, or alternatively a His6 tag, a TEV cleavage site, a GST tag, a second TEV cleavage site and a FLAG tag at the N-terminus of PP1C. Recombinant baculovirus stocks were prepared as described by the manufacturer (Life Technologies). In brief, donor plasmids were used for transformation of competent DH10BAC E. coli cells. Recombinant bacmid DNA was isolated and used for transfection of Sf9 cells. After 48 hr incubation at 28 C, supernatant from the culture medium was saved and used for further amplification. For preparation of PP1C, Sf9 cells were grown in suspension culture (28 C, rotating at 135 rpm) in 100 ml of Grace’s insect medium [supplemented with 0.33% yeastolate, 0.33% lactalbumin hydrolysate and 10% (v/v) fetal calf serum] in a 500 ml flask with a ventilation cap (note that Life Technologies serum-free medium, Sf-900 II SFM, should not be used since this results in a Sf9 PP1C preparation that dephosphorylates to some extent phospho-DARPP-32 and tyrosine-phosphorylated myelin basic protein). Sf9 cells (1.5  106 cells/ml) were infected at a MOI between 1 and 2. Cells were then incubated for 72 hr at 28 C, rotating at 135 rpm. Cells were then centrifuged at 500g for 5 min, and the pellets lyzed with gentle swirling in 5 ml (
2  107 cells/ml) of 50 mM Tris– HCl, pH 7.5, 100 mM NaCl (or 100 mM KCl), 1% NP40, 15 mM 2mercaptoethanol, 1 mM AEBSF, 30 g/ml E-64, 5 g/ml pepstatin A, 5 g/ml leupeptin, 5 g/ml chymostatin. Supernatants were prepared by

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centrifugation at 100,000g for 30 min and purified by affinity chromatography as described below.

Affinity Chromatography

Several different affinity chromatography procedures were used that utilized the His6 and GST tags attached to recombinant PP1. In addition, PP1 was also purified by affinity chromatography using a docking motif peptide derived from PNUTS (PNUTS[392-408]). In some cases more than one affinity chromatography step was used in a purification procedure. The supernatant obtained after cell lysis was typically diluted in appropriate binding buffer and filtered through a 0.45 m syringe-driven filter unit. For purification of His6-PP1C, filtered supernatant was incubated with Ni-NTA agarose (1–2 ml, Ni-NTA agarose resin, QIAGEN or equivalent) and the agarose slurry transferred into a column. The resin was washed with buffer containing 20 mM Tris–HCl, pH 8.5, 0.5 M KCl, 0.1 mM EGTA, 10% glycerol, 5 mM 2-mercaptoethanol and 20 mM imidazole, then washed with a buffer containing 20 mM Tris–HCl, pH 8.5, 1 M KCl, 0.1 mM EGTA, 10% glycerol, 5 mM 2-mercaptoethanol. Proteins were eluted with buffer containing 20 mM Tris–HCl, pH 8.5, 0.1 M KCl, 0.1 mM EGTA, 10% glycerol, and 150 mM imidazole. Fractions containing active PP1C were dialyzed against buffer containing 50 mM Tris–HCl, pH 7.0, 50% glycerol, 0.1 mM EGTA, and 15 mM 2mercaptoethanol. For purification of GST-tagged PP1C, cells (
3  107 cells per ml) were lysed with buffer containing 50 mM Tris–HCl, pH 7.5, 0.1 M NaCl and 1% NP-40, 15 mM 2-mercaptoethanol, 1 mM AEBSF, 30 g/ml E-64, for 30 min on ice. [In some cases, CompleteTM (EDTA-free) protease inhibitor cocktail tablets (Boehringer-Mannheim), replaced the protease inhibitors.] The supernatant (centrifugation at 100,000g for 30 min) was incubated with
1/10th volume of glutathione-Sepharose resin for 1 hr at 4 C, and the resin was pelleted by centrifugation and washed three times with phosphate-buffered saline. The resin was then resuspended in 50 mM Tris–HCl, pH 8.0, 10 mM EDTA, 1 mM DTT, 10% glycerol, and incubated with 200 units of His6-tagged TEV protease for 2 hr at 4 C. After cleavage, Ni-NTA agarose was added to the suspension for 1 hr at 4 C to remove the His6-tagged TEV protease. The suspension was centrifuged and the supernatant was used for protein phosphatase assay. The synthetic peptide PNUTS [392-408] (KGRKRKTVTWPEEGKLR, 1–2 mg) was coupled to HiTrap-NHS resin (1 ml) (Amersham Biosciences) according to the manufacturer’s instructions. The coupling efficiency

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(calculated based on the absorbance at 280 nm of the applied sample and first wash sample) was typically >80%. The PNUTS peptide resin was packed in a Hi-Trap column and attached to a FPLC (Pharmacia or equivalent), and the column equilibrated with 5 ml of buffer containing 20 mM triethanolamine, pH 7.0, 50 mM NaCl, 0.1 mM EGTA, 10% glycerol and 15 mM 2-mercaptoethanol, at a flow rate of 0.5 ml/min. The diluted cell lysate was applied to the column at the same flow rate. (Note that the flowthrough can be collected and reloaded onto the column.) After loading, the column was washed with 10 ml of buffer, then PP1C was eluted with a 19.5 ml linear gradient from 0.05 to 2 M NaCl in column equilibration buffer. Fractions of 1 ml were collected, and analyzed for PP1C activity or protein by enzyme assay or immunoblotting. Active PP1C eluted between 0.6 M and 2 M NaCl. (Note that immunologically cross-reactive PP1C that has low phosphatase activity is eluted in fractions between 0.2 and 0.4 M NaCl.) Fractions containing PP1C were dialyzed against 50 mM triethanolamine, pH 7.0, 0.1 mM EGTA, 50% glycerol, 15 mM 2-mercaptoethanol, 0.1 mM PMSF at 4 C overnight, and stored at 70 C until use. In some cases, the PNUTS affinity step was combined with affinity chromatography of His6-PP1 using Ni-NTA. In this case, the sample was applied first to the Ni-NTA column as described above (except that 50 mM triethanolamine, pH 7.0, replaced 20 mM Tris–HCl, pH 8.5). His6-PP1C was eluted with buffer containing 150 mM imidazole (as described above), and fractions containing PP1C activity were loaded directly onto the PNUTS peptide affinity column. (Note that the PNUTS affinity method must only be used after purification on Ni-NTA or glutathione-Sepaharose to avoid contamination with endogenous Sf9 cell PP1.) Although the His6 tag has no apparent influence on PP1C activity, this can be removed using TEV protease (Invitrogen) that is also polyhistidine tagged. The released tagged fragment and the TEV protease can then be removed from the PP1 by chromatography on Ni-NTA resin.

Immunoblotting of Recombinant PP1C

Proteins were separated by SDS-PAGE and electrophoretically transferred to Immobilon-P membrane using standard procedures. Membranes were incubated in PBS containing 0.1% Tween 20, 0.5% nonfat dry milk and 1 g/ml of anti-PP1C monoclonal antibody (E-9). Rabbit anti-mouse IgG antibody and 125I-labeled Protein A were used for detection of signal. Radioactivity was quantified using a PhosphorImager (Molecular Dynamics) and purified recombinant PP1C of known concentration was used as standard.

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Characterization of PP1C Expressed in E. coli using Method 1

PP1C has been expressed in E. coli and purified by several investigators using either conventional or affinity chromatography.6,10–14 This type of preparation has been used in many structural and biochemical studies of PP1C (including most studies of PP1C mutants), and was the preparation used in production of protein for X-ray crystallization studies.15,16 Recombinant E. coli PP1C is available commercially from several sources including NEB and Roche. Approximately 1 mg/liter of purified PP1C can be obtained after
460-fold purification with a yield of
50% from the cytosolic fraction of lysed E. coli (Table I). A major limitation in the yield is related to the amount of insoluble PP1C protein that is expressed. Reduced temperature of incubation, or shorter times of induction with IPTG may help to increase the relative expression level of soluble PP1C. Addition of 1 mM MnCl2 to the culture medium, and to buffers during the purification, is essential for preparation of active PP1C.10,11 As shown in several studies, E. coli PP1C exhibits a high specific activity comparable with native PP1C (usually prepared from rabbit skeletal muscle as a mixture of isoforms) [>35 units/mg under standard assay

PURIFICATION Purification step Crude lysate Heparin-Sepharose Phenyl-Sepharose Sephacryl S-200 Mono-Q

10

OF

TABLE I PP1C FROM E.

COLI

USING METHOD 1

Volume (ml)

Protein (mg)

Total activity (units)

Specific activity (units/mg)

Purification (-fold)

Yield (%)

287 130 107 56 7

3961 83.2 ND 8.4 4.1

312 266 ND 194 152

0.08 3.2 ND 23.1 36.9

1 40 ND 289 461

100 85 ND 62 49

Z. Zhang, G. Bai, S. Deans-Zirattu, M. F. Browner, and E. Y. C. Lee, J. Biol. Chem. 267, 1484–1490 (1992). 11 D. R. Alessi, A. J. Street, P. Cohen, and P. T. W. Cohen, Eur. J. Biochem. 213, 1055–1066 (1993). 12 Z. Zhang, S. Zhao, S. D. Zirattu, G. Bai, and E. Y. C. Lee, Arch. Biochem. Biophys. 308, 37–41 (1994). 13 J. Zhang, Z. J. Zhang, K. Brew, and E. Y. C. Lee, Biochemistry 35, 6276–6282 (1996). 14 J. H. Connor et al., J. Biol. Chem. 273, 27716–27724 (1998). 15 J. Goldberg et al., Nature 376, 745–753 (1995). 16 M. P. Egloff, P. T. W. Cohen, P. Reinemer, and D. Barford, J. Mol. Biol. 254, 942–959 (1995).

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conditions (Table I); Vmax of > 60 mol/min/mg and Km
10 M using phosphorylase a as substrate].6 The E. coli enzyme is also normal in terms of inhibition by microcystin, calyculin A and okadaic acid (see Table IV) and has therefore been very useful in structure/function studies related to these inhibitors.6,17 However, E. coli PP1C exhibits several anomalous properties. These include the fact that it is dependent on added Mn2 þ for activity, and is able to dephosphorylate phospho-tyrosine-containing substrates and para-nitrophenyl phosphate.6,11,18,19 E. coli PP1C is also relatively insensitive to inhibition by phospho-inhibitor-1 and phosphoDARPP-32, an effect that can be partly explained by the ability of the preparation to dephosphorylate either phospho-inhibitor protein (see Table IV, Ref. 6 and data not shown). Several of the anomalous properties of E. coli PP1C may be explained by a subtle change in the structure of the metal-binding ligands in the active site of the enzyme.6,16,20,21 Thus active E. coli PP1C contains two Mn2 þ ions in its active site rather than Fe2 þ and Zn2 þ , likely present in native PP1C. Subtle changes in the position of Tyr272 that overhangs the phosphate-binding site in the active site, also is the likely cause of the ability of the enzyme to dephosphorylate phospho-inhibitor-1, phosphoDARPP-32, pNPP, and phospho-tyrosine-containing substrates.21 It is notable that E. coli PP1C also exhibits a marked ability to autodephosphorylate phospho-Thr320 when incubated with cdc2 kinase, in contrast to native or Sf9 PP1C (Fig. 1 and see below). In addition to the abnormal catalytic properties, E. coli PP1C exhibits altered interaction with its regulatory subunits that are also likely to be caused by subtle structural changes, perhaps in the region of the protein that binds the docking motif. For example, E. coli PP1C is still relatively insensitive to inhibition by either thiophospho-DARPP-32 (Table IV) or thiophospho-inhibitor-1 (both of which are resistant to the dephosphorylation by PP1). E. coli PP1C is also relatively insensitive to regulation by spinophilin and PNUTS. For example, GST-PNUTS[382– 486] inhibits Sf9 PP1C with an IC50 value of
0.1 nM, but inhibits E. coli PP1C with an IC50 value of
80 nM (data not shown). E. coli PP1C appears to be inhibited normally by full-length inhibitor-2, but displays anomalous regulation by various fragments of inhibitor-2 (Table II). For example, deletion of up to 105 amino acids at the C-terminus of 17

L. F. Zhang, Z. J. Zhang, F. X. Long, and E. Y. C. Lee, Biochemistry 35, 1606–1611 (1996). C. MacKintosh et al., FEBS Lett. 397, 235–238 (1996). 19 S. Endo et al., Biochemistry 36, 6986–6992 (1997). 20 D. Barford, Trends Biochem. Sci. 21, 407–412 (1996). 21 T. Watanabe et al., Proc. Natl. Acad. Sci. U.S.A. 98, 3080–3085 (2001). 18

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FIG. 1. In vitro phosphorylation of PP1C expressed in E. coli or Sf9 insect cells by cdc2/ cyclin B. E. coli PP1C (800 nM, upper panel) and Sf9 PP1C (400 nM, lower panel) were phosphorylated at Thr320 for various times at 30 C by purified cdc2/cyclin B (NEB) with [ -32P]ATP in the absence or presence of 1 M microcystin. Proteins were separated by SDSPAGE (10% acrylamide) and [32P]-labeled PP1C was detected by autoradiography. In the absence of microcystin, phosphorylation of E. coli PP1C rapidly reached a steady-state where phosphorylation by cdc2 was balanced by autodephosphorylation. When autodephosphorylation was blocked by microcystin, phosphorylation by cdc2 kinase reached a maximal level that reflected stoichiometric phosphorylation. In contrast, phosphorylation of Sf9 PP1C by cdc2 kinase was not affected by addition of microcystin.

REGULATION

OF

TABLE II E. COLI PP1C AND NATIVE PP1C MUTANT INHIBITOR-2

BY

WILD-TYPE AND

Relative inhibitory activity Inhibitor Full-length I2[1–204] I2[1–180] I2[1–172] I2[1–164] I2[1–159] I2[1–140] I2[1–120] I2[1–99] I2[1–84] I2[9–204] I2[14–204] I2[19–204] I2[I10G] I2[K11E] I2[I13G]

E. coli PP1C

Native PP1C

100% (IC50 1.1 nM) 31.0 48.0 19.0 6.0 7.0 7.0 8.0 4.0 37.0 0.2 0.4 26.0 3.0 1.0

100% (IC50 1.0 nM) ND ND ND ND 50.0 37.0 38.0 7.1 63.0 0.1 0.1 21.0 2.2 0.9

Results for native PP1C are summarized from Huang et al.7 ND, not determined.

inhibitor-2 had little effect on the IC50 for inhibition of native PP1C. However, removal of 40–50 amino acids at the C-terminus of inhibitor-2 rendered the proteins poor inhibitors of E. coli PP1C. This result presumably highlights, in the altered E. coli enzyme preparation, a specific

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333

deficit in one or more of the multiple binding interactions that normally occur with inhibitor-2 and native PP1C.7,22,23 In summary, E. coli PP1C prepared by this standard method is easily obtained in high quantity, is highly active, and is useful for some structure/ function studies. However, as a result of its anomalous enzymatic properties, this preparation is less useful in studies of physiological substrates and of interactions with targeting and inhibitor proteins.

Characterization of PP1 Expressed in E. coli Using Method 2

Previous studies have included examination of the ability of other divalent cations to substitute for Mn2 þ in supporting the activity of E. coli PP1C preparations initially purified in the presence of Mn2 þ .6,24 These studies are possible since at least one of the two Mn2 þ ions required for PP1C activity can be removed by dialysis in the presence of EDTA. The best substitute was found to be Co2 þ , which could form a stable complex with PP1C and activate the enzyme. Based on these results, it seemed possible that if added to the culture medium in place of Mn2 þ , Co2 þ might form a stable complex with PP1C as it was being expressed and folded, and that this preparation might resemble native PP1C. To test this possibility, His6tagged PP1C was expressed in E. coli in the presence of either 1 mM MnCl2 or 100 M CoCl2. In these studies, cells were grown at room temperature, which was important for increasing the level of soluble PP1C. In addition, it was observed that concentrations of CoCl2 higher than 100 M inhibited cell growth. Using a one-step affinity chromatography step, PP1C of greater than 95% purity could be obtained rapidly in the presence of either Mn2 þ or Co2 þ , and approximately similar amounts of PP1C were obtained (yield of 3–5 mg/liter culture). The activity of the Mn-PP1C preparation could be increased
1.5-fold (added 1 mM Mn2 þ ) or
2-fold (added 1 mM Co2 þ ) when divalent cations were added to the assay. In contrast, the CoPP1C preparation was inhibited 20–30% by addition of 1 mM Mn2 þ or Co2 þ . The maximum specific activities of the two preparations were approximately the same (
3–8 mol/min/mg under standard assay conditions). Both preparations also showed similar sensitivity to inhibition by okadaic acid, while as shown previously6,24 Mn-PP1C but not CoPP1C could be inhibited by addition of 1 mM EDTA. However, 22

I.-K. Park and A. A. DePaoli-Roach, J. Biol. Chem. 269, 28919–28928 (1994). J. H. Connor et al., J. Biol. Chem. 275, 18670–18675 (2000). 24 Y. F. Chu, E. Y. C. Lee, and K. K. Schlender, J. Biol. Chem. 271, 2574–2577 (1996). 23

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Co-PP1C still displayed anomalous properties in that it dephosphorylated pNPP (specific activity was
50% of that of Mn-PP1C), and could dephosphorylate phospho-Thr34-DARPP-32 (Vmax 0.24 mol/min/mg and Km of 0.6 M compared to Mn-PP1C which had a Vmax of 1.42 mol/ min/mg and a Km of 3.3 M). Thus, while Co-PP1C exhibits properties that are closer to those of native PP1C, the anomalous activities still make it of limited use in many studies of PP1C function. Characterization of PP1 Expressed in Sf9 Cells Using Method 1

Given the anomalous properties of the E. coli PP1C preparations, an alternative procedure was developed using Sf9 cells and the baculovirus method. Purification procedures were developed for two different isoforms, PP1C and PP1C . The optimal time for maximal expression of soluble, active PP1 was established using small-scale cultures, and analysis of membrane and cytosol fractions using phosphatase assays and immunoblotting (data not shown). For PP1C, significant expression of active soluble phosphatase was observed after 30 hr, expression peaked between 50 and 60 hr, and decreased thereafter. At early time points (up to 36 hr), PP1C was mostly found in the cytosol (>60%), but thereafter PP1C was mostly found as an inactive species in the particulate fraction. For PP1C , the time course of expression of active enzyme was similar to that of PP1C, but >70% of the protein was found in the cytosol at all time points. Notably, the activity of PP1C in the soluble fraction did decrease after
60 hr, but the reason for this loss of activity of soluble enzyme was not apparent. Varying the MOI did not have a significant effect on the relative ratios of soluble and particulate PP1C or . Similar patterns of expression were also observed when Sf21, Hi5, or Mg-1 cells were used in place of Sf9 cells (data not shown, note that the amount of PP1C expressed per cell varied somewhat depending on cell size). The purification procedure for untagged PP1C or was very similar to that used for purification of untagged E. coli PP1C (method 1) (Table III shows typical results for PP1C; very similar results were obtained for PP1C ). Approximately 0.7 mg of PP1C (>90% purity) can be obtained after
200-fold purification with a yield of
25% from the cytosolic fraction of lysed Sf9 cells (from
1 liter of culture medium). Sf9 cell PP1C and exhibited high specific activities [up to 20 units/mg under standard assay conditions (Table III and data not shown)]. The two enzyme preparations were also normal in terms of inhibition by microcystin, calyculin A and okadaic acid (Table IV). The two preparations did not require Mn2 þ for activity, and similar to native PP1C, addition of 1 mM Mn2 þ inhibited

[24]

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TABLE III PURIFICATION OF PP1C FROM Sf9 CELLS Purification step Crude lysate DEAE-cellulose Heparin-Sepharose Sephacryl S-200

REGULATION

OF

USING

METHOD 1

Volume (ml)

Protein (mg)

Total activity Units

Specific activity (units/mg)

Purification (-fold)

Yield (%)

145 155 111 34

544 85 9.4 0.7

44.9 22.7 14.3 10.8

0.08 0.27 1.5 15.2

1 3 19 190

100 51 32 24

TABLE IV VARIOUS RECOMBINANT PREPARATIONS OF PP1C

BY INHIBITORS

IC50 (nM) Source of PP1C P-DARPP-32 S-DARPP-32 Microcystin-LR Calyculin A Okadaic acid Sf9 PP1C Sf9 PP1CMn2 þ E. coli PP1C Rabbit PP1C

2.2 370 450 2.9

5.4 120 115 1.5

0.1 0.3 0.1 0.1

0.8 1.1 0.6 0.8

17 34 36 22

P-DARPP-32 and S-DARPP-32 are proteins phosphorylated by PKA using ATP or thiopho-ATP, respectively.

enzyme activities
50%. Notably, PP1C and did not dephosphorylate phospho-DARPP-32 or phospho-inhibitor-1. Consistent with this, both PP1C and were inhibited by either phospho- or thiophospho-DARPP-32 with IC50 values in the low nM range (Table IV and data not shown), properties similar to those of native PP1C. In addition, PP1C and were both relatively resistant to autodephosphorylation during phosphorylation by cdc2 kinase (Fig. 1 and data not shown). Given the anomalous properties of E. coli PP1C which is expressed in the presence of Mn2 þ , the effect of addition of Mn2 þ to the Sf9 culture medium was also investigated. Addition of 1 mM MnCl2 during the incubation period and purification of PP1C increased the total level of expression, but a greater proportion of protein was found in the particulate fraction (>70% after 36 hr incubation for either PP1C or ). The specific activity of Sf9 PP1CMn was significantly higher (
2-fold) than that of Sf9 PP1C. Like E. coli PP1C, Sf9 PP1CMn was normal in terms of inhibition by microcystin, calyculin A and okadaic acid (Table IV). However, Sf9 PP1CMn was relatively insensitive to either phosphoor thiophospho-DARPP-32 (Table IV). Furthermore, this was associated

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with the ability of Sf9 PP1CMn to rapidly dephosphorylate phosphoDARPP-32 or phospho-inhibitor-1 (Vmax of 58 mol/min/mg and Km of 28 M for phospho-inhibitor-1). Thus the addition of extracellular Mn2 þ to Sf9 cells results in the production of PP1C preparation that is very similar to E. coli PP1C. The ability of extracellular Mn2 þ to influence the properties of PP1C (in a cellular environment where the protein has the ability to fold into a native structure) presumably reflects the nonphysiological levels of this divalent cation. Interestingly, chronic Mn2 þ poisoning has been characterized as an irreversible syndrome that bears a striking similarity to Parkinson’s disease, although the detailed neuropathology appears to be distinct.25,26 Specifically, with chronic Mn2 þ , neurodegeneration occurs downstream of the nigrostriatal dopaminergic projection in the striatum and pallidum, rather than in dopaminergic neurons of the substantia nigra (as occurs in Parkinson’s disease). In the medium spiny neurons of the striatum, PP1C has been found to play a key role in mediating the postsynaptic effects of dopamine as a consequence of its regulation by DARPP-32.2 Thus it is conceivable that some of the effects of chronic Mn2 þ poisoning are mediated via the nonphysiological consequences of Mn2 þ on the activities of PP1C in medium spiny neurons.

Characterization of PP1 Expressed in Sf9 Cells Using Method 2

Wild-type, mutant and chimeric PP1C proteins have all been prepared in Sf9 cells using method 2.21 For these various PP1 preparations, expression reached a maximum between 48 and 72 hr postinfection, and approximately 50% of expressed PP1 was recovered in the soluble fraction. The inclusion of the His6 (or GST) tags made the purification more straightforward. Using the two-step affinity chromatography method (either purification on Ni-NTA or glutathione-Sepharose resins then chromatography on the PNUTS affinity column), up to 10 g of purified protein (usually > 90% pure) was generally recovered from 1  108 cells in 100 ml culture. Alternatively, partially purified recombinant PP1 can be rapidly obtained by the one-step procedure, and this is usually sufficient

25

D. B. Calne, N. S. Chu, C. C. Huang, C. S. Lu, and W. Olanow, Neurology 44, 1583–1586 (1994). 26 D. Centonze, P. Gubellini, G. Bernardi, and P. Calabresi, Exp. Neurol. 172, 469–476 (2001).

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337

for the analysis of the effects of various inhibitors or proteins on PP1 activity (
0.2 ng of PP1 protein is sufficient for a single assay reaction). In general, one-step chromatography using glutathione-Sepharose resulted in a purer preparation compared to that of one-step chromatography using Ni-NTA resin. As discussed above for Sf9 cell method 1, tagged Sf9 PP1C exhibited properties essentially identical to those of native PP1, with respect to its lack of dependence on added Mn2 þ , and sensitivity to inhibition by phospho-DARPP-32 and inhibitor-2. Sf9 PP1C also exhibited an ability to bind tightly to spinophilin and PNUTS, and to be inhibited by these two targeting subunits. Importantly, Sf9 PP1C also failed to dephosphorylate tyrosine-phosphorylated myelin basic protein or phosphoDARPP-32. Given the native-like properties of Sf9 PP1C, it has been possible to carry out detailed structure-function analysis using the wild-type enzyme as a control.21 Thus, studies of the potential role of residues in the surface grooves near the active site have revealed that several acidic amino acids do not appear to contribute to regulation of the enzyme by thiophosphoDARPP-32. Other mutation studies of Sf9 PP1C revealed an important role for Tyr272 in enzyme activity. In contrast, previous studies in which Tyr272 has been mutated in E. coli PP1C had not revealed any effect on enzyme activity. In addition, mutation of Tyr272 resulted in an increase in the relative phosphatase activity towards both tyrosine-phosphorylated myelin basic protein and phospho-DARPP-32, with a much larger effect being observed for PP1[Tyr272Ala] than for PP1[Tyr272Phe]. Tyr272 is found in a loop between 12 and 13 strands, and overhangs the active site of the phosphatase.15,16 Based on comparison of the properties of Sf9 and E. coli PP1C, it seems likely that the precise position of this residue may be responsible for some of the abnormal activities of E. coli PP1C.21 In summary, Sf9 PP1 is an excellent preparation for almost all types of biochemical studies. The yield is lower than for E. coli PP1C and is less easy to scale up. However, it is convenient to use affinity chromatography procedures to rapidly obtain PP1C mutants that are suitable for enzyme assays. Most importantly, these studies indicate that PP1C folds in Sf9 cells in the same way that it does in mammalian cells, and the biochemical properties of Sf9 PP1C are indistinguishable from native PP1C typically purified from rabbit skeletal muscle. However, addition of extracellular Mn2 þ to Sf9 cell cultures is sufficient to perturb this normal process, in which case an enzyme preparation is generated that is much like that formed in E. coli. Presumably, a high intracellular level of Mn2 þ is generated that is sufficient to compete with Fe2 þ and Zn2 þ for binding to PP1C as it folds.

338

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Possibly, this effect of Mn2 þ might contribute to neurodegeneration associated with chronic Mn2 þ poisoning. Acknowledgments This work was supported by U.S. Public Health Grant MH40899 (to A. C. N. and P. G.).

[24] An Inducible System to Study the Growth Arrest Properties of Protein Phosphatase 2C By PAULA OFEK, DANIELLA BEN-MEIR, and SARA LAVI Introduction

The PP2C family of phosphatases is one of four major groups of serine/threonine phosphatases (PP1, PP2A, PP2B, and PP2C) in eukaryotes, which is distinguished from the other groups by its dependence on magnesium ions and its insensitivity to the tumor promoter okadaic acid. The UniGene database (UniGene, NIH) indicates that the human genome contains at least six PP2C paralogs. Several independent reports suggest that different members of the PP2C family regulate transcription of growth-related pathways in mammals.1–5 This protein is highly conserved in evolution.6 Roles for PP2C in response to stress were identified in Arabidopsis7,8 as well as in yeast and mammalian cells.3,9,10 PP2C (also referred as PPM1A) is the most characterized member of the PP2C group. It is a monomeric enzyme of about 42 kDa that shows

1

M. A. Guthridge, P. Bellosta, N. Tavoloni, and C. Basilico, Mol. Cell Biol. 17, 5485 (1997). M. Fiscella, H. Zhang, S. Fan, K. Sakaguchi, S. Shen, W. E. Mercer, G. F. Vande Woude, P. M. O’Connor, and E. Appella, Proc. Natl. Acad. Sci. U.S.A. 94, 6048 (1997). 3 Y. Tong, R. Quirion, and S. H. Shen, J. Biol. Chem. 273, 35282–35290 (1998). 4 A. Cheng, P. Kaldis, and M. J. Solomon, J. Biol. Chem. 275, 34744 (2000). 5 C. Leung-Hagesteijn, A. Mahendra, I. Naruszewicz, and G. E. Hannigan, EMBO J. 20, 2160 (2001). 6 P. Cohen, D. L. Schelling, and M. J. Stark, FEBS Lett. 250, 601 (1989). 7 J. Sheen, Proc. Natl. Acad. Sci. U.S.A. 95, 975 (1998). 8 S. Tahtiharju and T. Palva, Plant J. 26, 461 (2001). 9 M. Hanada, J. Ninomiya-Tsuji, K. Komaki, M. Ohnishi, K. Katsura, R. Kanamaru, K. Matsumoto, and S. Tamura, J. Biol. Chem. 276, 5753 (2001). 10 M. Hanada, T. Kobayashi, M. Ohnishi, S. Ikeda, H. Wang, K. Katsura, Y. Yanagawa, A. Hiraga, R. Kanamaru, and S. Tamura, FEBS Lett. 437(3), 172 (1998). 2

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Possibly, this effect of Mn2 þ might contribute to neurodegeneration associated with chronic Mn2 þ poisoning. Acknowledgments This work was supported by U.S. Public Health Grant MH40899 (to A. C. N. and P. G.).

[25] An Inducible System to Study the Growth Arrest Properties of Protein Phosphatase 2C By PAULA OFEK, DANIELLA BEN-MEIR, and SARA LAVI Introduction

The PP2C family of phosphatases is one of four major groups of serine/threonine phosphatases (PP1, PP2A, PP2B, and PP2C) in eukaryotes, which is distinguished from the other groups by its dependence on magnesium ions and its insensitivity to the tumor promoter okadaic acid. The UniGene database (UniGene, NIH) indicates that the human genome contains at least six PP2C paralogs. Several independent reports suggest that different members of the PP2C family regulate transcription of growth-related pathways in mammals.1–5 This protein is highly conserved in evolution.6 Roles for PP2C in response to stress were identified in Arabidopsis7,8 as well as in yeast and mammalian cells.3,9,10 PP2C (also referred as PPM1A) is the most characterized member of the PP2C group. It is a monomeric enzyme of about 42 kDa that shows

1

M. A. Guthridge, P. Bellosta, N. Tavoloni, and C. Basilico, Mol. Cell Biol. 17, 5485 (1997). M. Fiscella, H. Zhang, S. Fan, K. Sakaguchi, S. Shen, W. E. Mercer, G. F. Vande Woude, P. M. O’Connor, and E. Appella, Proc. Natl. Acad. Sci. U.S.A. 94, 6048 (1997). 3 Y. Tong, R. Quirion, and S. H. Shen, J. Biol. Chem. 273, 35282–35290 (1998). 4 A. Cheng, P. Kaldis, and M. J. Solomon, J. Biol. Chem. 275, 34744 (2000). 5 C. Leung-Hagesteijn, A. Mahendra, I. Naruszewicz, and G. E. Hannigan, EMBO J. 20, 2160 (2001). 6 P. Cohen, D. L. Schelling, and M. J. Stark, FEBS Lett. 250, 601 (1989). 7 J. Sheen, Proc. Natl. Acad. Sci. U.S.A. 95, 975 (1998). 8 S. Tahtiharju and T. Palva, Plant J. 26, 461 (2001). 9 M. Hanada, J. Ninomiya-Tsuji, K. Komaki, M. Ohnishi, K. Katsura, R. Kanamaru, K. Matsumoto, and S. Tamura, J. Biol. Chem. 276, 5753 (2001). 10 M. Hanada, T. Kobayashi, M. Ohnishi, S. Ikeda, H. Wang, K. Katsura, Y. Yanagawa, A. Hiraga, R. Kanamaru, and S. Tamura, FEBS Lett. 437(3), 172 (1998). 2

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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broad substrate specificity. The catalytic domain, of 300 amino acid residues in the N terminus, consists of six -helices and 11 -sheets and is common to all enzymes that belong to the PP2C family. The C-terminal domain includes a sequence of 80 amino acids that forms three -helices. This domain determines the substrate specificity of the enzyme and displays no similarity to the other paralogs except for the closely related PP2C .11 A growing list of substrates has been suggested to be specifically dephosphorylated by PP2C in eukaryotic cells.12–20 Still, due to the absence of specific inhibitors and the presence of multiple paralogs the natural role of PP2C in mammalian cells is not known. In human embryo kidney (HEK) 293 cells over expression of PP2C and PP2C were highly toxic to the cells and we were not able to obtain stable cell lines overexpressing PP2C. Hereby, we present a system that enabled us to study PP2C expression in 293 cells. Methods Establishment of Inducible PP2C Cells

Plasmid Constructions PP2C-pcDNA3.1 (wt-PP2C). A rat cDNA library21 was screened with probes derived from the rat PP2C gene,22 yielding several PP2C clones. A clone encoding the complete PP2C cDNA was isolated and converted to a plasmid, enabling CMV promoter-driven transcription of the inserted cDNA. The complete coding sequence of PP2C was amplified by PCR and cloned into expression vector pcDNA3.1 (Invitrogen), between the HindIII and ApaI sites. Sequence accuracy was verified by DNA sequencing after cloning. 11

A. K. Das, N. R. Helps, P. T. Cohen, and D. Barford, EMBO J. 15, 6798 (1996). T. S. Ingebritsen and P. Cohen, Eur. J. Biochem. 132, 255 (1983). 13 S. P. Davies, N. R. Helps, P. T. Cohen, and D. G. Hardie, FEBS Lett. 377, 421 (1995). 14 K. Fukunaga, L. Stoppini, E. Miyamoto, and D. Muller, J. Biol. Chem. 268, 7863 (1993). 15 M. Takekawa, T. Maeda, and H. Saito, EMBO J. 17, 4744 (1998). 16 A. A. Welihinda, W. Tirasophon, S. R. Green, and R. J. Kaufman, Mol. Cell Biol. 18, 1967 (1998). 17 T. Zhu, D. Dahan, A. Evagelidis, S. Zheng, J. Luo, and J. W. Hanrahan, J. Biol. Chem. 274, 29102 (1999). 18 A. Hishiya, M. Ohnishi, S. Tamura, and F. Nakamura, J. Biol. Chem. 274, 26705 (1999). 19 A. Cheng, K. E. Ross, P. Kaldis, and M. J. Solomon, Genes. Dev. 13, 2946 (1999). 20 E. T. Strovel, D. Wu, and D. J. Sussman, J. Biol. Chem. 275, 2399 (2000). 21 A. Zauberman, A. Lupo, and M. Oren, Oncogene 10(12), 2361 (1995). 22 E. Seroussi, N. Shani, D. Ben-Meir, A. Chajut, I. Divinski, S. Faier, S. Gery, S. Karby, Z. Kariv-Inbal, O. Sella, N. I. Smorodinsky, and S. Lavi, J. Mol. Biol. 312(3), 439 (2001). 12

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PP2CD239A (mut-PP2C). PCR was performed to introduce the mutation D239A into PP2C. Two PCR reactions were conducted in parallel, using PP2C-pcDNA3.1 as the template. In the first, the upstream primer introducing the mutation in the sense direction was 50 CTTGCATGTGCTGGCATCTGG; the downstream primer was SP650 ATTTAGGTGACACTATAG. In the second reaction, the upstream primer contained the internal PP2C sequence 50 ACACGGTGCAGATAGAAGTG and the downstream primer contained the mutation in the antisense direction 50 CCAGATGCCAGCACATGCAAG. Following purification, the resultant PCR products from the two parallel reactions were mixed and used as a template in a final PCR reaction mix containing the two external primers (the upstream primer with the internal PP2C sequence and the downstream with the SP6 primer). The final PCR product was cloned into PP2C-pcDNA3.1 via the EcoRI and ApaI sites. The sequence accuracy of the mut-PP2C was verified by sequencing. Inducible PP2C (PP2C-pcDNA4/TO): PP2C (wt or mut) was isolated from the pcDNA3 based vectors described above, via the HindIII and ApaI sites, and subcloned into the pcDNA4/TO vector (T-Rex system, Invitrogen). The T-RexTM System The T-RexTM system is a tetracycline-regulated mammalian expression system that uses regulatory elements from the E. Coli Tn10-encoded tetracycline (Tet) resistance operon.23 Tetracycline regulation in the T-RexTM system is based on the binding of tetracycline to the Tet repressor and derepression of the promoter controlling expression of the gene of interest.24 T-RexTM-293 cells, stably expressing the regulatory plasmid pcDNA6/ TR (Invitrogen), were transfected by calcium phosphate/ DNA precipitation with PP2C-pcDNA4/TO wt or mut, or an empty vector (pcDNA4/TO). 48 hr after transfection cells were seeded into Dulbecco’s modified Eagle’s medium containing blasticidin (5 g/ml) and zeocin (200 g/ml) (both purchased from Invitrogen, Carlsbad, CA). Several clones were isolated and characterized. Plating Efficiency

Cells were seeded (500 or 2000 cells/well in a 24-well or a 6-well plate respectively), and grown for 10 days. The resistant colonies were fixed with 4% formaldehyde in PBS, stained with Giemsa stain (Sigma) and counted. 23 24

W. Hillen and C. Berens, Annu. Rev. Microbiol. 48, 345 (1994). F. Yao, T. Svensjo, T. Winkler, M. Lu, C. Eriksson, and E. Eriksson, Hum. Gene Ther. 9(13), 1939 (1998).

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Antibodies

Monoclonal Anti-PP2C Antibodies (9F4) Coding sequences of PP2C were cloned into the pET-28b bacterial expression vector (Novagen) and the resulting plasmid was used to transform BL21 (DE3) E. coli. Culture of the transformants grown overnight at 30
C, following induction by 0.1 mM isopropyl D-thiogalactoside (IPTG), led to overexpression of soluble and active PP2C. The recombinant protein was purified on a nickel-agarose column (Qiagen) under nondenaturing conditions. BALB/c mice were immunized subcutaneously with 50 g of the recombinant PP2C emulsified in complete Freund’s adjuvant, followed by five subcutaneous boosts (50 g PP2C in incomplete Freund’s adjuvant), at 2–3 weeks intervals. Three days before fusion, the mice received an i.v. boost of 50 g PP2C in PBS. One mouse, which acquired a relatively high titer of anti-PP2C antibodies, was chosen for fusion. Hybridomas were prepared by fusion of spleen cells with mouse myeloma NSO (kindly provided by C. Milstein, Imperial College, Cambridge, UK). The hybridomas were screened by ELISA, immunofluorescence and immunoblotting of the supernatants. Anti-PP2C specific monoclonal antibodies (9F4) were chosen after two rounds of screening. Immunoglobulin heavychain isotyping was carried out with an IsoStrip Mouse Monoclonal Antibody Isotyping Kit (Roche, Germany) and was found to be IgG1. Cell Extracts and Western Blot Analysis

Cells were harvested with PBS and lysed in 50 mM Hepes, 150 mM NaCl, 1% Triton, pH 8.0 supplemented with protease inhibitors (Boehringer Mannheim). Cell debris was pelleted and protein concentrations were measured by the BCA reagent (Pierce Inc., Rockford, IL). Proteins were separated by SDS-PAGE.25 After electrophoresis, the proteins on the gel were transferred to a nitrocellulose membrane. The membrane was blocked and immunoblotted with anti-PP2C antibodies, followed by peroxidaseconjugated IgG (Jackson, West Grove, PA) and West Pico Chemiluminescent Substrate (Pierce Inc.). XTT Assay

This test is based on the ability of metabolic active cells to reduce the tetrazolium salt XTT26 to orange colored compounds of formazan. 25

F. M. Ausubel, R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, and J. A. Smith, ‘‘Current Protocols in Molecular Biology’’. John Wiley and Sons, Inc., New York, 1996. 26 D. A. Scudiero, R. H. Shoemaker, K. D. Paull, A. Monks, S. Tierney, T. H. Nofziger, M. J. Currens, D. Seniff, and M. R. Boyd, Cancer Res. 48(17), 4827 (1988).

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The greater the number of live cells in the well, the greater the activity of mitochondria enzymes, and the higher the concentration of the dye formed, which can then be measured and quantitated. T-Rex-293 cells expressing wt-PP2C, mut-PP2C, or empty vector TO (2  104 cells/ well in a 96-well plate) were incubated for 24 and 48 hr in the presence or absence of Tet (1 g/ml). 50 l of XTT reaction solution (Biological Industries, Israel) was added to each well and the plate was incubated at 37
C for 2 hr. The sample absorbance was measured with an ELISA reader at a wavelength of 450 nm. The reference absorbance (nonspecific readings) was measured at a wavelength of 630 nm. Protein Phosphatase Activity: Malachite-green Assay

This assay is specific for the PP2C family and distinguishes it from the other classes of protein phosphatases (PP1, PP2A, and PP2B). It is performed in the presence of okadaic acid that completely inhibits PP1 and PP2A, EGTA that neutralizes the Ca2 þ /calmodulin-dependent PP2B, and Mg2 þ that activates PP2C. Thus, PP2C activity is measured as the Mg2 þ dependent and okadaic acid-insensitive activity.27 The phosphopeptide substrate in our assay, FLRTpSCG is derived from AMP-activated protein kinase (AMPK) and was previously shown to be a good substrate for PP2C.28 Protein extracts were prepared from transfected cells and free phosphate was removed with a VivaSpin concentrator (cut-off 10,000 Da). Phosphatase activity was then measured colorimetrically as described.29 Briefly, the assay was performed in 30 l of assay buffer [50 mM Tris (pH 7.5) 0.1 mM EGTA] containing 5 g of cell extract and 0.5 mM of the substrate FLRTpSCG,28 in the presence of 30 mM MgCl2, 5 M okadaic acid and 5 g of bovine serum albumin. After an incubation of 30 min at 30
C, the reaction was terminated by adding 70 l of cold assay buffer, followed by 25 l of malachite green/ammonium molybdate reagent. Measurements were taken at 630 nm in an ELISA reader (Dynatech MR5000). Results Establishment of Tet-inducible PP2C-Expressing Cells

Earlier studies in our laboratory have demonstrated that overexpression of PP2C leads to cell cycle arrest in the G2/M phase or to induction of 27

P. Cohen and P. T. Cohen, J. Biol. Chem. 264, 21435 (1989). A. E. Marley, J. E. Sullivan, D. Carling, W. M. Abbott, G. J. Smith, I. W. Taylor, F. Carey, and R. K. Beri, Biochem. J. 320(Pt 3), 801 (1996). 29 A. A. Baykov, O. A. Evtushenko, and S. M. Avaeva, Anal. Biochem. 171(2), 266 (1988). 28

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FIG. 1. 293 cells were transfected with 5 g of the following expression vectors: pcDNA3 empty vector (cont.), wt-PP2C (wt), or mut-PP2C (mut). Cells were grown in medium containing G418 (1.2 mg/ml) for two weeks until resistant colonies appeared. The colonies were then stained with Giemsa dye.

apoptosis. Thus, stable cell lines expressing PP2C could not be established (Fig. 1). To facilitate the analysis of PP2C in human cells we constructed stable clones expressing PP2C under the regulation of a tetracycline-inducible promoter (T-RexTM). Expression of PP2C under the control of a tetracycline repressor permitted the isolation of stable clones in which PP2C expression and its consequent killing effect could be tightly modulated by the addition of tetracycline (Tet). Clones capable of expressing high levels of wild type PP2C (wt-PP2C) could be easily isolated as long as they were grown in the absence of tetracycline. In parallel, cultures expressing inducible mutated PP2C (mutPP2C), or control cultures harboring the empty vector (pcDNA4-TO) were also constructed. T-Rex-293 PP2C (wt or mut)/TO cells extracts were immunoblotted with anti-PP2C antibodies. In the absence of Tet, all the clones expressed only the basal level of PP2C, indicating that there is a tight suppression of PP2C expression and that the clones are not leaky. Clones expressing the highest levels of PP2C in the presence of Tet were selected for further experiments. A pool of mut-PP2C transfected cells selected for further experiments also expressed high levels of PP2C in the presence of Tet. Only the basal level of PP2C was detected in clones expressing the empty vector (pcDNA4-TO) when grown either in the presence or absence of tetracycline (data not shown).

Characterization of T-Rex-293 PP2C Cells

PP2C Expression Level is Regulated by Tetracycline Concentration and Time Dependent T-Rex-293 wt-PP2C cells were treated for 24 hr with different concentrations of tetracycline (0, 6, 12, 25, 50, 100 ng/ml). Cells were lysed

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FIG. 2. T-Rex-293 wt-PP2C cells were treated for 24 hr with different concentrations of tetracycline (0, 6, 12, 25, 50, 100 ng/ml). (A) Cells were lysed and extracts were assayed for PP2C expression level with anti-PP2C antibodies (9F4). (B) Phosphatase activity was determined by the Malachite Green assay. (C) T-Rex-293 wt-PP2C cells were grown in several concentrations of Tet (0, 3, 12, 50, or 200 ng/ml) for 17 days, after that resistant colonies were stained with Giemsa dye.

and extracts were assayed for PP2C expression with anti-PP2C antibodies. As can be seen in Fig. 2A, cells grown without Tet express only basal PP2C levels. PP2C overexpression is proportional to the Tet concentration. In a parallel experiment, cells expressing wt or mut-PP2C were incubated with Tet (1 g/ml) for 0, 24, or 48 hr. PP2C expression level was monitored in the cells extracts by Western blot, and was increased by the lapsing time (Fig. 3A).

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FIG. 3. T-Rex-293 cells expressing PP2C (wt or mut) were incubated for 24 and 48 hr with Tet (1 g/ml). In parallel, control cultures were incubated without Tet. (A) The cells were lysed, electrophoresed and immunoblotted with anti-PP2C antibodies. (B) The cells were then assayed for viability using the XTT assay. The percent of viable Tet-treated cells relative to the control (non-treated cells) was calculated and the average of four different wells from a representative experiment was plotted þ / std.dev. wt and mut-PP2C expressing cells are represented by black and white squares, respectively.

PP2C Phosphatase Activity is Regulated by Tetracycline Concentration T-Rex-293 wt-PP2C cells were treated for 24 hr with different concentrations of Tetracycline. Cells were lysed and assayed for phosphatase activity using the malachite green phosphatase assay (Fig. 2B). The PP2C specific activity increased proportionally to the Tet concentration. These experiments demonstrate that the Tet-induced PP2C is enzymatically active. PP2C Induction and Its Effects

Cell Proliferation and Colony Formation are Inhibited by PP2C Overexpression T-Rex-293 PP2C (wt or mut) cells were incubated with Tet (1 g/ml) for 0, 24, or 48 hr and assayed for PP2C induction (Fig. 3A) and proliferation rate. As shown in Fig. 3B, less than 40% of the wt-PP2C expressing cells remained viable after 48 hr incubation with Tet. Tetracycline-induced growth inhibition and wt-PP2C expression were time dependent and tightly correlated. Induction of mut-PP2C did not

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inhibit the cells proliferation rate (Fig. 3). T-Rex-293 cells expressing the control empty vector (TO) were not affected by Tet (data not shown). T-Rex-293 PP2C cells were seeded in different Tet concentrations and grown to form colonies. The cells’ plating efficiency decreased with increasing levels of Tet (Fig. 2C), in direct correlation with the enhanced expression and activity of PP2C (Fig. 2A and B). PP2C Overexpression Causes Cell Cycle Arrest and Apoptosis T-Rex-293 wt-PP2C (wt/mut) cells were incubated for 0, 24, 48, or 72 hr with tetracycline. The cell cycle distribution was assayed by fluorescence-activated cell sorter (FACS). As shown in Fig. 4, wt-PP2C expression dramatically affected the cell cycle in the Tet induced cells. 24 hr after wt-PP2C induction there was a

FIG. 4. T-Rex-293 wt-PP2C (wt/mut) cells were incubated for 0, 24, 48, or 72 hr with tetracycline 1 g/ml, and analyzed by flow cytometry. (A) The percent of cells in every cell cycle step was monitored for each treatment. (B) Protein extracts (10 g) from each sample were electrophoresed and immunoblotted with anti-PP2C monoclonal antibodies.

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very pronounced arrest of the cells in the G2/M phase. At 48 hr, in addition to the G2/M arrest, there was a pronounced apoptosis as the percentage of sub G1 cells increased. At 72 hr most of the cells underwent apoptosis. In contrast, overexpression of the mutant protein did not alter the cell cycle. Further experiments performed on PP2C using this system are reported in more detail in P. Ofek et al.30

Acknowledgments We wish to thank N. Kazhdan for her assistance and S. Karby and N. Smorodinsky for their help in the preparation of the antibodies. We are also grateful to O. Sagi-Assif and B. Slobodin for their help with the FACS analysis.

30

P. Ofek, D. Ben-Meir, Z. Kariv-Inbal, M. Oren, and S. Lavi, J. Biol. Chem. 278, 14299 (2003).

[25] Use of Tetracycline-Regulatable Promoters for Functional Analysis of Protein Phosphatases in Yeast By JOAQUI´N ARIN˜O and ENRIC HERRERO

Introduction

Ser/Thr protein phosphatases regulates a large variety of important biological processes, often necessary for cell survival. Given the high level of evolutionary sequence conservation for many of these enzymes, yeast cells (particularly Saccharomyces cerevisiae) represent a very useful bench work for phosphatase research, primarily because of the power of the molecular biology and genetics of this organism. However, the essential roles of these enzymes prevent in many cases straightforward approaches to analyze gene function, such as gene disruptions (resulting in lethality in the case of single genes, such as GLC7, encoding the catalytic subunit of type 1 protein phosphatase) or high-copy overexpression (yielding very poor growth and other defects). Often, these problems can be circumvented by the use of regulatable expression systems. In this chapter we introduce the reader to the tetracycline-regulatable promoters in yeast and we describe its use for functional analysis using the Sit4 and Ppz1 Ser/Thr protein phosphatases as example.

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very pronounced arrest of the cells in the G2/M phase. At 48 hr, in addition to the G2/M arrest, there was a pronounced apoptosis as the percentage of sub G1 cells increased. At 72 hr most of the cells underwent apoptosis. In contrast, overexpression of the mutant protein did not alter the cell cycle. Further experiments performed on PP2C using this system are reported in more detail in P. Ofek et al.30

Acknowledgments We wish to thank N. Kazhdan for her assistance and S. Karby and N. Smorodinsky for their help in the preparation of the antibodies. We are also grateful to O. Sagi-Assif and B. Slobodin for their help with the FACS analysis.

30

P. Ofek, D. Ben-Meir, Z. Kariv-Inbal, M. Oren, and S. Lavi, J. Biol. Chem. 278, 14299 (2003).

[26] Use of Tetracycline-Regulatable Promoters for Functional Analysis of Protein Phosphatases in Yeast By JOAQUI´N ARIN˜O and ENRIC HERRERO

Introduction

Ser/Thr protein phosphatases regulates a large variety of important biological processes, often necessary for cell survival. Given the high level of evolutionary sequence conservation for many of these enzymes, yeast cells (particularly Saccharomyces cerevisiae) represent a very useful bench work for phosphatase research, primarily because of the power of the molecular biology and genetics of this organism. However, the essential roles of these enzymes prevent in many cases straightforward approaches to analyze gene function, such as gene disruptions (resulting in lethality in the case of single genes, such as GLC7, encoding the catalytic subunit of type 1 protein phosphatase) or high-copy overexpression (yielding very poor growth and other defects). Often, these problems can be circumvented by the use of regulatable expression systems. In this chapter we introduce the reader to the tetracycline-regulatable promoters in yeast and we describe its use for functional analysis using the Sit4 and Ppz1 Ser/Thr protein phosphatases as example.

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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Regulation of Gene Expression in Yeast Cells Using the Tet System

The Tet system is being used for the regulation of gene expression in cells of different organisms (such as yeasts, plants, Drosophila, or mammals), under the control of doxycycline and other antibiotic molecules of the tetracycline family.1 In its initial form, it was designed for regulated expression in mammalian cells.2 It is based on the activity of a hybrid transcriptional regulator (tTA) formed by the TetR DNA-binding domain (the tetracycline-inducible repressor from the Escherichia coli Tn10 transposon) fused to the VP16 activator domain of the herpes simplex virus. The tTA molecules are able to interact with high affinity with tetO operator sequences, this interaction being prevented by doxycycline or other tetracyclines. In this original direct system, tTA acts as an activator of the expression of genes under the control of an array of tetO sequences, and rapid decay of expression is attained upon doxycycline addition, allowing tight control of gene transcription. Later, a reverse system was developed,3 based on a mutated TetR moiety (rTetR) that allows binding of the modified regulator (rtTA) to tetO sequences only in the presence of the antibiotic. This system displays high levels of gene expression upon doxycycline addition. More recently, new rtTA mutants have been selected after random and directed mutagenesis that show higher induction levels by doxycycline, higher stability and lower background expression in mammalian cell lines.4 These properties make them a useful tool for the in vivo control of gene expression in transgenic organisms.5 The Direct Tet System in Yeast Cells

Regulated gene expression is an essential tool for functional analysis in yeast cells, as well as for biotechnological applications. In Saccharomyces cerevisiae, vectors based on promoters such as those of GAL1, MET3, PHO5, or CUP1 have been traditionally used for those purposes.6 However, controlling gene expression from these promoters involves changes in culture conditions that may cause undesired pleiotropic effects on cell 1

U. Baron and H. Bujard, Methods Enzymol. 327, 401 (2000). M. Gossen, A. L. Bonnin, and H. Bujard, Trends Biochem. Sci. 18, 471 (1993). 3 M. Gossen, S. Freundlieb, G. Bender, G. Muller, W. Hillen, and H. Bujard, Science 268, 1766 (1995). 4 S. Urlinger, U. Baron, M. Thellmann, M. T. Hasan, H. Bujard, and W. Hillen, Proc. Natl. Acad. Sci. USA 97, 7963 (2000). 5 E. Vigna, S. Cavalieri, L. Ailles, M. Geuna, R. Loew, H. Bujard, and L. Naldini, Mol. Ther. 5, 252 (2002). 6 M. J. Stark, Yeast gene analysis, in ‘‘Methods in Microbiology’’ (A. J. P. Brown and M. Tuite, eds.), Vol. 26, p. 83. Academic Press, San Diego, 1998. 2

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metabolism, therefore obscuring the phenotypic effects of switching-on or off the studied gene. For instance, induction of gene expression from the GAL1 promoter (the more commonly used of them) requires shifting yeast cells from glucose or raffinose as carbon sources to galactose-based growth medium. On the contrary, the employment of molecules of the tetracycline family for regulated gene expression in S. cerevisiae and other yeasts offer an alternative to the above systems, as the antibiotic molecules (i) are not metabolized by the yeast cells, (ii) are not growth-inhibitory (at least up to 50 g/ml concentration in the culture medium in the case of doxycycline), (iii) do not cause other apparent side effects during growth, and (iv) regulation can occur in rich medium such as YPD for S. cerevisiae, without requiring culture media of specific composition. For this purpose, we adapted the Tet system from mammalian cells to S. cerevisiae.7 The initial plasmid vector constructions (Fig. 1A) contained an expression cassette with the ADH1 terminator (to avoid read-through expression from adjacent vector sequences), the tetO operator region followed by a multiple cloning site, and a CYC1 terminator to make termination of tetO-driven transcription independent of other genome sequences. Importantly, the vector with the promoter cassette also contains the direct tTA transactivator gene (constitutively expressed from a heterologous cytomegalovirus promoter), making unnecessary use of specific yeast strains with a chromosome-integrated activator. Based on this, a range of vectors have been constructed that differ on: (i) copy number (centromeric, for low-copy number or episomal, for high-copy number), (ii) selectable marker (URA3 or TRP1), (iii) the number of tetO unit sequences in tandem (two or seven), and (iv) the presence of a spacer region between the TetR and VP16 moieties of tTA to increase the flexibility of the molecule and its transcriptional activity. Under optimal conditions (multicopy plasmid, a tetO7 operator, and a modified tTA with one or two spacer region), about 60% expression levels are obtained relative to the GAL1 promoter under otherwise comparable conditions (using E. coli lacZ as reporter gene). Use of the above Tet plasmids to study the terminal phenotypes of null mutants in essential genes would require the construction of strains (by conventional yeast genetic methods) carrying an inactivating mutation in the chromosomal gene and transformed with a Tet plasmid conditionally expressing the gene. As an alternative, we have developed a one-step method8 that substitutes the endogenous gene promoter for a tetO2 or tetO7

7 8

E. Garı´ , L. Piedrafita, M. Aldea, and E. Herrero, Yeast 13, 837 (1997). G. Bellı´ , E. Garı´ , M. Aldea, and E. Herrero, Yeast 14, 1127 (1998).

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FIG. 1. (A) Structure of the EcoRI-HindIII fragment present in the Tet plasmids, based on the tetO-CYC1 TATA box hybrid promoter (Ref. 7). This promoter can contain two or seven tetO boxes in tandem. The direct tTA regulator (TetR-VP16 activator domain) is expressed under the control of the human cytomegalovirus promoter 1E. ADH1t and CYC1t correspond respectively to the downstream terminator regions of the S. cerevisiae genes ADH1 and CYC1. The plasmid multiple cloning site (MCS) is indicated with the restriction sites that are unique. (B) Scheme of the promoter substitution cassette based on the tetO-CYC1 TATA box promoter and kanMX4 as marker. The flanking regions with the restriction sites in tandem are shown. For more details, see Ref. 8.

promoter carried in a cassette (Fig. 1b) that also contains the tTA transactivator and kanMX4 (coding for geneticin resistance) as reporter gene for transformants. The method and the kanMX4-based cassette is an extension of the previously reported one-step strategy for construction of null mutants.9 It relies on the fact that PCR-amplified DNA fragments containing flanking sequences displaying about 40 bp homology with chromosomal regions are capable of chromosomal substitutive integration by homologous integration. The use of this one-step method to evaluate the functional interaction between the Sit4 and Ppz1/Hal3 phosphatase systems will be described in detail in the second half of this chapter. Other authors have separately developed a Tet system for regulated gene expression in S. cerevisiae using different constructions from the above ones, although displaying similar efficiency.10 In this case, two separate chromosomally integrative constructions are employed to transform yeast 9

A. Wach, A. Brachat, R. Po¨hlmann, and P. Philippsen, Yeast 10, 1793 (1994). S. Nagahashi, H. Nakayama, K. Kamada, H. Yang, M. Arisawa, and K. Kitada, Mol. Gen. Genet. 255, 372 (1997).

10

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cells: one for substitution of the endogenous gene promoter by a hybrid regulatable tetO-HOP1 promoter, and another for integration of a construction expressing one of the two alternative chimerical transactivators formed by the TetR DNA binding domain and the activator domains of the yeast proteins Gal4 or Hap4. This system has been adapted with slight modifications for the pathogenic yeast Candida glabrata.11 It is remarkable that the heterologous VP16 activating domain of tTA chimerical transactivator employed in our constructions7,8 is as much efficient as the activating domains of the yeast proteins Gal4 or Hap4. Construction of a Reverse Tet System for Yeast Cells

The original reverse Tet system for mammalian cells (doxycyclineinducible expression from tetO) based on mutant forms of TetR has also been adapted for S. cerevisiae.13 In our system, the reverse rtTA transactivator is also constitutively expressed from a cytomegalovirus promoter. Using growth conditions and antibiotic concentrations similar to those described for the direct system, we have observed detectable (by Northern analysis) induction of expression of a number of yeast genes after a short time (10–20 min) of doxycycline addition. Maximum levels are attained after 6 to 12 hr depending on the overexpressed gene. As expected, the kinetics of expression induction using the reverse system is more rapid than with the direct system, since in the former case it is not necessary to dilute the internal pool of doxycycline molecules to attain sufficient level of functional transactivator molecules. Even higher levels of overexpression can be reached with an autocatalytical system in which rtTA induces its own transcription under the control of a tetO-based promoter,13 besides activating expression of the desired gene. However, in this system the behavior of the individual yeast cells in the population is random, with a proportion of nonexpressing cells that depends on doxycycline concentration. The Dual Activator–Repressor System Allows Tight Control of Gene Expression

Many applications of regulated expression systems require a tight control of expression. Not only the attainment of high overexpression levels 11

H. Nakayama, M. Izuta, S. Nagahashi, E. Y. Sihta, Y. Sato, T. Yamazaki, M. Arisawa, and K. Kitada, Microbiology 144, 2407 (1998). 12 G. Bellı´ , E. Garı´ , L. Piedrafita, M. Aldea, and E. Herrero, Nucleic Acids Res. 26, 942 (1998). 13 A. Becskei, B. Se´raphin, and L. Se´rrano, EMBO J. 20, 2528 (2001).

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FIG. 2. Scheme of the dual (activator/repressor) Tet system in S. cerevisiae. The direct (doxycycline-repressible) and reverse (doxycycline-activable) versions are shown. The repressor moiety corresponds to the entire Ssn6 molecule or, alternatively, to a truncated Tup1 peptide lacking the N-terminal 72 amino acids of the native protein. See Ref. 12 for more details.

is experimentally important, but also that the basal expression levels were as low as possible. The latter is required specially when small amounts of the product can be functional, therefore masking the mutant phenotype. Although the original direct and reverse Tet systems allow high levels of expression in induced conditions, we have modified it by constructing a dual activator–repressor system12 in which the basal expression of different genes in repressing conditions is practically undetectable. In this system (Fig. 2) the tTA or rtTA regulators coexist in the same cell respectively with a rTetR-repressor or TetR-repressor chimerical protein, where the repressor moiety corresponds to the S. cerevisiae Ssn6 or Tup1 proteins. In our more frequently employed version of the dual system, the rTetR-Ssn6 or TetRSsn6 repressors are expressed constitutively from the cytomegalovirus promoter, and these constructions are chromosomally integrated at the LEU2 locus. The levels thus reached for the repressor are sufficient to compete with the activator at the appropriate doxycycline conditions. Experimental conditions are equivalent to those employed for the original direct or reverse systems.

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FIG. 3. Scheme of plasmids based on the direct Tet system for N- or C-terminal tagging with the 3xHA epitope and a 6xHis peptide. The plasmids are derived from the respective vectors described in Ref. 7. Only the expression cassette region is shown (not in scale). All the plasmids also contain URA3 as selective marker and the tTA activator. In order to make these constructions, a double-strand DNA fragment (with adequate extruding ends) coding for the 3xHA and 6xHis tags and also containing inframe ATG or stop codons was obtained from partially complementary synthetic oligonucleotides. It was then cloned in the multiple cloning site of the expression cassette shown in Fig. 1A.

Other Constructions Based on the Tet System

We have additionally constructed a number of Tet plasmids (Fig. 3) that besides the previously described elements also contain the coding sequence for: (i) three herpes simplex virus hemagglutinin-based epitopes in tandem (3xHA) for N- or C-terminal tagging of proteins overexpressed from the tetO7 -CYC1 promoter, and (ii) six histidine residues. The expression cassettes are inserted in vectors of the YEplac, YCplac, and Ylplac series,14 with URA3 as selection marker, and contain the direct tTA activator. They are useful for determination of the levels of the overproduced protein using

14

R. D. Gietz and A. Sugino, Gene 74, 527 (1988).

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standard Western blotting techniques with anti-HA antibody,12 and for purification of the protein using nickel columns. Construction and Phenotypic Analysis of a sit4 hal3 Conditional Mutant by Using a One-step Promoter Substitution Cassette and the Tet-regulatable Dual Expression System Background

SIT4 encodes a S. cerevisiae type 2A related Ser/Thr phosphatase that acts as a positive regulator of the G1/S transition of the yeast cell cycle, by affecting cyclin expression and budding. Consequently, sit4
mutants are either unviable (in ssd1-d backgrounds) or exhibit a slow growth phenotype.15 HAL3/SIS2 encodes a multicopy suppressor of the sit4 slow growth defect and this effect is mediated through the type 1 related Ser/Thr phosphatase Ppz1, which is inhibited by Hal3.16,17 Therefore, Hal3 can be considered a regulatory subunit of Ppz1. While a hal3 mutant grows normally under standard conditions, a sit4 hal3 mutant displays synthetic lethality, presumably due to a block at the G1/S transition.16,18 To investigate this situation, a conditional sit4 hal3 mutant strain was constructed by replacing the HAL3 promoter by a Tet regulatable promoter derived from plasmid pCM224,8 which contains two copies of the tetO operator sequences. Substitution of the HAL3 Promoter by a Doxycycline-Regulatable Promoter

Amplification of the Substitution Cassette 1.

The substitution cassette (approx. 3.9 kbp) was amplified from plasmid pCM224 by PCR using Expand High fidelity DNA polymerase (Roche) and standard methods. The oligonucleotides used were:

HAL3Prom-50 HAL3Prom-30

15

CCCTGGGTTAATGCCATGATCGTCTTTGATG TATCTCTAATGCAGCTGAAGCTTCGTACGC GTCAGCATCTTGCTTCCCGCTAGTAGAGGC GACGGCAGTCATATAGGCCACTAGTGGATC TG

A. Sutton, D. Immanuel, and K. T. Arndt, Mol. Cell. Biol. 11, 2133 (1991). J. Clotet, E. Garı´ , M. Aldea, and J. Arin˜o, Mol. Cell. Biol. 19, 2408 (1999). 17 E. de Nadal, J. Clotet, F. Posas, R. Serrano, N. Go´mez, and J. Arin˜o, Proc. Natl. Acad. Sci. USA. 95, 7357 (1998). 18 C. J. Di Como, R. Bose, and K. T. Arndt, Genetics 139, 95 (1995). 16

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The sequence in italics correspond to nt 578117–578158 (HAL3Prom-50 ) and 577718–577759 (HAL3Prom-30 ), of chromosome XI that correspond to the region
300/
260 and þ 1/ þ 42, respectively with respect to the initiating HAL3 Met codon. Underlined sequences represents fragments of the pCM224 plasmid multicloning site flanking the substitution cassette. 2.

The PCR reaction was run on a 0.7% agarose gel and the band excised from the gel and purified by available commercial systems (Geneclean II, Bio 101 Inc.). Although more time-consuming, we prefer this method to the alternative standard sodium acetate/ ethanol precipitation, as the latter do not efficiently remove long oligonucleotides.

Construction of the sit4
tetO:HAL3 Strain

Transformation of a sit4
Strain Usually, 1–5 g of linear amplified DNA are used for each transformation. We routinely prepare S. cerevisiae competent cells according to the method described by Gietz and Woods,19 with minor modifications. This method yields satisfactory and consistent results. It is recommended to maintain the starting culture in rich medium at relatively low density (not higher than 1.5  107 cells/ml). The addition of dimethyl sulfoxide to 10% (v/v) final concentration prior to the heat shock step has been found to increase the efficiency of transformation significantly. After the heat shock treatment, centrifuge the samples for 1 min at 5000 rpm in a microfuge, remove the supernatant and resuspend the cells in 1 ml of YPD (1% yeast extract, 2% peptone, 2% glucose) medium. Selection of Transformants As the substitution cassette contains the KanMX4 marker, selection is made on the basis of G418 (geneticin) resistance. 1. 2. 3.

19

Transfer the suspension into culture tubes containing 3 ml of YPD medium and incubate with shaking for 3 hr. Take a 1.5 ml aliquot into an Eppendorf tube, centrifuge at 5000 rpm for 1 min and remove approximately 1.1 ml of the supernatant. Resuspend the cells in the remaining volume of medium and disperse the suspension on YPD plates containing 200 mg/liter of G418 Sulfate (geneticin, Calbiochem).

R. D. Gietz and R. A. Woods, Yeast gene analysis, in ‘‘Methods in Microbiology’’ (A. J. P. Brown and M. Tuite, eds.), Vol. 26, p. 53. Academic Press, San Diego, 1998.

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5.

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Incubate plates at 30 C for 2–3 days. After this period, positive clones should appear as relatively big (about 3 mm diameter) colonies. Do not expect a large number of positives. The appearance of a relatively large number of small colonies (< 1 mm in diameter) probably is quite common, due to nonintegrative transformation events. Always use freshly prepared (no more than 2–3 days old) YPD-G418 plates at this step, as this greatly reduces the appearance of small colonies. Pick each big colony from the transformation plate and streak out the cells on YPD-G418 plates to isolate single cells. Only colonies that can grow from single cells can be considered as a possible positive for integration. This second round of selection do not require freshly prepared YPD-G418 (plates from the batch prepared for the first round of selection can be used here).

Verification of Correct Integration We verify on routine basis the correct integration checking the formation of both novel ends by PCR analysis using a pair of oligonucleotides external to the region of insertion and two more oligonucleotides internal to the kanamycin marker. The basis for this strategy and a detailed protocol for PCR amplification directly from yeast colonies can be found in Ref. 20.

Functional Analysis of a sit4 hal3 Conditional Mutant

As indicated in the ‘‘Introduction’’ section, this kind of regulatable promoters can serve to analyze the terminal phenotype of otherwise unviable mutants. In our case, positive clones (strain JC002) carrying a replacement of the HAL3 promoter were tested under different conditions to monitor the terminal phenotype derived from switch off the expression of Hal3 in a sit4 phosphatase deficient-background. Growth of Strain JC002 in Plus/Minus Doxycycline Conditions 1.

20

Preparation of doxycycline plates. Prepare standard YPD/agar medium, autoclave and let it cool down to 55 C. Add doxycycline (doxycycline hydrochloride, Sigma Chemical Co., St. Louis, MO)

A. Wach, C. Brachat, Rebischung, S. Steiner, K. Pokorni, S. Heesen, and P. Philippsen, Yeast gene analysis, in ‘‘Methods in Microbiology’’ (A. J. P. Brown and M. Tuite, eds.), Vol. 26, p. 67. Academic Press, San Diego, 1998.

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from a stock solution (5 mg/ml in 50% ethanol, kept at
20 C) to the desired concentration. In many cases, concentrations ranging from 1 to 5 g/ml are sufficient for optimal inhibition of expression in S. cerevisiae, although concentrations up to 50 g/ml can be used without significant alteration of growth. It might be advisable to test different concentrations to define the best conditions. Note: Regulation of the Tet system is equally effective on S. cerevisiae cells growing exponentially in rich YPD medium or in defined SD medium [0.67% yeast nitrogen base w/o amino acids (Difco), 2% glucose] with the auxotrophic requirements, and in the temperature range from 25 to 37 C. However, in some cases the physiological effects resulting from switching-off gene expression are more evident in rich medium. 2.

Inoculate the cells. Dilute exponentially growing cultures with the same medium up to an OD660 of 0.005. Dispense 3 l of the cultures, being careful that the plates are not too wet (to avoid dispersion of the drop). Allow 15–20 min to absorb the drops and incubate plates for 2–3 days at the desired temperature (normally, 28 C).

As shown in Fig. 4A, strain JC002 grows well in YPD plates in the absence of doxycycline (in fact, it grows better than a sit4 mutant, probably due to that tetO-driven expression of the HAL3 ORF is stronger than from its own promoter). In contrast, when these cells are grown in the presence of 20 g/ml doxycycline essentially no growth is observed even after one week (Fig. 4A). A similar effect can be observed when the same experiment is carried out in liquid rich medium (Fig. 4B). Previous data on the relevance of Sit4 and the Hal3/Ppz1 phosphatase system in cell cycle G1/S transition suggested that the inability of the sit4 hal3 mutant to grow could be due to a strong G1/S arrest. This hypothesis can be easily tested in our conditional mutant. To this end, cultures of the JC002 strain were grown and split in two. One of them was made 20 g/ml doxycycline. Growth was resumed and samples were taken at specific periods of time and processed for DNA content monitoring by flow cytometry, according to Ref. 21. As shown in Fig. 4C, exposure to doxycycline for 12 hr results in a substantial increase in cells with one content of DNA (as JC002 is a haploid strain). At this moment, the percentage of cells with small buds was lower than 25%, while this index

21

C. Gallego, E. Gari, N. Colomina, E. Herrero, and M. Aldea, EMBO J. 16, 7196 (1997).

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FIG. 4. Terminal phenotypes of the conditional S. cerevisiae sit4 tetO : HAL3 phosphatase mutant. Panel A: 3 l of strain JC002 cultures (OD660 0.005) were inoculated on YPD/agar plates with ( þ DOX) or without (
DOX) 20 g/ml doxycycline and growth resumed at 28 C for the indicated periods of time. Panel B: The same starting cultures were inoculated into YPD medium in the presence (dotted line) or the absence (continuous line) of 20 g/ml doxycycline, and growth monitored by measuring the optical density of the culture at 660 nm. Panel C: JC002 cells were grown to an initial OD660 of 0.15 and then split in two aliquots. One of them received 20 g/ml doxycycline. Growth was resumed and samples were taken at the indicated periods of time and processed for DAPI staining and DNA content monitoring by flow cytometry.

was around 70% in cells grown in the absence of doxycycline. These parameters were indicative that a halt in cell cycle at the G1/S transition occurred due to shut-off of the tetO-driven expression of HAL3. This dramatic effect on growth has been used in our laboratory to successfully develop a screening method in search of putative positive effectors of the G1/S transition.22,23 Acknowledgments We thank Gemma Bellı´ and Eloi Garı´ for their contribution to the generation of the dual system constructs, and Josep Clotet and Ernesto Simon for their work in the development and characterization of strain JC002. Work in the laboratory of the authors has been supported by grants PB98-0565-C04-02 and BMC2002-04011-C05-04 from the ‘‘Ministerio de Educacio´n y Cultura’’ to J. A.) and by the EUROFAN program (to E. H.). 22 23

E. Simo´n, J. Clotet, F. Calero, J. Ramos, and J. Arin˜o, J. Biol. Chem. 276, 29740 (2001). I. Mun˜oz, E. Simo´n, N. Casals, J. Clotet, and J. Arin˜o, Yeast 20, 157 (2003).

Section V Knockdown and Knockout Technologies

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Introduction

Protein serine/threonine phosphatases are a diverse group of enzymes that participate in many aspects of cellular signal transduction. Biochemical and molecular biological approaches have provided numerous insights into the molecular structure and regulatory properties of these enzymes. In contrast, much less is known about their physiological functions. This gap in our knowledge is due, in large part, to the lack of inhibitors that can be used for pharmacological suppression of individual phosphatases. None of the naturally occurring phosphatase inhibitors has sufficient specificity and permeability to adequately distinguish individual serine/threonine phosphatases in vivo. For example, okadaic acid is a cell permeable toxin that inhibits PP2A with a Ki of 0.2 nM, PP1 with a Ki of 2 nM, and does not inhibit PP2B/calcineurin, PP2C/PPM, or PP7.1,2 However, PP43,4 and PP55 are just as sensitive to okadaic acid as PP2A. Consequently, even though okadaic acid can be used to distinguish the functions of PP2A from PP1 and the okadaic acid-insensitive enzymes, it cannot distinguish between the PP2A-like enzymes (PP2A, PP4, PP5, and PP6). The inability of current inhibitors to adequately define roles for individual serine/threonine phosphatases is further compounded by the existence of different forms of the same phosphatase. Both PP1 and PP2A are composed of a common catalytic subunit that interacts with a wide variety of targeting and regulatory proteins, each of which has a distinct function. Therefore, the current inhibitors, which are all directed at phosphatase catalytic sites, cannot distinguish between the numerous oligomeric forms of these enzymes in vivo or in vitro. Functional analysis of protein serine/threonine phosphatases would be greatly facilitated by a method capable of ablating the function of individual 1

P. Cohen, C. F. Holmes, and Y. Tsukitani, Trends Biochem. Sci. 15, 98 (1990). X. Huang and R. E. Honkanen, J. Biol. Chem. 273, 1462 (1998). 3 N. D. Brewis, A. J. Street, A. R. Prescott, and P. T. W. Cohen, EMBO J. 12, 987 (1993). 4 C. J. Hastie and P. T. W. Cohen, FEBS Lett. 431, 357 (1998). 5 M. X. Chen, A. E. McPartlin, L. Brown, Y. H. Chen, H. M. Barker, and P. T. Cohen, EMBO J. 13, 4278 (1994). 2

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phosphatase catalytic and regulatory subunits in intact cells. In the past several years, double stranded RNA (dsRNA)-mediated RNA interference (RNAi) has proven to be a useful method to routinely ablate the expression of proteins in Caenorhabditis elegans and Drosophila melanogaster.6–10 More recently, RNAi-based methods for ablating (or ‘‘knocking down’’) the levels of proteins in mammalian cells have been developed.11–15 RNAi makes use of the posttranscriptional gene silencing (PTGS) mechanism that has evolved to protect mammalian cells from harmful transposons or invading viral genetic material. When the PTGS is triggered by sequence-specific dsRNA it is termed RNAi. Double stranded RNA corresponding to a particular mRNA enters Drosophila Schneider 2 (S2) cells when added to the culture medium. Once inside the cell, 100–800 nucleotide dsRNA is cleaved into 21–23 nucleotide smallinterfering RNA (siRNA) by Dicer RNAase III. These 21–22 nucleotide fragments assemble into a multicomponent RNA-induced silencing complex (RISC), which catalyzes the cleavage of target mRNA in a sequence-specific manner.16,17 RNAi in the S2 cell tissue culture system is sequence-specific, dose-dependent, and can spread catalytically from cell to cell.18 We developed an RNA interference strategy to ‘‘knockdown’’ protein phosphatases and their regulatory subunits to examine their functions in Drosophila Schneider 2 tissue culture cells (ATCC#: CRL-1963). The S2 cell line was established from 20 to 24 hr Drosophila embryos and has an epithelial cell morphology.19 While the ease of performing RNAi

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A. M. Silverstein, C. A. Barrow, A. J. Davis, and M. C. Mumby, Proc. Natl. Acad. Sci. U.S.A. 99, 4221 (2002). 7 J. C. Clemens, C. A. Worby, N. Simonson-Leff, M. Muda, T. Maehama, B. A. Hemmings, and J. E. Dixon, Proc. Natl. Acad. Sci. U.S.A. 97, 6499 (2000). 8 R. W. Carthew, Curr. Opin. Cell Biol. 13, 244 (2001). 9 A. Fire, Trends Genet. 15, 358 (1999). 10 C. P. Hunter, Curr. Biol. 9, R440 (1999). 11 T. R. Brummelkamp, R. Bernards, and R. Agami, Science 296, 550 (2002). 12 N. J. Caplen, S. Parrish, F. Imani, A. Fire, and R. A. Morgan, Proc. Natl. Acad. Sci. U.S.A. 98, 9742 (2001). 13 S. M. Elbashir, J. Harborth, W. Lendeckel, A. Yalcin, K. Weber, and T. Tuschl, Nature 411, 494 (2001). 14 P. J. Paddison, A. A. Caudy, and G. J. Hannon, Proc. Natl. Acad. Sci. U.S.A. 99, 1443 (2002). 15 G. Sui, C. Soohoo, el B. Affar, F. Gay, Y. Shi, W. C. Forrester, and Y. Shi, Proc. Natl. Acad. Sci. U.S.A. 99, 5515 (2002). 16 J. Martinez, A. Patkaniowska, H. Urlaub, R. Luhrmann, and T. Tuschl, Cell 110, 563 (2002). 17 T. Tuschl, Chembiochem. 2, 239 (2001). 18 N. J. Caplen, J. Fleenor, A. Fire, and R. A. Morgan, Gene 252, 95 (2000). 19 I. Schneider, J. Embryol. Exp. Morphol. 27, 353 (1972).

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experiments makes them an attractive experimental system, signaling pathways in these cells are poorly characterized compared to mammalian cell lines. However, even though some specific signaling molecules are lacking, it appears that many signaling pathways and signal responsive systems are present in S2 cells. Some of these include MAP kinase pathways,6,7,20,21 G protein-coupled receptors,22,23 steroid receptors,24 the apoptotic pathway,25,6,26 the NF-kappaB pathway,27 JAK/STAT pathways,28,29 the Hedgehog pathway,30 the Wnt signaling pathway,31 pathways controlling the c-myb32 and myo-D33 transcription factors, and pathways regulating the eIF2B translation initiation factor.34 While some of these studies required the introduction of missing components by transfection, they indicate that S2 cells present a viable cell system for studying signal transduction. This chapter describes the application of RNA interference to study the functions of PP2A, PP4, and PP5 in Drosophila S2 cells. However, the functions of any phosphatase, phosphatase regulatory protein, or targeting protein can be studied with this method. In the case of PP2A, Drosophila S2 cells are a particularly attractive experimental system. In contrast to mammals where each PP2A subunit is encoded by multiple genes, Drosophila has a single gene encoding the PP2A scaffold/A subunit,35 the catalytic

20

G. Cornelius and M. Engel, Cell Signal 7, 611 (1995). S. J. Han, K. Y. Choi, P. T. Brey, and W. J. Lee, J. Biol. Chem. 273, 369 (1998). 22 H. Torfs, H. B. Oonk, J. V. Broeck, J. Poels, W. Van Poyer, A. De Loof, F. Guerrero, R. H. Meloen, K. Akerman, and R. J. Nachman, Arch. Insect Biochem. Physiol. 48, 39 (2001). 23 W. Van Poyer, H. Torfs, J. Poels, E. Swinnen, A. De Loof, K. Akerman, and J. Vanden Broeck, Insect Biochem. Mol. Biol. 31, 333 (2001). 24 S. K. Yoshinaga and K. R. Yamamoto, Mol. Endocrinol. 5, 844 (1991). 25 S. Hisahara, H. Kanuka, S. Shoji, S. Yoshikawa, H. Okano, and M. Miura, J. Cell Sci. 111 (Pt 6), 667 (1998). 26 X. Li, A. Scuderi, A. Letsou, and D. M. Virshup, Mol. Cell Biol. 22, 3674 (2002). 27 A. Avila, N. Silverman, M. T. Diaz-Meco, and J. Moscat, Mol. Cell Biol. 22, 8787 (2002). 28 M. Boutros, H. Agaisse, and N. Perrimon, Dev. Cell 3, 711 (2002). 29 S. M. Sweitzer, S. Calvo, M. H. Kraus, D. S. Finbloom, and A. C. Larner, J. Biol. Chem. 270, 16510 (1995). 30 T. Fukumoto, R. Watanabe-Fukunaga, K. Fujisawa, S. Nagata, and R. Fukunaga, J. Biol. Chem. 276, 38441 (2001). 31 S. Yanagawa, J. S. Lee, and A. Ishimoto, J. Biol. Chem. 273, 32353 (1998). 32 M. Okada, H. Akimaru, D. X. Hou, T. Takahashi, and S. Ishii, EMBO J. 21, 675 (2002). 33 Q. Wei, G. Marchler, K. Edington, I. Karsch-Mizrachi, and B. M. Paterson, Dev. Biol. 228, 239 (2000). 34 D. D. Williams, G. D. Pavitt, and C. G. Proud, J. Biol. Chem. 276, 3733 (2001). 35 R. E. Mayer-Jaekel, S. Baumgartner, G. Bilbe, H. Ohkura, D. M. Glover, and B. A. Hemmings, Mol. Biol. Cell 3, 287 (1992). 21

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subunit,36 and the R2/B regulatory subunit.37 The R5/B0 regulatory subunit is encoded by two genes,38 and we identified a single gene encoding the R3/B00 (or PR72) subunit in Drosophila.6 PP439 and PP540 also have a single gene encoding their catalytic subunits in Drosophila. The presence of a limited number of genes encoding protein phosphatase subunits makes interpretation of the results much easier. This chapter provides simple methods for using dsRNA-mediated RNA interference to knockdown specific protein phosphatase subunits in S2 cells.

Identifying Drosophila Protein Phosphatase Homologs and Preparation of cDNA

Drosophila protein phosphatase subunits are identified by carrying out BLAST searches (http://www.fruitfly.org/blast/) of the Berkeley Drosophila Genome Project (BDGP) database using published Drosophila cDNA sequences or the protein sequences of their mammalian homologs. The coding sequences of the targeted genes are used to search the Drosophila EST and cDNA databases to identify corresponding cDNA clones. Multiple cDNA clones of 500–800 nucleotides are identified for each of the targeted phosphatase subunits. It is extremely important to then BLAST the coding sequences of these cDNA clones against the Drosophila cDNA and genome databases to check for significant overlap with other Drosophila proteins. This step helps insure the specificity of the cDNAs that will be used for production of double stranded RNA. Individual Drosophila cDNA clones transformed into bacteria are purchased from Research Genetics (www.resgen.com) (Birmingham, AL). It is important to choose several cDNA clones for each gene. Some of the bacterial stab cultures may have bacteriophage contamination or may not grow. In addition, different dsRNA sequences directed against the same gene can have different knockdown efficiencies. It is necessary to test multiple dsRNA sequences, corresponding to different regions of the mRNA, in

36

S. Orgad, N. D. Brewis, L. Alphey, J. M. Axton, Y. Dudai, and P. T. Cohen, FEBS Lett. 275, 44 (1990). 37 R. E. Mayer-Jaekel, H. Ohkura, R. Gomes, C. E. Sunkel, S. Baumgartner, B. A. Hemmings, and D. M. Glover, Cell 72, 621 (1993). 38 M. Berry and W. Gehring, EMBO J. 19, 2946 (2000). 39 N. R. Helps, N. D. Brewis, K. Lineruth, T. Davis, K. Kaiser, and P. T. Cohen, J. Cell Sci. 111(Pt 10), 1331 (1998). 40 L. Brown, E. B. Borthwick, and P. T. Cohen, Biochim. Biophys. Acta 1492, 470 (2000).

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order to select those that are most effective. Sequencing of the purchased cDNAs is recommended to verify whether the sequences are correct. The cDNAs from Research Genetics are provided in either of two vectors, pBluescript and pOT2, that have been transformed into TOP10F0 bacteria. Upon receipt, streak the bacteria glycerol stocks onto Luria Broth plates 1% Bacto Tryptone (Becton, Dickinson, Co., Sparks, MD), 0.5% Bacto Yeast extract (Becton, Dickinson, Co.), 0.5% NaCl, 0.1% 1 N NaOH, 1.5% Bacto Agar (Becton, Dickinson, Co.), containing 12.5 g/ml chloramphenicol (for the pOT2 vector) or 50 g/ml ampicillin (for the pBluescript vector) and incubate overnight at 37
. Select at least four individual colonies to streak onto new plates that are incubated overnight at 37
. Using a pipette tip, choose at least four individual colonies from each of these plates and dilute into a 25 ml conical tube containing 5 ml of Luria Broth (same recipe as above, but without the Bacto Agar) with the appropriate antibiotic. Incubate overnight at 37
in a rotating incubator set at 250 rpm. Purify plasmid DNA from 4.5 ml of the culture using Qiaprep Miniprep kits (Qiagen) according to the manufacturer’s instructions. Check the identities of the purified cDNAs by restriction enzyme digestion and by sequencing. For sequencing from the 50 end of the cDNA inserts in the pOT2 vector, use a sequence corresponding to the T7 promoter, and for sequencing from the 30 end of the cDNA inserts in the pOT2 vector, use a sequence termed PM001 (50 -CGT TAG AAC GCC GCT ACA AT -30 ) by the BDGP. For sequencing from the 50 end of the cDNA inserts in the pBluescript vector, use a sequence corresponding to the T3 promoter, and for sequencing from the 30 end of the inserts in the pBluescript vector, use a sequence corresponding to the T7 promoter. Identify the correct clones and make 10% glycerol stocks from the remaining 0.5 ml of the overnight bacterial culture and store the bacterial clones at 80
. Complimentary DNA clones for some Drosophila proteins are not available from commercial sources. An example is the R3/PR72 subunit of Drosophila PP2A. In this case, a 700 base pair fragment of the R3/PR72 subunit is amplified by the polymerase chain reaction (PCR) from a 48 hr Drosophila embryo cDNA library (kindly provided by Denis McKearin, Univ. Texas Southwestern Medical Center). PCR primers designed to correspond to coding sequences identified in the Drosophila R3/PR72 gene are engineered to contain bacteriophage T7 RNA polymerase binding sites at the 50 ends of both the sense and antisense primers. PCR is done with Pfu Turbo DNA polymerase (Stratagene) according to the manufacturer’s instructions. Verify the correct size of the PCR product by electrophoretic mobility in a 1% agarose gel, and purify the PCR product from the gel via the QIAquick Gel Extraction Kit

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(Qiagen) following the manufacturer’s protocol. Verify the sequences of the purified PCR fragments by sequencing using the same primers used for PCR.

Production of Double Stranded RNA

Production of dsRNA involves two steps: amplification of cDNA fragments and synthesis of RNA using the PCR fragments as templates. One microgram of the phosphatase encoding plasmids or PCR product are used as templates in PCR reactions in which both the sense and antisense primers contain a T7 polymerase binding site (50 -GAA TTA ATA CGA CTC ACT ATA GGG AGA -30 ) at the 50 end as described above. PCR conditions for the individual clones are optimized using the Optiprime PCR Optimization Kit (Stratagene) according to the manufacturer’s instructions. Clean the PCR products of excess nucleotides and polymerase in a Microcon spin concentrator (Millipore) by following the manufacturer’s instructions and dilute in sterile ddH2O to a final concentration of 200 ng/l, determined by OD260. Use 1 g of PCR product to synthesize dsRNA using a large-scale T7 transcription kit from Novagen (Madison, WI) according to the manufacturer’s instructions. PCR products with a T7 polymerase binding site on the 50 end of both the sense and antisense strands permits the synthesis of the sense and antisense RNA strands in the same reaction. Then precipitate the RNA by adding 0.1 volumes of 3 M sodium acetate, pH 5.2 and 2.5 volumes of 100% ethanol. Mix the samples gently and incubate at 20
for 30 min. After centrifugation at 15,000 rpm for 15 min in a microcentrifuge, aspirate the supernatant and wash the RNA pellet with 100 l of 70% ethanol. Centrifuge the sample at 15,000 rpm for 15 min in a microcentrifuge and remove the supernatant fraction by aspiration. Resuspend the RNA pellet in 50 l sterile, RNAse-free water. To promote annealing of the single stranded RNA, incubate the samples for 30 min at 65
in a beaker of water placed in a water bath. Move the beaker to a bench top and allow it to cool to room temperature. Determine the dsRNA concentration by absorbance at 280 nm using the formula [dsRNA] ¼ (OD260nm)(dilution factor)(45 g/ml), and dilute all dsRNAs to a final concentration of 3 mg/ml in RNAse-free water (provided with the T7 transcription kit) and store at 70
. As a negative control for the nonspecific effects caused by treating cells with dsRNA, the pEGFP-C3 vector (Clontech) is used as a template to make enhanced green fluorescent protein (EGFP) dsRNA as described above. EGFP is unlikely to promote RNAi of Drosophila genes.

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Cell Culture and dsRNA Treatment

Serum free medium (SFM)-adapted Schneider S2 cells (D.Mel-2) and Drosophila SFM are from Life Technologies. S2 cells are maintained in Drosophila SFM supplemented with 16.5 mM L-glutamine and 46 g/ml gentamicin at 28
, 0% CO2 in T-75 EasyFlasks with Filtercaps (Nalge Nunc International, Rochester, NY). Nonconfluent S2 cells grow as adherent cells, and continue to grow in suspension once they have become confluent. S2 cells are passaged every 3–4 days by removing 1 ml of the medium containing nonadherent cells to a new T-75 flask containing 9 ml of room temperature Drosophila SFM. For dsRNA treatments, S2 cells are harvested from flasks by triturating the medium in order to wash as many adherent cells as possible from the bottom of the flask. The triturated S2 cell suspension is removed to a 50 ml conical tube and the cells are pelleted at 1000 rpm for 5 min. The medium is aspirated, 10 ml of room temperature Drosophila SFM is added, and the cells are resuspended by repeated pipetting with a 10 ml pipette. Count the resuspended S2 cells using a hemacytometer and dilute to 1  106 cells/ml in Drosophila SFM. For each individual knockdown experiment, pipette 1 ml of the cell suspension into the wells (35 mm) of a 6 well Multidish (Nalge Nunc International) and add 15 g of dsRNA using a pipette. Mix by gently rocking the plate. For replicate samples, a larger volume can be prepared by mixing 15 g of dsRNA per each 1 ml of cell suspension by triturating in a 15 ml conical tube and then immediately pipetting into individual wells. The cells are incubated with dsRNA for approximately 4.5 hr at 28
, and then diluted by the addition of 2 ml of room temperature Drosophila SFM. The dsRNA is left in the dish until the cells are harvested. For the phosphatase subunits tested, maximal knockdown is achieved after 72 hr in the presence of dsRNA. Note that incubation times need to be varied depending on the half-life of individual proteins and the dsRNAs used to target them. It is recommended that a time course following the disappearance of mRNA or, preferably, protein be carried out with each RNAi target.

Cytosol Preparation and Western Blotting

Following incubation with dsRNA, most of the cells are adherent and the medium is removed by aspiration. The cells are lysed by adding 200 l of RIPA buffer (20 mM Tris, pH 8.0; 150 mM NaCl; 1% Nonidet P-40; 0.5% Deoxycholate; 0.1% SDS; 0.2% sodium vanadate; 10 mM sodium fluoride; 0.4 mM EDTA; and 10% glycerol) to the wells and incubating for 5 min at room temperature. A cell scraper is used to remove any

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residual adherent cells before transferring the lysis buffer to 1.5 ml microcentrifuge tubes. Centrifuge the tubes at 14,000 rpm for 10 min at 4
in a microcentrifuge, and transfer the supernatant fractions to new 1.5 ml microcentrifuge tubes. Determine the protein concentrations using the bicinchoninic acid (BCA) protein assay (Pierce) according to the manufacturer’s protocol using 5 l of sample and 500 l of the BCA reagent. When antibodies are available, the knockdown of proteins targeting by RNAi can be determined by Western blotting. Equal amounts of protein (30–60 g) from individual RNAi experiments are boiled for 10 min in 2X Laemmli SDS sample buffer (0.125 M Tris, pH 6.8; 4% SDS; 20% glycerol; 0.01% bromophenol blue; and 10% b-mercaptoethanol), and proteins are resolved by electrophoresis on 10% SDS-polyacrylamide gels. Proteins are transferred to PROTRAN nitrocellulose (Schleicher & Schuell, Keene, NH) by electroblotting overnight at 0.15 amps. The nitrocellulose membranes are then incubated in blocking buffer (25 mM Tris, pH 7.4; 150 mM NaCl; 0.1% Tween 20; and 0.5% nonfat dry milk) for 20 min at room temperature. The protein-containing membranes are cut horizontally with a razor blade to generate strips corresponding to different molecular weight ranges. The alignment of the cuts is determined using the Kaleidoscope prestained molecular weight markers (Biorad, Hercules, CA) that are applied to the SDS gel and transferred to the membranes along with the samples. Cutting the membranes into separate strips allows detection of multiple proteins, using different primary and secondary antibodies, within a single transfer experiment. Individual strips of membrane are incubated overnight at 4
with gentle rocking in blocking buffer containing 0.02–0.1% v/v of the appropriate primary antibody. For detection of PP2A subunits, PP4, and PP5, the following antibodies have been used: F725 antiserum against the A subunit of PP2A;41 C-20 antiserum against the PP2A catalytic subunit (Affinity Bioreagents, Neshanic Station, NJ); M878 antiserum against the PP2A B0 /B56 subunit;41 anti-PP5 antiserum (kindly provided by Michael Chinkers, Univ. South Alabama, Mobile, AL); and affinity purified anti-PP4 polyclonal antibody (kindly provided by Brian Wadzinski, Vanderbilt University, Nashville, TN). Wash the membranes three times for 5 min in TBST (25 mM Tris, pH 7.4; 150 mM NaCl; 0.1% Tween 20) and incubate for one hour with the appropriate anti-mouse or anti-rabbit secondary antibodies that are conjugated to horseradish peroxidase. Wash the membranes four times for 10 min in TBST and once for 5 min 41

C. Kamibayashi, R. Estes, R. L. Lickteig, S.-I. Yang, C. Craft, and M. C. Mumby, J. Biol. Chem. 269, 20139 (1994).

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in PBS (10 mM Na2HPO4; 3 mM KH2PO4; 3 mM KCl; 137 mM NaCl, pH 7.2). Immunoreactive bands are then detected by enhanced chemiluminescence with ECL Western Blotting Detection Reagents (Amersham Biosciences). In some cases, antibodies against mammalian protein phosphatase subunits do not cross-react with the corresponding Drosophila protein. An example of this is the R2/B subunit of PP2A. In this case a peptide corresponding to a putative immunogenic peptide derived from the Drosophila protein sequence can be sent to Capralogics (Hardwick, MA) for production of the appropriate rabbit polyclonal antibody. This antibody can then be used for Western blotting as described above. The Western blot results from a typical RNAi experiment are shown in Fig. 1A. The method results in nearly complete loss of detectable protein for most of the phosphatase subunits tested. The most notable exception is the R2/B subunit of PP2A where significant immunoreactivity is still detected after 72 hr in the presence of dsRNA. The protein band present in samples treated with dsRNA for the A subunit of PP2A is a cross-reacting protein that migrates just below the A subunit (top row of Fig. 1A). As reported previously,6 knockdown of either the A or C subunits of PP2A causes a corresponding loss of R5/B56-1 and R2/B subunit proteins.

Reverse Transcription (RT)-PCR

In some cases, antibodies of adequate specificity for Drosophila proteins cannot be produced. Examples include the Drosophila R5/B56-2 and R3/PR72 homologs of PP2A subunits. In these cases, the effectiveness of the RNAi can be monitored by loss of mRNA using the reverse transcriptionpolymerase chain reaction (RT-PCR). S2 cells in 35 mm dishes are treated with dsRNA for 72 hr as described above and total RNA is isolated using the RNeasy Mini Kit (Qiagen) according to the manufacturer’s instructions. RT-PCR is performed with the SUPERSCRIPT One-Step RT-PCR with PLATINUM Taq kit (Invitrogen) as described by the manufacturer. Total RNA (1.5 g) is used as the template, and the primers are identical to those used to make PCR fragments for RNA production described above. Five microliter (1/10) of the RT-PCR reaction mixture is resolved on a 1% agarose gel. An example of the effects of RNAi on phosphatase subunit mRNA detected by RT-PCR is shown in Fig. 1B. In all cases, no phosphatase mRNA is detected in cells treated with the corresponding dsRNA. Due to the sensitivity of RT-PCR, the absence of PCR products in samples treated with specific dsRNAs and the presence of strong signals in

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FIG. 1. Knockdown of protein phosphatase 2A, 4, and 5 subunits in Drosophila S2 cells by RNAi. Drosophila S2 cells were incubated with dsRNA targeted to EGFP or to the specific phosphatase subunits indicated at the top of each panel for 72 hr. (A) Equal amounts of protein (30–60 g) were separated on SDS-PAGE gels and phosphatase subunits were detected by Western blotting with antibodies against the proteins indicated at the right. (B) Total RNA was extracted from the cells treated with the dsRNA indicated at the top. RT-PCR was performed as described in the text using PCR primers for the phosphatase subunits indicated across the bottom.

the sample treated with the control EGFP is a definitive indicator of a positive RNAi effect. However, care must be taken when interpreting these results at the level of the corresponding proteins. Other properties, including the half-life of the protein, must be taken into account. Even though RNAi ablates the mRNA, substantial amounts of the protein may still be present in the cells. Once knockdown of the target protein has been verified, the physiological consequences of ablating the protein can be determined in dsRNA-treated cells. The readouts that will be assayed vary with the target

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and are usually predicated on some knowledge of the potential roles of the proteins. Physiological and signaling parameters that can be examined include, but are not limited to, changes in cell morphology, changes in cell growth, or changes in one or more of the signaling pathways described in ‘‘Introduction.’’ As discussed below, the use of RNAi can also lead to the discovery of novel functions of protein phosphatases.

Summary

Double stranded RNA-mediated RNA interference is an effective method to downregulate the levels of protein phosphatases in Drosophila S2 cells. In many cases, nearly complete ablation of the targeted protein can be achieved. RNAi-mediated knockdown of protein phosphatases is akin to pharmacological inhibition with drugs and can be used to determine the roles of specific protein phosphatases in intact cells. RNAi can avoid the problems associated with less than adequate specificity of phosphatase inhibitors. Although information about the signaling pathways present in Drosophila S2 cells is not as well developed as many mammalian cell lines, the Drosophila system is particularly attractive for the study of oligomeric phosphatases like PP2A. Drosophila has far fewer isoforms for the phosphatases we have examined. This is especially true of the genes for PP2A regulatory subunits where over 50 isoforms are present in mammals but only four are present in Drosophila. Once hypotheses regarding phosphatase function have been generated from RNAi experiments in S2 cells, they can potentially be tested utilizing recent advances in the use of siRNAs to conduct RNAi experiments in mammalian cell lines.11,13–15,42 RNAi in Drosophila S2 cells has proven to be a powerful technique for identifying physiological functions of signaling proteins.7 The RNAi method is straightforward and works routinely with almost all proteins. RNAi in S2 cells can be used to assess the role of signaling proteins in specific pathways and as a screening tool to identify new roles for signaling molecules. For example, results from RNAi analysis of PP2A show that regulation of MAP kinase signaling involves the R2/B regulatory subunit and that the R5/B56 subunits play a previously unidentified role in apoptosis.6 While RNAi in Drosophila S2 cells is a powerful tool for analyzing protein function, the method does have limitations. Foremost, cells may exhibit an RNAi response to any nonspecific dsRNA, even in the absence of interferon.43 Therefore, 42 43

T. Tuschl, Nat. Biotechnol. 20, 446 (2002). G. Geiss, G. Jin, J. Guo, R. Bumgarner, M. G. Katze, and G. C. Sen, J. Biol. Chem. 276, 30178 (2001).

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physiological processes that respond to nonspecific dsRNA will be difficult to study. A second limitation is the need to produce antibodies that react with Drosophila isoforms. We have found that many antibodies to mammalian protein phosphatases do not cross-react with the corresponding Drosophila proteins. Finally, the physiology and signaling pathways of S2 cells have not been extensively studied. This lack of information limits the number of available readouts that can be used when assessing the effects of protein knockdowns.

[27] Regulating the Expression of Protein Phosphatase Type 5 By TERESA A. GOLDEN and RICHARD E. HONKANEN

Introduction Serine/threonine Phosphatase type 5 (PP5)

Serine/threonine phosphatase 5 (PP5) is an okadaic acid, microcystin, calyculin A sensitive phosphatase that is highly conserved among species and expressed ubiquitously in mammalian tissues. PP5 has a catalytic domain that is structurally similar to that of PP1–PP7 family of phosphatases, yet PP5 contains an extended N-terminal region with three TPR (tetratricopeptide repeat) domains (Fig. 1). PP5 has been reported to associate with the atrial natriuretic peptide receptor,1 the heat shock protein 90 (Hsp-90)-glucocorticoid receptor (GR)-heterocomplex,2 cryptochrome 2,3 the CDC16/CDC27 subunits of the anaphase-promoting complex,4 apoptosis signal-regulating kinase 1 (ASK1),5 Hsp90-dependent hemeregulated eIF2alpha kinase,6 and the G12-alpha/G13-alpha subunits of heterotrimeric G proteins.7 In estrogen responsive breast carcinoma cells, the expression of PP5 is induced by treatment with estrogen, and the constitutive expression of PP5 converts MCF-7 cells into an estrogen 1

M. Chinkers, Proc. Natl. Acad. Sci. U.S.A. 91, 11075 (1994). M. S. Chen, A. M. Silverstein, W. B. Pratt, and M. Chinkers, J. Biol. Chem. 271, 32315 (1996). 3 S. Zhao and A. Sancar, Photochem. Photobiol. 66, 727 (1997). 4 V. Ollendorff and D. J. Donoghue, J. Biol. Chem. 272, 32011 (1997). 5 K. Morita, M. Saitoh, K. Tobiume, H. Matsuura, S. Enomoto, H. Nishitoh, and H. Ichijo, EMBO J. 20, 6028 (2001). 6 J. Shao, S. D. Hartson, and R. L. Matts, Biochemistry 41, 6770 (2002). 7 Y. Yamaguchi, H. Katoh, K. Mori, and M. Negishi, Curr. Biol. 12, 135 (2002). 2

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physiological processes that respond to nonspecific dsRNA will be difficult to study. A second limitation is the need to produce antibodies that react with Drosophila isoforms. We have found that many antibodies to mammalian protein phosphatases do not cross-react with the corresponding Drosophila proteins. Finally, the physiology and signaling pathways of S2 cells have not been extensively studied. This lack of information limits the number of available readouts that can be used when assessing the effects of protein knockdowns.

[28] Regulating the Expression of Protein Phosphatase Type 5 By TERESA A. GOLDEN and RICHARD E. HONKANEN

Introduction Serine/threonine Phosphatase type 5 (PP5)

Serine/threonine phosphatase 5 (PP5) is an okadaic acid, microcystin, calyculin A sensitive phosphatase that is highly conserved among species and expressed ubiquitously in mammalian tissues. PP5 has a catalytic domain that is structurally similar to that of PP1–PP7 family of phosphatases, yet PP5 contains an extended N-terminal region with three TPR (tetratricopeptide repeat) domains (Fig. 1). PP5 has been reported to associate with the atrial natriuretic peptide receptor,1 the heat shock protein 90 (Hsp-90)-glucocorticoid receptor (GR)-heterocomplex,2 cryptochrome 2,3 the CDC16/CDC27 subunits of the anaphase-promoting complex,4 apoptosis signal-regulating kinase 1 (ASK1),5 Hsp90-dependent hemeregulated eIF2alpha kinase,6 and the G12-alpha/G13-alpha subunits of heterotrimeric G proteins.7 In estrogen responsive breast carcinoma cells, the expression of PP5 is induced by treatment with estrogen, and the constitutive expression of PP5 converts MCF-7 cells into an estrogen 1

M. Chinkers, Proc. Natl. Acad. Sci. U.S.A. 91, 11075 (1994). M. S. Chen, A. M. Silverstein, W. B. Pratt, and M. Chinkers, J. Biol. Chem. 271, 32315 (1996). 3 S. Zhao and A. Sancar, Photochem. Photobiol. 66, 727 (1997). 4 V. Ollendorff and D. J. Donoghue, J. Biol. Chem. 272, 32011 (1997). 5 K. Morita, M. Saitoh, K. Tobiume, H. Matsuura, S. Enomoto, H. Nishitoh, and H. Ichijo, EMBO J. 20, 6028 (2001). 6 J. Shao, S. D. Hartson, and R. L. Matts, Biochemistry 41, 6770 (2002). 7 Y. Yamaguchi, H. Katoh, K. Mori, and M. Negishi, Curr. Biol. 12, 135 (2002). 2

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FIG. 1. Homology of the Ser/thr protein phosphatases. A schematic comparison of the PP1PP7 family of PPases. The PPases contain a common catalytic core domain that is extremely conserved among species. PP1, PP2A, PP4 and PP6 are highly homologous enzymes, differing primarily in their N- and C-terminal domains. PP5 contains an extended N-terminal region with three tetratricopeptide (TPR) domains. PP2B differs in that it contains a Ca2 þ -calmodulin (CaM) binding site. PP7 contains five EF- and EF-like hand domains (indicated by black boxes) in the C-terminal calcium binding domain. Both PP2B and PP7 contain an insert in the catalytic core (indicated by open boxes) that alters the okadaic acid/microcystin toxin binding sites contained in PP1, PP2A, PP4 and PP5.

independent phenotype.8 Nonetheless, determining the physiological/ pathological roles of PP5 has proven difficult for many reasons. First, in crude cell homogenates PP5 exists predominately in an inactive state. Thus, in phosphatase assays the basal activity of PP5 is masked by the abundant activity of PP1 and PP2A, which are also expressed ubiquitously at high levels. Second, the physiological activators of PP5 are unknown. In vitro PP5 can be activated by a brief treatment with trypsin, which cleaves the Nterminal domain and relieves auto-inhibitory activity. Still, to date, there is no direct evidence that PP5 is activated in vivo by a protease. PP5 can also become activated by the addition of polyunsaturated lipids, yet again the physiological relevance of lipid activation remains to be determined. Finally, to date no selective small molecule inhibitors of PP5 have been reported, and genetic studies in Drosophila suggest that the lack of PP5 expression 8

G. Urban, T. Golden, I. V. Aragon, J. G. Scammell, N. M. Dean, and R. E. Honkanen, J. Biol. Chem. 276, 27638 (2001).

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results in an embryonic lethal phenotype.9 Therefore, a reliable method to measure direct changes in the activity of PP5 in vivo or in crude cell homogenates is currently lacking.

Antisense and Its Proper Use

Although studying changes in PP5 activity is difficult, the expression of PP5 in mammalian cells can be suppressed by treatment with antisense oligonucleotides or small double stranded molecules of RNA. Antisense oligonucleotides are synthetic, small DNA molecules, designed to bind in a Watson-Crick fashion to a target mRNA. For optimal use in tissue culture or in animals there are several important considerations for antisense design. Most importantly the oligonucleotides must have sufficient affinity and specificity of hybridization to the target mRNA such that binding blocks or inactivates its function. mRNA inactivation is achieved primarily via RNase-H-mediated degradation of hybridized mRNA, with antisense targeting any region in which the molecule binds with sufficient affinity (Fig. 2). This includes the 50 and 30 -untranslated regions of the target mRNA. Other mechanisms include the steric blockade of ribosomal subunit attachment to the target mRNA at the 50 capsite and interference with proper mRNA splicing via antisense binding to splice acceptor or splice donor sites. To be useful the oligonucleotides must also be at least partially resistant to both 30 and 50 exonucleases and to endonucleases, which is necessary to prevent their rapid cleavage and degradation. The desired cells and tissues of interest must also take up the oligonucleotides in sufficient quantities to produce a biological response in vivo, and the oligonucleotides must not bind tightly to proteins of the serum, complement, or clotting pathways. These issues and strategies to address them will be examined further below.

Chemical Composition

Due to their natural susceptibility to exo- and endonucleases, unmodified small oligodeoxynucleotides are rapidly degraded in cells or in the serum. Specific modifications can improve resistance against endogenous nucleases otherwise oligodeoxynucleotides would not be useful in most biological systems. The most commonly employed and commercially readily available form of antisense oligonucleotides is the phosphorothioate oligodeoxynucleotides. In these oligonucleotides a nonbridging oxygen 9

L. Brown, E. B. Borthwick, and P. T. Cohen, Biochim. Biophys. Acta 1492, 470 (2000).

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FIG. 2. Antisense: Mechanism of action. (1) Antisense is introduced into cells via transfection using cationic liposomes. (2) The antisense oligonucleotides bind a complementarily site of the target mRNA by forming a local duplex structure via hydrogen bonding. The specificity for the target mRNA is derived from the encoded bases of the antisense oligonucleotide. (3) The antisense bound mRNA is recognized by RNase H, which hydrolyzes the phosphodiester backbone of the target mRNA. (4) Lacking protection provided by the 50 cap or the poly-A tail, the target mRNA is then rapidly degraded by endogenous nucleases.

atom of the internucleoside phosphate group is replaced with sulphur, forming a phosphorothioate backbone. These phosphorothioate (PS) containing oligonucleotides have substantial resistance to nucleases compared with phosphodiester oligodeoxynucleotides, and despite the potential pitfalls of phosphorothioate oligodeoxynucleotides, they can be used effectively to show true antisense-mediated inhibition of gene expression. However, the incorporation of the sulphur comes at a cost, with each incorporation of a PS generating a chiral center that reduces the hybridization affinity for the targeted mRNA by 1–1.5
C.10,11 To compensate for the lower affinity, phosphorothioate oligodeoxynucleotides must often be employed at higher concentrations than other chemical modifications, which can produce side effects that do not arise from the suppression of the target protein. True-antisense gene suppression by bound phosphorothioate oligodeoxynucleotides is likely mediated by an RNase-H catalyzed mechanism of degradation of the target message (Fig. 2). 10 11

S. Akhtar, R. Kole, and R. L. Juliano, Life Sci. 49, 1793 (1991). J. M. Campbell, T. A. Bacon, and E. Wickstrom, J. Biochem. Biophys. Methods 20, 259 (1990).

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To overcome the biological limitations of PS-oligodeoxynucleotides (reviewed in Agrawal,12 and Levin13) the development of additional oligonucleotide chemistries was essential. Improvements have lead to new ‘‘second-generation’’ chemistries demonstrating increased activity, reduced toxicity and a reduction in nonantisense effects associated with the use of PSoligodeoxynucleotides. Most useful modifications consist of additions to the 20 -carbon of the ribose,14 and there are several, such as: 20 -fluoribonucleotides,15 20 -propoxyribonucleotides (20 -O-propyl),16 and 20 -O-(2-methoxy) ethyl ribonucleotides (20 -MOE).17 Additions of 20 -O-propyl or 20 -MOE improve both nuclease resistance and hybridization affinity. Nonetheless, many of these modifications do not support the RNase-H mediated cleavage, which appears to greatly limit their usefulness in a biological setting. Many backbone modifications have also been tested, including oligonucleotide conjugates,18 methylphosphonate,19–21 phosphoramidate,22,23 morpholino,24,25 amide,26 peptide nucleic acids (PNA),27,28 boranophosphate,29 methylene (methylimino) (MMI),30 and 30 -methylene 12

S. Agrawal, Biochim. Biophys. Acta 1489, 53 (1999). A. Levin, Biochim. Biophys. Acta 1489, 69 (1999). 14 N. M. Dean and R. H. Griffey, Antisense Nucl. Acid Drug Dev. 7, 229 (1997). 15 A. M. Kawasaki, M. D. Casper, S. M. Freier, E. A. Lesnik, M. C. Zounes, L. L. Cummins, C. Gonzalez, and P. D. Cook, J. Med. Chem. 36, 831 (1993). 16 R. A. McKay, L. L. Cummins, M. J. Graham, E. A. Lesnik, S. R. Owens, M. Winniman, and N. M. Dean, Nucl. Acids Res. 24, 411 (1996). 17 R. A. McKay, L. Miraglia, L. Cummins, S. Owens, H. Sasmor, and N. M. Dean, J. Biol. Chem. 274, 1715 (1999). 18 M. Manoharan, K. L. Tivel, and L. K. Andrade, Nucleosides Nucleotides 14, 969 (1995). 19 M. D. Disney, S. M. Testa, and D. H. Turner, Biochemistry 39, 6991 (2000). 20 P. S. Miller, R. A. Cassidy, T. Hamma, and N. S. Kondo, Pharmacol. Ther. 85, 159 (2000). 21 M. A. Reynolds, R. I. Hogrefe, J. A. Jaeger, D. A. Schwartz, T. A. Riley, W. B. Narvin, W. J. Daily, M. M. Vaghefi, T. A. Beck, S. K. Knowles, R. E. Klem, and L. J. Arnold, Nucl. Acids Res. 24, 4584 (1996). 22 J. M. Dagle, M. E. Andracki, R. J. DeVine, and J. A. Walder, Nucl. Acids Res. 19, 1805 (1991). 23 S. M. Gryaznov, Biochim. Biophys. Acta 1489, 131 (1999). 24 R. M. Hudziak, J. Summerton, D. D. Weller, and P. L. Iversen, Antisense Nucl. Acid Drug Dev. 10, 163 (2000). 25 J. Summerton, Biochim. Biophys. Acta 1489, 141 (1999). 26 A. DeMesmaeker, K. H. Altmann, A. Waldner, and S. Wendeborn, Curr. Opin. Struct. Biol. 5, 343 (1995). 27 O. Buchardt, M. Egholm, R. H. Berg, and P. E. Neilsen, Trends Biotechnol. 11, 384 (1993). 28 H. J. Larson, T. Bentin, and P. E. Nielsen, Biochim. Biophys. Acta 1489, 159 (1999). 29 B. R. Shaw, D. Sergueev, K. He, K. Porter, J. Summers, Z. Sergueeva, and V. Rait, Methods Enzymol. 313, 226 (2000). 30 H. Kishioka, N. Fukuda, M. Nakayama, W. Y. Hu, C. Satoh, K. Kanmatsuse, and M. Manoharan, Eur. J. Pharmacol. 392, 129 (2000). 13

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derivatives.31 Like the modifications of the ribose mentioned, details of how these modifications are designed and how they function can be found in the papers cited. Briefly, each modification has its specific uses, benefits, and costs.

Experimental Design and Interpretations

The simple concept of using antisense to inhibit or alter expression of a target protein often belies a very real need for the understanding of its limitations and the absolute necessity of controls. Issues that must be addressed involve: the selection of a specific sequence as a target, the selection of which chemistries will be used, and finally, the uptake of the oligonucleotide by the desired cells or tissues. Although there are published recommendations for the selection of antisense target sites, there are no guaranteed formulas to preselect the most effective target sequence for a given gene. This is likely due to proteins bound to the mRNA in vivo, RNA secondary structure, and/or accessibility of RNase H to bound mRNA. Therefore, a good initial, but sometimes costly strategy, involves the synthesis and evaluation of  15–20 oligonucleotides that target a number of different sequences selected from sites along the entire length of the gene to find an active antisense oligonucleotide (see below). Other more involved methods are based on predictions of secondary structure, combinatorial screening, and binding of mRNA transcripts to an array of oligonucleotides (reviewed in Cooper et al.32). There are at least some general guidelines that have been developed to help eliminate problem sequences.33 Antisense sequences should have a high Tm with no self-complementary regions. Runs of four contiguous guanine residues must also be avoided due to their ability to create a secondary structure that can prevent Watson-Crick hybridization.34 The immune system can also be effected by the use of palindromes of six bases and greater or CG dinucleotides, so these too should also be avoided.35–37 31

H. An, T. Wang, M. A. Maier, M. Manoharan, B. S. Ross, and P. D. Cook, J. Org. Chem. 66, 2789 (2001). 32 S. R. Cooper, J. K. Taylor, L. J. Miraglia, and N. M. Dean, Pharmacol. Ther. 82, 427 (1999). 33 K. J. Myers and N. M. Dean, Trends Pharmacol. Sci. 21, 19 (2000). 34 J. R. Wyatt, P. W. Davis, and S. M. Freier, Biochemistry 35, 8002 (1996). 35 A. M. Krieg, Biochim. Biophys. Acta 1489, 107 (1999). 36 D. K. Monteith, S. P. Henry, R. B. Howard, S. Flournoy, A. A. Levin, C. F. Bennett, and S. T. Crooke, Anticancer Drug Des. 12, 421 (1997). 37 S. Yamamoto, T. Yamamoto, T. Kataoka, E. Kuramoto, O. Yano, and T. Tokunaga, J. Immunol. 148, 4072 (1992).

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Productive use of antisense is also critically dependent on the selection of the correct chemistry for oligonucleotide synthesis. For example, it is known that the highly charged PS-backbones have a strong propensity to bind serum and cellular proteins in a nonspecific manner, which may contribute to minor toxicities commonly found in whole animal antisense therapy (i.e., a decrease in blood clotting time and complement activation38,39). Therefore, when PS-oligonucleotides are employed, phenotypes observed in tissue culture cells need to be examined carefully to verify that they are produced as a result of a true antisense effect. Ways to verify a true antisense effect include: conducting dose-response studies, careful assessment to determine if there is a decrease only in the expression of the target protein, and by a lack of an effect when mismatched control oligonucleotides are employed. On the other hand, occasionally nonspecific binding may provide a slight advantage, by helping to retain the oligonucleotides in a certain tissue for a greater duration. Modifications like the 20 -MOE have advantages over the phosphorothioate alone due to increased plasma stability, but if 20 -MOEs are used with a phosphodiester backbone instead of a phosphorothioate backbone the oligos are rapidly cleared from the plasma.40 The selection of the appropriate chemistry therefore involves a strategy of balancing the costs and benefits of each chemistry with the ultimate experimental or drug development goals. Another important factor that must be considered in any antisense experiment is the limitations associated with cellular uptake and retention. For tissue culture, many agents that enhance transfection have been described, and these agents are reviewed by Dean et al.41 Cationic lipids, such as LipofectinÕ , are an efficient means of getting oligonucleotides into many types of cells and are therefore widely used. Employing cationic lipids, the inhibition profile of phosphorothioate oligonucleotides is reproducible. For a constitutively expressed message a decrease of approximately 80% is expected using oligonucleotide at doses between 0.2 and 1 M.42 The antisense response typically lasts 24–48 hr, and may differ with cell lines. For proteins with long half-lives more stable oligonucleotide chemistries or

38

W. M. Galbraith, W. C. Hobson, P. C. Giclas, P. J. Schechter, and S. Agrawal, Antisense Res. Dev. 4, 201 (1994). 39 S. P. Henry, W. Novotny, J. Leeds, C. Auletta, and D. J. Kornbrust, Antisense Nucl. Acid Drug Dev. 7, 503 (1997). 40 R. S. Geary, T. A. Watanabe, L. Truong, S. Freier, E. A. Lesnik, N. B. Sioufi, H. Sasmor, M. Manoharan, and A. A. Levin, J. Pharmocol. Exp. Ther. 296, 890 (2001). 41 N. M. Dean, S. R. Cooper, W. Shanahan, J. Taylor, and K. Myers, J. Clinical Ligand Assay 23, 43 (2000). 42 N. M. Dean, R. McKay, T. P. Condon, and C. F. Bennett, J. Biol. Chem. 269, 16416 (1994).

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repeated treatments may be required. The nuclear accumulation of oligonucleotides is temporally associated with the reduction of target gene expression. This suggests that the nucleus is the functional site of the oligo. Therefore cationic lipids are widely used to deliver antisense to cultured cells because they provide the added advantage of promoting oligonucleotide accumulation in the nucleus.43,44 Unlike in tissue culture cells, in animals the oligonucleotides can exert effects without the addition of delivery enhancing agents.45 The pharmacology of antisense oligonucleotides has been examined in many animal species46,47 (see reviews Dean et al.,41 Marcusson et al.48). Following intravenous injection, both oligonucleotides composed of phosphorothioates as well as those containing certain 20 -C modifications distribute rapidly into liver, kidney, bone marrow, skeletal muscle and skin.47,49,50 PS-oligonucleotides do not cross the blood-brain barrier, and they are not orally bioavailable.51 Oligonucleotides containing 20 -C modifications often have improved permeability over PS-oligonucleotides.48 Still, hybrid oligonucleotides containing 20 -C modifications have also demonstrated decreased distribution and/or concentration in certain tissues.40

Screening for a Useful Antisense Molecule

To account for possible nonantisense effects and obtain meaningful data from experiments utilizing antisense oligonucleotides, it is absolutely necessary to conduct numerous controls. To illustrate the methods and the controls needed for the identification of an antisense oligonucleotide capable of inhibiting the expression of a single human protein phosphatase, we will 43

C. F. Bennett, M. Y. Chiang, H. Chan, J. E. E. Shoemaker, and C. K. Mirabelli, Mol. Pharmacol. 41, 1023 (1992). 44 E. G. Marcusson, B. Bhat, M. Manoharan, C. F. Bennett, and N. M. Dean, Nucl. Acids Res. 26, 2016 (1998). 45 N. M. Dean and R. McKay, Proc. Natl. Acad. Sci. U.S.A. 91, 11762 (1994). 46 S. Agrawal, J. Temsamani, W. Galbraith, and J. Tang, Clin. Pharmacokinet. 28, 7 (1995). 47 P. A. Cossum, H. Sasmor, D. Dellinger, L. Truong, L. Cummins, S. R. Owens, P. M. Markham, J. P. Shea, and S. Crooke, J. Pharmacol. Exp. Ther. 267, 1181 (1993). 48 E. G. Marcusson, B. R. Yacyshyn, W. R. Shanahan, and N. M. Dean, Molec. Biotech. 12, 1 (1999). 49 S. Agrawal, J. Temsamani, and J. Y. Tang, Proc. Natl. Acad. Sci. U.S.A. 88, 7595 (1991). 50 H. Sands, L. J. Gorey-Feret, A. J. Cocuzza, F. W. Hobbs, D. Chidester, and G. L. Trainor, Mol. Pharmacol. 45, 932 (1994). 51 P. L. Nicklin, D. Bayley, J. Giddings, S. J. Craig, L. L. Cummins, J. G. Hastewell, and J. Phillips, Pharm. Res. 15, 583 (1998).

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FIG. 3. Illustration of the ‘‘gap’’ and ‘‘wing’’ structure contained in chimeric antisense oligonucleotides. The winged region contains 20 -O-methoxyethylribonucleotide (20 -MOE) that provide high affinity binding to the target mRNA. The wings flank a phosphorothioate core, also referred to as a PS gap, that supports RNase H recognition. Examples of 20 -MOE modifications are highlighted by the gray outline, and an S indicates internucleoside sulfur groups. These chimeric oligonucleotides display increased target specificity, while supporting RNase H mediated degradation of the target mRNA.

examine the development of antisense oligonucleotides that target human serine/threonine protein phosphatase type 5. To study the function of PP5 in living cells an RNase-H mediated approach using ‘‘second generation’’ antisense oligonucleotides was chosen for development. The chemistry was 20 -O-(2-methoxy)ethylphosphothioate ribonucleotides for the wings and phosphorothioate oligodeoxynucleotides (PS) for the core. The oligos tested were 20 bases in length and contained central phosphorothioate oligodeoxy residues (‘‘oligodeoxy gap’’) flanked by 20 -O-(2-methoxy) residues on both the 30 and 50 ends (Fig. 3). Active oligonucleotides were identified by their ability to suppress the expression of PP5 mRNA levels in A549 cells. In screening for effective antisense molecules, several oligonucleotides that were predicted to hybridize to different regions of human PP5 mRNA were synthesized. The oligonucleotides were designed to target specific regions in the protein coding region, the 50 -untranslated or the 30 -untranslated region of human PP5 mRNAs (Fig. 4A). Active oligonucleotides were rapidly identified by their ability to suppress the expression of PP5 mRNA levels using Northern blot analysis, which is an effective method to identify antisense oligonucleotides that act via an RNase-H mediated mechanism. In the screen, as well as subsequent studies, the A549 cell cultures were treated when they reached  70% confluency and the oligonucleotides were delivered in a solution of DMEM containing 15 g/ml DOTMA/DOPE (LipofectinÕ ). Careful attention should be paid to the confluency of the cell

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FIG. 4. Inhibition of PP5 mRNA and protein levels following treatment with antisense oligonucleotides. (A) PP5 mRNA levels from Northern blot analysis expressed as a percentage of the levels in control cells following normalization to GAPDH levels for each antisense oligo. Thirty-eight antisense oligos targeting different regions of PP5 were evaluated for their ability to inhibit the expression of PP5 in cultured A549 cells. The asterisk indicates that ISIS 15534 and ISIS 14504 target the same sequence. The 14504 (as well as other listed 14000 oligos) contain a

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cultures at the time of treatment because the lipid to cell membrane ratios appears to greatly affect uptake of the antisense. After  24 hr, the cells were harvested and the ability of each oligonucleotide to specifically inhibit the expression of PP5 was determined by Northern blot analysis probing for levels of PP5 mRNA. PP5 mRNA was detected using a PP5 specific cDNA probe that forms a hybrid with a single  1.5 kb transcript. This probe was generated by PCR of a full length human cDNA clone, and labeling of the 1.5 kb product was achieved with 32P using random labeling with Klenow enzyme (Decaprime kit, Ambion). A comparison graph of normalized PP5 mRNA levels in A549 cells treated with antisense oligonucleotides targeting PP5 in the presence of cationic lipids is shown in Fig. 4. The reduction in PP5 mRNA levels observed in response to treatment was varied from little or no effect on the inhibition of PP5 mRNA levels, through to pronounced effects. Several antisense oligonucleotides with activity against PP5 mRNA were identified, and ISIS 15534 which targets a region in the 30 -untranslated regions of the PP5 mRNA was chosen for further analysis (Table I). To demonstrate a concentration dependent decrease of PP5 mRNA and to assess the potency of ISIS 15534-mediated inhibition of PP5 mRNA expression, dose-response studies were conducted (Fig. 4B). As controls for nonantisense effects, several mismatch control analogues for ISIS 15534 were constructed and tested (e.g., ISIS 15521). These mismatched controls contain the same base composition as ISIS 15534. However, the sequences in mismatch controls are scrambled and noncomplementary to PP5 (Table I). After  24 hr treatment, ISIS 15534

PS-core consisting of 8 nucleosides and wings containing six 20 -O-methoxyethyl (20 -MOE) modified ribonucleotides. ISIS 15534 (and all others besides the 14,000 oligos) contains a PS-core consisting of 10 nucleosides flanked by wings with five 20 MOE-modified ribonucleosides. The relative position of the predicted hybridization sites for each of the antisense oligos evaluated is shown at the bottom. (B) Inhibition of PP5 mRNA levels by ISIS 15534. Reprinted from The Journal of Biochemistry [Z. Zuo, N. M. Dean, and R. E. Honkanen, J. Biol. Chem. 273, 12250 (1998)]. A549 cells were treated with the indicated amount of ISIS 15534 or a mismatch control analogue 15521 (see Table I for sequences). Total mRNA was prepared 24 hr later and analyzed for PP5 and GAPDH mRNA levels by Northern blot analysis. () control cells treated with liposomes without oligonucleotides. (C) Target specific inhibition of PP5 mRNA. A549 cells were treated with the indicated amount of ISIS 15534 and total mRNA was prepared 24 hr later and analyzed for PP1, PP2A, and PP4 mRNA levels by Northern blot analysis. (D) Target specific suppression of PP5 protein. A549 cells were treated with 300 nM ISIS 134217 and ISIS 134218, which target PP5, or with the indicated mismatched control analogues (ISIS 216345, 216346, 216347, and 216348; see Table I for sequences). After 24 hr, the cells were scraped in 2x sample buffer (lacking BME and dye), protein concentration measured with the DC protein Assay (BioRad), BME and dye added at the appropriate concentration and 30 ng of protein of each run on an SDS-PAGE gel and processed for Western analysis. Samples were analyzed for PP5, PP2A and PP6 protein levels using type specific polyclonal antibodies.

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TABLE I SEQUENCES siRNA duplex and sequence of RNA oligonucleotides

PP5 RNA oligo name siRNA:

PP5TG1 PP5TGMM1 PP5TG2

PP5TG3

PP5 oligo name ISIS:

pBS/U6:

50 -GGGCCCTATTGCTTGAGTGG-30 50 -GTGCGATCGTTGCGGTTAGC-30 50 -TGGCCTTCACCTCACCCTCG-30 50 -GCTCCCTCACCTCATCCGGT-30 50 -CTGGCTCCTACCTACCTCGC-30 50 -GCCAACGGGAGCCACTCGAA-30

216347 MM 216348 MM

50 -AAGCACGGGAGCCACTACCG-30 50 -GCCATCACGAGAACGGCGAA-30

30 -UTR (same region as #1 but longer) Coding region (same region as PP5TG2 but longer)

Coding region (starts approximately 39 basepairs after ISIS 134217)

Sequence of PP5 pBS/U6 inserted duplex oligonucleotides 0

Target region

0

A: 5 -GGTGAGGTGAAGGCCAAGTGA-3 30 -CCACTCCACTTCCGGTTCACTTCGA-50 B: 50 -AGCTTCACTTGGCCTTCACCTCACCCTTTTT G-30 30 -AGTGAACCGGAAGTGGAGTGGGAAAAACTTAA-50

Target region Same general region as PP5TG2 and ISIS 134217 (16 bases identical)

383

PP5#5-1 5-2 5-3 6-1

Coding region approximately 930 bp from 1st AUG in cDNA Start codon [begins at U from 1st AUG in cDNA; based on ISIS 14493 (not shown)]

Sequence of ISIS antisense (in antisense orientation)

15534 15521 MM 134217 216345 MM 216346 MM 134218

PP5 RNAi plasmid name

30 -UTR

PROTEIN PHOSPHATASE TYPE 5

PP5TGMM2

50 -CCACUCAAGCAAUAGGGCCTT TTGGUGAGUUCGUUAUCCCGG-50 50 -GUGCGAUCGUUGCGGUUAGTT TTCACGCUAGCAACGCCAAUC-50 50 -GAGGGUGAGGUGAAGGCCATT TTCUCCCACUCCACUUCCGGU-50 50 -GGUAGGACGAGAGGUAGGCTT TTCCAUCCUGCUCUCCAUCCG-50 50 -UGGCGAUGGCGGAGGGCGATT TTACCGCUACCGCCUCCCGCU-50

Target region

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produced a dose-dependent reduction of PP5 mRNA levels with an apparent IC50 of 50 nM. No effect was observed following treatment with the mismatch controls. To demonstrate equal RNA loading, the blots were probed with a glycerol-3-phosphate dehydrogenase cDNA probe (GAPDH). Studies with different human cell lines produced similar results, and Western analysis was then employed to confirm that treatment also effectively decreased PP5 protein levels (Fig. 4D). These studies revealed that protein suppression also occurred in a dose-dependent manner after  24 hr, with PP5 protein levels remaining repressed for  3 days (not shown). The specificity of the antisense oligonucleotides targeting PP5 was then examined. Because the sequence targeted by ISIS 15534 is not contained in the other PPases, ISIS 15534 would not be expected to inhibit the expression of these PPases if they indeed inhibit the expression of PP5 mRNA via an antisense mediated mechanism. To test this, Northern analysis was performed to determine if ISIS 15534 had an effect on the expression of structurally related PPases (PP1, PP2A, PP4). As seen in Fig. 4C, ISIS 15534 proved to specifically suppress the expression of PP5. The mismatch control oligonucleotide, ISIS 15521, had no effect, and similar results were also observed with two additional human cell lines (not shown). Furthermore, other selected antisense molecules, ISIS 134217 and 134218 also reduce PP5 expression levels without affecting the levels of other PPases. In contrast, their respective mismatch controls do not alter mRNA (not shown) or protein levels (Fig. 4D). Although time and labor intensive, these extensive controls are required to thoroughly demonstrate a true antisense effect, appropriate concentrations for further studies, and the usable time frame in which the oligonucleotides can be employed. Employing ISIS 15534 the suppression of PP5 expression has been shown to result in many changes including a marked increase in the ability of glucocorticoid agonist to induce GR-dependent gene expression, the nuclear accumulation of GRs in the absence of glucocorticoids, and the suppression of p53 induced p21Waf1/Cip1 expression.52–54

Double-stranded RNA Interference

Recently, double-stranded RNA interference (dsRNAi), a mechanism for the inhibition of protein expression that was formerly known to occur 52

D. A. Dean, G. Urban, I. V. Aragon, M. Swingle, B. Miller, S. Rusconi, M. Bueno, N. M. Dean, and R. E. Honkanen, BMC Cell Biol. 2, 6 (2001). 53 G. Urban, J. G. Scammell, N. M. Dean, T. K. McLean, I. Aragon, and R. E. Honkanen, Biochemistry 38, 8849 (1999). 54 Z. Zuo, N. M. Dean, and R. E. Honkanen, J. Biol. Chem. 273, 12250 (1998).

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only in nematodes and plants, has been shown to also function in Drosophila and mammalian cells.55,56 In dsRNAi an RNA molecule (whether an RNA with a certain secondary structure, or a small synthetic molecule) can under certain conditions down regulate the expression of the gene that encodes it in a posttranscriptional manner. To date the exact mechanism of how dsRNAi functions is not clear, but there are numerous articles addressing the evolutionary reasons why it occurs and probable mechanisms of action (e.g., Matzke et al.,57 Hutvagner and Zamore58). In practical use, small, synthetic RNA oligonucleotides (siRNA) can be employed to effect a reduction of a specific protein in mammalian cells. These synthetic oligonucleotides contain 19 basepairs of specific target sequence and end with two 20 -deoxy-thymidine (needed at least for stability). The oligonucleotides are then annealed and transfected into cells in culture. The exact workings of this method are being advanced daily, but with current procedures it is already a powerful technique. As with the antisense oligonucleotides, the reduction of the target protein with siRNA derives its specificity from the sequence of the mRNA. This being the case, it is obvious that methods employed above to screen, select, and evaluate antisense molecules are very similar to those needed to develop effective RNA interference experiments. To date there is little evidence for ‘‘nonspecific’’ RNAi-effects, as long as the siRNA is less than 30 basepairs in length. Nonetheless the use of dsRNA to suppress the expression of target mRNAs is still at an early stage and the possibility of unrecognized complications exist. Therefore, again careful controls are the safest way to avoid the potential artifacts produced by nontarget interactions. To identify dsRNA that would act against human PP5, we made three siRNA oligonucleotides based on the sequences of proven antisense molecules that targeted different regions of PP5 (see Table I). They encode a region in the 30 UTR (PP5TG1), a region about two-thirds of the way through the protein coding region (PP5TG2) and a region that contains the initiating codon (PP5TG3). The RNA was synthesized as single strands (purchased from a commercial supplier) and then annealed. The annealing reactions are done using each single strand at a final concentration of 20 M in RNase-free annealing buffer (1x: 100 mM potassium acetate, 30 mM HEPES-KOH at pH 7.4, 2 mM magnesium 55

S. M. Elbashir, W. Lendeckel, and T. Tuschl, Genes Devel. 15, 188 (2001). S. M. Elbashir, J. Harborth, W. Lendeckel, A. Yalcin, K. Weber, and T. Tuschl, Nature 411, 494 (2001). 57 M. Matzke, A. Matzke, and J. M. Kooter, Science 293, 1080 (2001). 58 G. Hutvagner and P. D. Zamore, Curr. Opin. Genet. Devel. 12, 225 (2002). 56

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acetate56) for 1 min at 90
C followed by 1 hr at 37
C. RNA oligonucleotide concentrations were determined by Spectroscopic Methods after resuspension in RNAse-free dH2O (note: many commercial providers have web site information to aide in exact calculation of RNA concentrations). For treatment, A549 cells were plated at 200,000 to 300,000 per dish (P60). Twenty-four hours later the annealed doublestranded RNA oligonucleotides were transfected using LipofectinÕ and employing the same methods used for the antisense oligonucleotides described above. Although a final concentration of 2.5 g/ml dsRNA was used for transfection much lower concentrations may also prove effective. Northern analysis/screening at 48 hr after transfection revealed that PP5TG2 and PP5TG3 were effective at reduction of PP5 mRNA, and could produce results similar to those achieved with ISIS 15534 (Fig. 5A). In contrast no reduction of mRNA was noted after transfection with the mismatched control siRNA. Interestingly, PP5TG1, which is based on the sequence for ISIS 15534, causes little reduction of PP5 mRNA, suggesting that the mechanism by which dsRNAi and antisense oligonucleotides act is different. Analysis of dose-response studies from samples obtained 48 hr after transfection revealed little difference in PP5 protein over the range tested (Fig. 5B) suggesting that once a minimal threshold level of dsRNA was achieved inside a cell the process that occurs is initiated to a maximal extent. siRNA-mediated reduction of PP5 protein was effective for at least 72 hr (Fig. 5C), and siRNA targeting PP5 did not produce a reduction in the expression of structurally related phosphatases [e.g., PP2A or PP6 (Fig. 5D and E)]. As observed with the antisense oligonucleotides, siRNA was effective in many other cell types including MCF-7, T47D, and human fibroblasts (data not shown). However, for each cell type the conditions used for optimal transfection (i.e., cell density, duration and amount of LipofectinÕ ) differed slightly and must be optimized with preliminary studies. Currently the synthesis of synthetic single stranded RNA is rather expensive, and the identification of siRNA that suppresses >80% of the target protein often requires the testing of several target sites. A more economical technique for identifying effective dsRNA oligonucleotides for RNA interference studies involves using plasmids such as pSUPER,59 and pBS/U6 (also known as pSILENCER from Ambion),60 in which the

59 60

T. R. Brummelkamp, R. Bernards, and R. Agami, Science 296, 550 (2002). G. Sui, C. Soohoo, E. B. Affar, F. Gay, Y. Shi, W. C. Forrester, and Y. Shi, Proc. Natl. Acad. Sci. U.S.A. 99, 5515 (2002).

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FIG. 5. Suppression of PP5 expression with small synthetic RNA (siRNA). (A) Double stranded RNA targeting PP5 (PP5TG1, PP5TG2, and PP5TG3) or respective mismatch controls (PP5TG1MM and PP5TG2MM) were transfected as described in the text into A549

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RNA oligonucleotide is coded on a plasmid and transcribed in a cell by an RNA Polymerase III promoter, such as U6 or H1. In the case of the pBS/U6 plasmid the RNA transcript is designed to encode both the sense and antisense sequences in one RNA strand with a spacer region in between. Transcription results in a hairpin structured double-stranded RNA. The termination signal for this promoter consists of five thymidines such that the actual resulting transcript in the cell is cleaved to leave a short 30 -polyU overhang similar to the siRNAs. The cell processes this RNA transcript into small double-stranded RNA, which induces RNA interference. This method was also effective at suppressing PP5 protein levels. A sequence nearly identical to siRNA PP5TG2 was cloned into pBS/U6 (see Table I, Fig. 6A) and this plasmid (pBS/U6-PP5) was transiently transfected into A549 cells as described above, except 20 g/ml LipofectAMINE (Life Technologies) was used as the transfection agent which was preincubated for 30 min with the vector before transfection. Unlike studies conducted with antisense oligonucleotides or transfection with double stranded RNA in which the suppression of PP5 protein levels can be achieved in 24–48 hr, when plasmids are used to drive siRNA a decrease in PP5 protein levels required 72 hr (Fig. 6B). Also, notable for this procedure is the need for very high transfection efficiency. Furthermore, it is more difficult to get a whole plasmid into cell compared to a short double-stranded oligonucleotide. Due to these issues, the cost advantage of making these plasmids makes it an excellent sequence screening tool to identify effective RNA sequences that can then be used experimentally in the more expensive synthetic double stranded form.

cells. After 48 hr, PP5 and GAPDH control mRNA levels were examined by Northern blot analysis. The suppression of PP5 mRNA levels obtained with ISIS 15534 (targeting PP5) and mismatch ISIS 15521 (mismatched control for ISIS 15534) are shown for comparison. (B) Western analysis of PP5 protein levels in A549 cells 48 hr after treatment with double stranded siRNA targeting PP5 or mismatched controls (PP5TG1, PP5TG2 of PP5TG1MM, PP5TG2MM, respectively). Cells were treated with 0–7.5 g of the siRNA indicated, and protein extracts were prepared 48 hr later. Western analysis was then performed with each lane containing 30 g of protein. (C) Time course analysis of PP5 protein levels. A549 cells were treated with PP5TG1, PP5TG2 of PP5TG1MM, PP5TG2MM, ISIS 15534 or ISIS 15521, and after 48 or 72 hr PP5 protein levels were examined by Western blot analysis as described above. (D) Specificity of siRNA and antisense targeting PP5. A549 cells were treated with double stranded PP5TG1, PP5TG2 of PP5TG1MM, PP5TG2MM, ISIS 15534 or ISIS 15521 as described above. After 48 hr PP5, PP2A, and PP6 protein levels were determined by Western blot analysis using type-specific polyclonal antibodies.

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FIG. 6. Suppression of PP5 protein with expression vectors producing siRNA that targets PP5. (A) Schematic of a siRNA vector targeting PP5 (pBS/U6 PP5). Double stranded oligo A (see Table I) was ligated into pBS/U6 after it was digested with ApaI, blunted by Klenow, and then digested with HindIII. This plasmid was then isolated and purified. Next, double-stranded oligo B (see Table I) was cloned into the HindIII and EcoRI site of the new plasmid (produced above). (B) Western analysis of PP5 and Actin protein levels from untreated cells (control) or after transfection with pBS/U6, or pBS/U6PP5. A549 cells were transfected with  1.6 g of the indicated plasmids with lipofectamine as described in text. Protein levels were measured  72 hr after transfection, with 30 g of protein loaded in each lane. The multiple pBS/U6 PP5 lanes represent replicate dishes.

Conclusions

When employed along with proper controls, either antisense oligonucleotides or dsRNAi can be used as a powerful tool to determine how the suppression PP5 alters the biological activity of animal cells. For both techniques target specificity is derived from the precise sequence of the target mRNA. Thus, unless the sequence of the target mRNA is identical, the oligonucleotides are species specific. For human PP5, effective target sites are provided in Table I. Unfortunately, there is no completely reliable way to predict a region in the target mRNA that will serve as a suitable

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target, so for both techniques screening is still required to identify effective oligonucleotides in other species. Acknowledgments This work was supported by grants from the National Institutes of Health (CA60750 and HL59154). We would like to thank the Yang Shi Lab for providing us with pBS/U6 as well as Dr. David Virshup and Dr. Robert Schackmann for advice and assistance with siRNA. We thank Ileana Aragon and Sylvia Mayo for technical assistance.

[28] Transgenic and Knockout Models of PP2A By JU¨RGEN GO¨TZ and ANDREAS SCHILD Introduction

Protein phosphatase 2A (PP2A) is a serine/threonine-specific protein phosphatase with important roles in cell growth, embryonic development, and human disease. All PP2A holoenzymes have in common a catalytic subunit C36 and a structural scaffolding subunit A65. These core subunits assemble with various regulatory B subunits to form heterotrimers with distinct functions in the cell. Four families of B subunits have been identified so far, termed B/PR55, B0 /PR56/PR61, B00 /PR72 and B000 /PR93/PR110.1 In contrast to Drosophila, where each subunit is encoded by a single gene, mammals express at least two A, two C, four B, at least eight B0 , four B00 , and two B000 isoforms. Thus, a total of about 75 PP2A holoenzymes can be generated. Taking the different splice variants into account, even more holoenzymes can be formed.2 This complexity, in addition to posttranslational modifications, including phosphorylation and methylation of the C subunits, provides a vast variety of possibilities for the regulation of PP2A activity.1 Despite the identification of an increasing number of in vitro substrates of PP2A, the specific PP2A holoenzymes involved in dephosphorylation are unknown for most substrates. Even more so, it is far from being understood how substrate specificity is achieved in vivo, and how the vast variety of PP2A holoenzymes is regulated. In the light of the accumulating evidence that distinct holoenzymes are involved in human disease, understanding the 1 2

V. Janssens and J. Goris, Biochem. J. 353, 417 (2001). K. Schmidt, S. Kins, A. Schild, R. M. Nitsch, B. A. Hemmings, and J. Gotz, Eur. J. Neurosci. 16, 2039 (2002).

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target, so for both techniques screening is still required to identify effective oligonucleotides in other species. Acknowledgments This work was supported by grants from the National Institutes of Health (CA60750 and HL59154). We would like to thank the Yang Shi Lab for providing us with pBS/U6 as well as Dr. David Virshup and Dr. Robert Schackmann for advice and assistance with siRNA. We thank Ileana Aragon and Sylvia Mayo for technical assistance.

[29] Transgenic and Knockout Models of PP2A By JU¨RGEN GO¨TZ and ANDREAS SCHILD Introduction

Protein phosphatase 2A (PP2A) is a serine/threonine-specific protein phosphatase with important roles in cell growth, embryonic development, and human disease. All PP2A holoenzymes have in common a catalytic subunit C36 and a structural scaffolding subunit A65. These core subunits assemble with various regulatory B subunits to form heterotrimers with distinct functions in the cell. Four families of B subunits have been identified so far, termed B/PR55, B0 /PR56/PR61, B00 /PR72 and B000 /PR93/PR110.1 In contrast to Drosophila, where each subunit is encoded by a single gene, mammals express at least two A, two C, four B, at least eight B0 , four B00 , and two B000 isoforms. Thus, a total of about 75 PP2A holoenzymes can be generated. Taking the different splice variants into account, even more holoenzymes can be formed.2 This complexity, in addition to posttranslational modifications, including phosphorylation and methylation of the C subunits, provides a vast variety of possibilities for the regulation of PP2A activity.1 Despite the identification of an increasing number of in vitro substrates of PP2A, the specific PP2A holoenzymes involved in dephosphorylation are unknown for most substrates. Even more so, it is far from being understood how substrate specificity is achieved in vivo, and how the vast variety of PP2A holoenzymes is regulated. In the light of the accumulating evidence that distinct holoenzymes are involved in human disease, understanding the 1 2

V. Janssens and J. Goris, Biochem. J. 353, 417 (2001). K. Schmidt, S. Kins, A. Schild, R. M. Nitsch, B. A. Hemmings, and J. Gotz, Eur. J. Neurosci. 16, 2039 (2002).

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function and regulation of the individual subunits may eventually help to combat human disease. The structural subunits A/PR65 and A/PR65b, for example, have been identified as tumor suppressors. Their genes are mutated in melanomas and carcinomas, as well as tumor-derived cell lines.3–5 Then, in humans with spinocerebellar ataxia, an expanded CAG repeat has been identified immediately upstream of the B/PR55b gene, indicating that dysregulation of PR55 may have functional consequences in neurodegenerative disorders.6 Finally, PP2A dysfunction has been implicated in Alzheimer’s disease (AD). The microtubule-associated protein tau is a substrate of PP2A,7 and hyperphosphorylated forms of tau form insoluble intracellular deposits in several human neurodegenerative diseases, including AD.8 In AD brain homogenates, PP2A activity is 30% reduced compared with controls,9 and in the hippocampus of AD brains, of five subunits analyzed, C, B/PR55 and B0 /PR61" mRNA expression is quantitatively decreased.10 In this chapter we review different experimental approaches in mice aimed to dissect PP2A function in vivo. The limitations of these approaches are discussed, as well as their implications for the understanding of human disease.

Classification of Transgenic Approaches

Transgenic approaches to address PP2A function in mice can be classified as follows: (1) complete or inducible versus tissue-specific knockouts of the catalytic, regulatory, or structural subunits of PP2A, (2) transgenic overexpression of regulatory subunits, and (3) dominant negative mutant approaches using either the structural subunit A or the catalytic 3

S. S. Wang, E. D. Esplin, J. L. Li, L. Huang, A. Gazdar, J. Minna, and G. A. Evans, Science 282, 284 (1998). 4 G. A. Calin, M. G. di Iasio, E. Caprini, I. Vorechovsky, P. G. Natali, G. Sozzi, C. M. Croce, G. Barbanti-Brodano, G. Russo, and M. Negrini, Oncogene 19, 1191 (2000). 5 R. Ruediger, H. T. Pham, and G. Walter, Oncogene 20, 10 (2001). 6 S. E. Holmes, E. E. O’Hearn, M. G. McInnis, D. A. Gorelick-Feldman, J. J. Kleiderlein, C. Callahan, N. G. Kwak, R. G. Ingersoll-Ashworth, M. Sherr, A. J. Sumner, A. H. Sharp, U. Ananth, W. K. Seltzer, M. A. Boss, A. M. Vieria-Saecker, J. T. Epplen, O. Riess, C. A. Ross and R. L. Margolis, Nat. Genet. 23, 391 (1999). 7 C. X. Gong, I. Grundke-Iqbal, Z. Damuni, and K. Iqbal, FEBS Lett. 341, 94 (1994). 8 J. Gotz, Brain Res. Brain Res. Rev. 35, 266 (2001). 9 C. X. Gong, S. Shaikh, J. Z. Wang, T. Zaidi, I. Grundke-Iqbal, and K. Iqbal, J. Neurochem. 65, 732 (1995). 10 V. Vogelsberg-Ragaglia, T. Schuck, J. Q. Trojanowski, and V. M. Lee, Exp. Neurol. 168, 402 (2001).

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PHENOTYPE Genotype

OF

TABLE I PP2A TRANSGENIC

Transgene expression


/


PP2A C (knockout) PP2A A/PR65 mutant
5 (transgenic) PP2A B0 /PR56 (transgenic) PP2A C mutant L199P (transgenic) PP2A C mutant L309A (transgenic)

Muscle

Lung Postnatally in neurons Postnatally in neurons

AND

KNOCKOUT MICE

Phenotype

Reference

Embryonic lethality, Gotz, 1998 no mesoderm induction and 200214,15 Form of dilated cardiomyopathy Brewis, 200027 (often premature death) Neonatal death, small lungs, absence of b-catenin Hyperphosphorylation and aggregation of tau Hyperphosphorylation and aggregation of tau

Everett, 200217 Kins, 200122 Schild, 200324

subunit C as template. All of these three approaches have been pursued (Table I). The advantages and limitations of the different approaches are discussed. Animal Models: PP2A Knockout Mice

The high degree of homology of the two catalytic subunits C and Cb in mice11 implies that deletion of one subunit (C) would be compensated by upregulation of the second subunit (Cb) resulting in a very moderate phenotype. The A and B subunits are less homologous than C, but the degree of homology is still quite high,2 meaning that targeting of these subunits should result in a more robust phenotype. PP2A C Knockout Mice

The PP2A C gene has a short promoter so that deletion of 0.5 kb immediately upstream of the transcription start site of the rat C gene abolishes expression in the chloramphenicol acetyltransferase (CAT) assay.12 We are able to confirm this in embryonic stem cells and eventually in mice by replacing more than 0.5 kb of the C promoter, exon 1, and 0.3 kb of intron 1 by a neomycin resistance cassette constitutively transcribed from a phosphoglycerate kinase promoter inserted in opposite transcriptional orientation. Electroporation of the embryonic stem cell line GS1 with a 11 12

J. Gotz and W. Kues, Biol. Chem. 380, 1117 (1999). Y. Kitagawa, H. Shima, K. Sasaki, and M. Nagao, Biochim. Biophys. Acta 1089, 339 (1991).

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targeting construct in which the neomycin cassette is flanked by 1 kb and 6 kb of C sequences, respectively, allows to isolate homologously targeted clones at a frequency of at least 3 in 100. To obtain homologous recombinants, a plate with embryonic stem cells is grown to subconfluency. The cells are washed with 10 ml CMF-PBS and incubated with 2 ml/plate of a trypsin/EDTA solution (Gibco BRL) for 5 min at 37 C. The cells are disaggregated by trituration, washed once in CMF-PBS, and resuspended at 5  106 in 0.8 ml CMF-PBS. This cell suspension is transferred to a 0.4 cm wide electroporation cuvette containing 30 g of linearized targeting construct. A single pulse of 240 V and 500 F is applied using the BioRad gene pulser. For other instruments, the optimal conditions have to be determined experimentally. The cells are transferred into a 50-ml tube containing 30 ml of embryonic stem cell medium. Cell viability is determined and the cells are distributed onto six 6-cm plates containing a layer of neomycin-resistant murine embryonic fibroblasts.13 Selection with G418 (geneticin) is initiated after 24 hr and neomycin-resistant clones become visible after 10 days. These clones are picked and screened by PCR to determine homologous recombinants. By this approach, we are able to identify several independent embryonic stem cell clones of which, in a recent experiment, two allowed germ line transmission of the mutated allele upon injection of the targeted cells into blastocysts. By this, we are able to establish two mouse strains with identical phenotypes. Analysis of both strains shows that no C RNA is transcribed from the mutant allele. C
/
embryos develop normally until postimplantation (around embryonic days 5.5–6.0). When they start degenerating, while no C protein is expressed in C
/
embryos, we find unaltered levels of total PP2A C. Degenerated embryos can be recovered even at day E13.5, indicating that the embryonic tissue is still capable of proliferating without C but that its normal differentiation is significantly impaired. Here, the primary germ layers ectoderm and endoderm are formed, but mesoderm formation is absent in degenerating C
/
embryos.14 Using a blastocyst culture system (as described below) we are able to show that during normal early embryonic development, C is predominantly present at the plasma membrane whereas the highly homologous isoform Cb is localized in the cytoplasm and nucleus. This shows us that Cb cannot compensate for vital functions of C in C
/
embryos. C is found in a 13

B. Hogan, R. Beddington, R. Constantini, and E. Lacy, ‘‘Manipulating the Mouse Embryo: A Laboratory Report,’’ Cold Spring Harbor Laboratory Press, Plainview, NY, 1994, pp. 253–290. 14 J. Gotz, A. Probst, E. Ehler, B. Hemmings, and W. Kues, Proc. Natl. Acad. Sci. U.S.A. 95, 12370 (1998).

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complex containing E-cadherin and b-catenin. In C
/
embryos, Ecadherin and b-catenin are redistributed from the plasma membrane to the cytosol, and cytosolic concentrations of b-catenin are low. Our results suggest that C is required for stabilization of E-cadherin/b-catenin complexes at the plasma membrane.15 Regulatory Subunit Knockout Mice

No reports of a PP2A regulatory subunit knockout mouse have been published so far. As the promoters are not as extensively characterized and splice variants have been identified with variable first exons more than 50 kb apart from each other,2 a targeting approach, as outlined above for C, is not feasible. We have initiated targeting of the B/PR55 subunit based on the following strategy: B/PR55 has nine coding exons.2 Exon 1 ends with the first nucleotide of an amino acid-encoding triplet, exon 2 ends with the third, exon 3 with the first and exon 4 again with the third nucleotide (Fig. 1). This has significant consequences for the design of the targeting construct. Replacing exon 2 by a neomycin resistance gene could, by readthrough transcription and splicing out exons 2 and 3, restore the open reading frame and result in a truncated protein encoded by exons 1 and 4 through 9. Similarly, replacing exon 3 could, by splicing out exons 3 and 4, result in a protein encoded by exons 1, 2, and 5 through 9. Similar splicing artifacts have been reported. Instead of a phenotype caused by the null mutation, it may in fact be caused by an aberrantly spliced protein.16 To exclude this artifact, a neomycin resistance gene (neoR) cassette is best inserted so that only the 50 portion of exon 3 and the 30 portion of exon 4 are retained (Fig. 1). As an additional measure of precaution, three stop codons, one for each reading frame, are included. The neoR gene is under the transcriptional control of the herpes simplex virus-thymidine kinase (HSVtk) promoter, inserted in an antisense orientation. As the neomycin resistance cassette is flanked by two loxP sites, it can be removed subsequently by Cre-mediated recombination (Fig. 1). Animal Models: PP2A Regulatory Subunit Transgenic Mice

Of the many regulatory subunits of PP2A, only B0 /PR56 has been expressed in transgenic mice so far. In the developing mouse embryo, PR56 is present in the lung, and protein levels reach maximal levels at embryonic 15 16

J. Gotz, A. Probst, C. Mistl, R. M. Nitsch, and E. Ehler, Mech. Dev. 93, 83 (2000). U. Muller, N. Cristina, Z. W. Li, D. P. Wolfer, H. P. Lipp, T. Rulicke, S. Brandner, A. Aguzzi, and C. Weissmann, Cell 79, 755 (1994).

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FIG. 1. Gene targeting of B/PR55 in mice. As the open reading frame is disrupted at the intron/exon boundaries, this has significant consequences for the design of the targeting construct. A simple replacement of 50 exons by a neomycin resistance cassette could result in splicing artifacts that have been reported. Instead of a null mutant, one might in fact observe a phenotype that is caused by an aberrantly spliced protein. To circumvent this, the 50 portion of exon 3 and the 30 portion of exon 4 is fused to a neomycin resistance cassette (neoR) that is under the transcriptional control of the herpes simplex virus-thymidine kinase (HSV-tk) promoter, inserted in an antisense orientation. As the neoR cassette is flanked by two loxP sites, it can be removed subsequently by Cre-mediated recombination. Three stop codons, one for each reading frame, are introduced as an additional measure of precaution.

day E17.17 To develop mice that overexpress PR56 in the lung, the full length PR56 human cDNA is best cloned into an expression vector that contains both the 3.7 kb long lung-specific human surfactant protein promoter and an SV40 small T-antigen poly A/splice cassette. Indeed, pronuclear injection of the linearized construct allows the production of transgenic mice that express PR56 specifically in lung. Mice obtained by this approach die neonatally, and lack a normal peripheral lung structure. In lung tissue of PR56 transgenic embryos, b-catenin, a major component of the wnt signaling pathway, is absent, suggesting a role of PR56 in wnt signaling during lung airway morphogenesis. Animal Models: PP2A C Dominant Negative Mutant Mice

Mutations in the catalytic subunit fall into two categories: (1) When they are introduced into the carboxy-terminus, a region known to modulate the binding of regulatory subunits, the catalytic activity is unaffected, but either no or only a subset of heterotrimers are assembled. (2) Mutations in the catalytic site of C reduce or abolish the catalytic activity without affecting holoenzyme assembly. The expression of mutant C occurs at the expense of the endogenous C as the catalytic subunit is subject to a potent autoregulatory mechanism that keeps total levels of C constant.18 We are able to express these two types of mutant forms of C in neurons of transgenic mice, as outlined below. 17

A. D. Everett, C. Kamibayashi, and D. L. Brautigan, Am. J. Physiol. Lung Cell Mol. Physiol. 282, L1266 (2002). 18 Z. Baharians and A. H. Schonthal, J. Biol. Chem. 273, 19019 (1998).

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The Dom1 Line

By mutagenesis of human C in Saccharomyces cerevisiae a number of mutants can be studied in vivo. One of them, L199P, is catalytically impaired, probably due to a disruption of metal- or substrate-binding implicated in catalytic function rather than due to a disturbed subunit interaction through misfolding.19 The cDNA of the human C mutant L199P, which we used to generate transgenic mice, is derived from the yeast plasmid YEpDE2.5.12 that carries a 978 bp HindIII/BamHI fragment encoding the HA-PP2AC-2512 mutant allele. To be able to discriminate mutant C from the highly homologous endogenous murine C, it is advantageous to epitope tag the mutant cDNA. As the highly conserved carboxy-terminus contains a leucine at position 309 that is methylated, the tag is better fused to the aminoterminus immediately downstream of the start codon. The HA (hemagglutinin)-PP2AC-2512 allele contains the t196 ! c transition mutation encoding an L199P amino acid substitution (numbering is for the untagged C).19 The 978 bp fragment is subcloned into the neuron-specific murine Thy1.2 expression vector.20,21 20 g of plasmid DNA are digested with appropriate restriction enzymes to remove the prokaryotic vector sequences. To preclude damage of the DNA by UV light, the DNA in the preparative agarose gel is not stained with ethidium bromide. Instead, the DNA is visualized by the addition of 2 g/ ml crystal violet (Sigma) to the gel, without the requirement of UV light. The DNA fragment for microinjection is cut out and purified with the PrepAGene DNA purification kit (BioRad). The DNA is further purified, to remove residual agarose, by spinning through a Millipore Ultrafree-MC 0.45 m filter unit at 5000g. The DNA is eluted with microinjection buffer (8 mM Tris pH 7.4, 0.15 mM EDTA) and adjusted to 2–3 ng/l. Before final use, the DNA is again purified using a Millipore Ultrafree-MC 0.22 m filter unit. Transgenic mice are produced by pronuclear microinjection of B6D2F1  B6D2F1 embryos cultured in Hepes-buffered M2 medium (Sigma). Founder animals are typically intercrossed with C57BL/6 mice to establish lines. Continuous backcrossing (up to 10 times) allows to obtain mice with the transgene expressed on a virtually pure C57BL/6 background that is suitable for behavioral studies. To test the role of the PP2A transgene 19

D. R. Evans, T. Myles, J. Hofsteenge, and B. A. Hemmings, J. Biol. Chem. 274, 24038 (1999). A. Luthi, H. Putten, F. M. Botteri, I. M. Mansuy, M. Meins, U. Frey, G. Sansig, C. Portet, M. Schmutz, M. Schroder, C. Nitsch, J. P. Laurent, and D. Monard, J. Neurosci. 17, 4688 (1997). 21 A. Probst, J. Gotz, K. H. Wiederhold, M. Tolnay, C. Mistl, A. L. Jaton, M. Hong, T. Ishihara, V. M. Lee, J. Q. Trojanowski, R. Jakes, R. A. Crowther, M. G. Spillantini, K. Burki, and M. Goedert, Acta Neuropathol. (Berl.) 99, 469 (2000). 20

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in lesion paradigms, it is more appropriate to backcross onto an FVB background that is susceptible to kainic acid or pilocarpine lesions. By this approach we are able to obtain transgenic mice with a chronic reduction of PP2A activity in brain. Our results also show that reduced PP2A activity is associated with altered compartmentalization, hyperphosphorylation and ubiquitination of tau, resembling a key pathological finding in AD.22

The Dom5 Line

The C-terminal leucine residue of C modulates the binding of the B/PR55 subunits through reversible methylation. In S. cerevisiae, mutation of Pph22p Leu-377, the equivalent to human C Leu-309, to alanine inhibits PP2A activity in vitro by preventing the binding of the B/PR55 regulatory subunits to the core dimer of PP2A.23 In order to produce transgenic mice that express C L309A under the control of a neuron-specific promoter, a yeast plasmid containing a 978 bp HindIII/BamHI fragment encoding the C wild-type allele is fused to a single hemagglutinin epitope located immediately downstream of the start codon. To obtain the L309A amino acid substitution in C, the CTG triplet at position 309 is changed to GCG using the QuickChange site-directed mutagenesis kit from Stratagene. The mutated fragment is subcloned into the neuron-specific murine Thy1.2 expression vector. As for Dom1, the HA-tagged cDNA under the control of the mThy1.2 promoter allows to obtain transgenic mice showing high levels of transgene expression in neurons. In these mice, similar to the Dom1 mice, tau is hyperphosphorylated and translocated to the somatodendritic domain of neurons, suggesting a role of the B/PR55 family in the tau pathogenesis of AD.24 Animal Models: PP2A PR65 Dominant Negative Mutant Mice

The scaffolding subunit A/PR65 consists of 15 nonidentical repeats. The first ten repeats bind the regulatory B subunits whereas the remaining are required for binding of the C subunit. The B subunits can only be recruited into the complex when the C subunit is already bound to A.25 In vitro and 22

S. Kins, A. Crameri, D. R. Evans, B. A. Hemmings, R. M. Nitsch, and J. Gotz, J. Biol. Chem. 276, 38193 (2001). 23 D. R. Evans and B. A. Hemmings, Genetics 156, 21 (2000). 24 A. Schild, R. M. Nitsch, and J. Go¨tz (2003), in preparation. 25 R. Ruediger, M. Hentz, J. Fait, M. Mumby, and G. Walter, J. Virol. 68, 123 (1994).

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in vivo studies demonstrated that the mutant A
5 lacking repeat 5 binds the C subunit, but not the regulatory B subunits.25,26 To generate transgenic mice that express high levels of the dominant negative mutant form of A, A
5, in heart, skeletal and smooth muscle, a nine amino acid epitope tag (EE) is introduced by PCR at the 30 end of the human A subunit-cDNA open reading frame. The cDNA is subcloned into a vector that contains the CMV enhancer, the chicken b-actin promoter, and the rabbit b-globin polyadenylation signal to yield high gene expression in muscle tissue. The DNA is purified from the gel using the DNA purification kit from Qiagen. It is concentrated with an Elutip column (Schleicher & Schuell), precipitated with ethanol and dissolved in 5 mM Tris (pH 7.5) and 0.1 mM EDTA at a concentration of 2.0 g/ml. To produce transgenic mice, pronuclei of fertilized CB6 F1 eggs are microinjected with the 3.7-kb fragment at a concentration of 2.0 g/ml. Surviving embryos are transferred into the oviducts of pseudopregnant ICR mice. By this approach it is possible to obtain A
5 transgenic mice in which the ratio of core enzyme to holoenzyme is increased in heart. At day 1 after birth, transgenic mice have an increased heart weight-to-body weight ratio that persists throughout life. End-diastolic and end-systolic dimensions are increased while fractional shortening is decreased. Finally, the thickness of the septum and of the left ventricular posterior wall is reduced.27 Protein Phosphatase Activity Measurements

To measure phosphatase activities, total brain extracts are prepared from adult mice with a teflon homogenizer (15 strokes at 100 rpm) in 1x TBS, 1% (v/v) Triton X-100, in the presence of protease inhibitors (CompleteTM with EDTA, Roche). This buffer is optimized for PP2A, to inhibit cation-dependent PP2B and PP2C activities. Homogenates are prepared in duplicate and two phosphatase assays are performed with each homogenate using the phosphatase kit V2460 from Promega. Endogenous free phosphate is removed on a Sephadex G-25 column, followed by normalization for protein content. Over a period of 10 min, release of phosphate is measured from a chemically synthesized phosphopeptide (RRA(pT)VA; pT ¼ phosphothreonine).28 This peptide is a substrate that

26

R. Ruediger, N. Brewis, K. Ohst, and G. Walter, Virology 238, 432 (1997). N. Brewis, K. Ohst, K. Fields, A. Rapacciuolo, D. Chou, C. Bloor, W. Dillmann, H. Rockman, and G. Walter, Am. J. Physiol. Heart Circ. Physiol. 279, H1307 (2000). 28 A. Donella Deana, C. H. Mac Gowan, P. Cohen, F. Marchiori, H. E. Meyer, and L. A. Pinna, Biochim. Biophys. Acta 1051, 199 (1990). 27

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can be dephosphorylated by PP2A, PP2C, and although with substantially lower efficiency, PP2B. PP1, in contrast, does not dephosphorylate peptide substrates at all. The amount of released phosphate is determined by measuring the absorbance of a molybdate : malachite green : phosphate complex at 595 nm. We are able to use this assay to compare phosphatase activities in Dom1 mice with controls. After 10 min of incubation, the assay reveals a reduction in activity of 34% (n ¼ 6, SEM ¼  3%) in Dom1 mice as compared to control homogenates.22 A similar decrease of PP2A activity is found in 5 and 12 month-old mice. To further confirm that the measured decrease in phosphatase activity is due to a reduced activity of PP2A, brain homogenates are incubated with okadaic acid (OA), a potent inhibitor of PP2A. Concentrations of 10 nM OA induce a half-maximal reduction of phosphatase activity in both wild-type and transgenic brain homogenates, normalized for the respective activities in wild-type and transgenic brain homogenates in the absence of OA. As the ratios remain the same, our data indicate that PP2A, and not another serine/ threonine-directed phosphatase, is inhibited in brains of Dom1 mice. Additional assays in the presence of 100 mM EGTA have, as compared to standard assay conditions, no influence on the ratio of inhibition, suggesting that PP2B is not a significant contaminant. Together, these data demonstrate that a neuronal, postmitotic expression of C L199P is sufficient to induce a chronic, 34% reduction of PP2A activity in brains of transgenic mice.22 While the technique described above is well suited for transgenic mice with a generally reduced PP2A activity, as in L199P mutant Dom1 mice, it bears some problems when the total PP2A activity is not affected, and when an altered substrate specificity has to be detected. This is the case for L309A mutant Dom5 mice, since the mutation prevents only the B/PR55 family of regulatory subunits from binding to the core dimer. As a consequence, we expect that in these mice other B subunits are recruited into the dimer that is present at limited amounts. By using the artificial phosphopeptide RRA(pT)VA as substrate, general phosphatase activities are likely to be unaffected. Instead, we expect that B/PR55 subunit-specific substrates, like vimentin, have to be used to monitor changes in substrate-specific PP2A activities in L309A mutant Dom5 mice. Blastocyst Cultures

PP2A is an essential enzyme as illustrated by the embryonic lethal phenotype of the PP2A C null mutant. To further assess PP2A function despite the observed early embryonic lethal phenotype, one may use

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[29]

embryonic C
/
stem cells as a source of C-deficient cell lines. This can be achieved by three means, either by starting with a heterozygous C þ /
embryonic stem cell line that is transfected with a second targeting construct carrying another selection marker (e.g., hygromycin resistance gene),29 or by increasing the neomycin concentration to obtain a C
/
cell line by gene conversation. Alternatively, one can monitor the developmental potential of C
/
embryos that are derived from C þ /
 C þ /
matings.15 To obtain homozygous C
/
blastocysts, heterozygous C þ /
female mice are hormonally stimulated with gonadotropins (Veterinaria), and crossed with heterozygous C þ /
male mice. Matings with wild-type male mice are included as controls. Blastocysts are obtained 3.5 days postcoitum by flushing the uterine horns, and are cultured on gelatin-coated plates in DMEM (Gibco BRL) supplemented with 20% fetal calf serum (Eurobio). After 2–4 days, blastocysts lose the zona pellucida, adhere to the culture dishes, and the inner cell mass of the blastocysts becomes visible on top of a layer of trophectodermal cells. Blastocyst cultures are washed in PBS, fixed with 4% formaldehyde in microtubule stabilization buffer (65 mM PIPES, 25 mM HEPES, 10 mM EGTA, 3 mM MgCl2, pH 6.9) for 3.5 hr, washed with 0.1% Triton X-100 in PBS, and incubated three times for 10 min in freshly prepared sodium borohydrate (10 mg/ml). Fixed blastocysts are either dehydrated in 100% methanol for storage at
20 C, or directly processed. For in situ immunohistochemistry, blastocyst cultures are fixed in Dent’s fixative (Methanol/DMSO ¼ 4 : 1) for 3 hr at 4 C. After blocking with 5% sheep serum (Sigma) and 1% BSA in TBS for 3 hr at room temperature, primary antisera are added in 1% BSA and 0.05% Tween-20 in TBS overnight at 4 C. Following washes in TBS/0.5% Tween-20, the secondary FITC- or Cy5-labeled antisera are added for overnight incubations at 4 C. Labeled blastocysts are washed and mounted in 0.1 M Tris–HCl pH 9.5/glycerol (3 : 7) including 50 mg/ml n-propyl gallate as anti-fading reagent. Blastocysts are viewed by confocal microscopy to determine the functional consequences of the lack of PP2A C. Advantages and Limitations of the Various Transgenic and Knockout Approaches

The advantages and inherent limitations of the different techniques (Fig. 2) aimed to dissect the function of PP2A in vivo are outlined in the following paragraph. 29

Z. W. Li, G. Stark, J. Gotz, T. Rulicke, U. Muller, and C. Weissmann, Proc. Natl. Acad. Sci. U.S.A. 93, 6158 (1996).

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FIG. 2. PP2A subunit composition and activities of the different transgenic and knockout mouse models are summarized.

Knockout approaches in general can have three potential outcomes, either (1) an embryonic or postnatal lethal phenotype (as has been shown for the PP2A C knockout14), (2) no detectable phenotype due to compensatory mechanisms, or (3) a clear but not life-threatening phenotype

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[29]

that may be either expected from known functions of the targeted gene or may not be expected at all (Table I). Lethality can be overcome by more sophisticated gene targeting approaches using either tissue-specific or inducible promoters. Redundancies, on the other hand, can be overcome by creating multiple knockouts, or by analyzing the mice even more carefully to detect subtle phenotypic alterations. Overexpression of regulatory subunits as in the case of PP2A may be complicated by the fact that total levels of PP2A are tightly regulated.18,30 As this imposes a limitation to the number of regulatory subunits that can be recruited into the functional PP2A holoenzyme complex, this implies that overexpression of one (transgenic) subunit titrates additional, endogenous regulatory subunits. Therefore, the observed phenotype may not be due to an overexpression of a distinct regulatory subunit, but may rather be caused by a lower concentration of holoenzymes containing other endogenous subunits. This requires a careful monitoring of many regulatory subunits, a task that is not easy, as it is likely that not all PP2A regulatory subunits have been identified as yet, and as specific antibodies are not available for all of them. Nonetheless, transgenic overexpression of PP2A regulatory subunits provides a rapid, first insight into tissue-specific functions of PP2A. The dominant negative mutant approach that makes use of C mutants is based on the screening of yeast mutants.19,23 In general, it is difficult to predict whether a particular dominant negative mutant phenotype in yeast may also be achieved in multiple-tissue organisms, such as mice. Additionally, in the presence of the endogenous catalytic subunit, PP2A inhibition is never 100%. This difficulty could be circumvented by crossing onto a null background, if the homozygous C mutants were viable. Nonetheless, analysis of C dominant negative mutant mice provided significant insight into the role of PP2A in the pathogenesis of tau,22 and in the regulation of ERK and JNK kinases.31 Conclusion

Considering the putative role of PP2A in the pathogenesis of human diseases, the development of more transgenic and knockout models of PP2A may provide insight into the regulation of PP2A. This may eventually lead to the discovery of therapeutic agents that can specifically counteract PP2A dysfunction. Considering the laborious task of producing and analyzing 30

S. Wera, A. Fernandez, N. J. Lamb, P. Turowski, M. Hemmings-Mieszczak, R. E. MayerJaekel, and B. A. Hemmings, J. Biol. Chem. 270, 21374 (1995). 31 S. Kins, P. Kurosinski, R. M. Nitsch, and J. Go¨tz, Am. J. Pathol., in press.

[29]

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transgenic and knockout mice, the recent advent of RNAi approaches for efficient downregulation of transcription is likely to help in the transgene design, and in the prediction of the expected phenotype.32 Acknowledgments We thank Birgit Ledermann and Jay Tracy for helpful suggestions.

32

A. M. Silverstein, C. A. Barrow, A. J. Davis, and M. C. Mumby, Proc. Natl. Acad. Sci. USA 99, 4221 (2002).

[29] Saccharomyces Gene Deletion Project: Applications and Use in the Study of Protein Kinases and Phosphatases By WAYNE A. WILSON and PETER J. ROACH

Introduction

The budding yeast, Saccharomyces cerevisiae, has a number of features which make it the ideal eukaryotic microorganism for many studies. Yeast is inexpensive to grow, nonpathogenic, has a rapid growth rate and can be handled by the same sorts of techniques as used for bacteria (basic techniques are reviewed in a previous volume in this series1). However, the prime advantage of yeast is its ease of genetic manipulation. Yeast cells can be maintained as either haploids or diploids. Mating of haploids, sporulation of the resulting diploid and analysis of the progeny is relatively straightforward, resulting in a powerful system for classical genetic analysis. Transformation with foreign DNA is both simple and efficient. Furthermore, since integration of transforming DNA occurs very efficiently by homologous recombination, this technique provides an easy means for gene replacement or disruption. In 1996, the entire yeast genome was sequenced, making this the first eukaryotic genome available. The sequencing project identified over 6000 open reading frames (ORFs). The vast majority of genes (some 96% of the total) have been individually deleted by a consortium of laboratories

1

C. Guthrie and G. R. Fink, Methods Enzymol. 194.

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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transgenic and knockout mice, the recent advent of RNAi approaches for efficient downregulation of transcription is likely to help in the transgene design, and in the prediction of the expected phenotype.32 Acknowledgments We thank Birgit Ledermann and Jay Tracy for helpful suggestions.

32

A. M. Silverstein, C. A. Barrow, A. J. Davis, and M. C. Mumby, Proc. Natl. Acad. Sci. USA 99, 4221 (2002).

[30] Saccharomyces Gene Deletion Project: Applications and Use in the Study of Protein Kinases and Phosphatases By WAYNE A. WILSON and PETER J. ROACH

Introduction

The budding yeast, Saccharomyces cerevisiae, has a number of features which make it the ideal eukaryotic microorganism for many studies. Yeast is inexpensive to grow, nonpathogenic, has a rapid growth rate and can be handled by the same sorts of techniques as used for bacteria (basic techniques are reviewed in a previous volume in this series1). However, the prime advantage of yeast is its ease of genetic manipulation. Yeast cells can be maintained as either haploids or diploids. Mating of haploids, sporulation of the resulting diploid and analysis of the progeny is relatively straightforward, resulting in a powerful system for classical genetic analysis. Transformation with foreign DNA is both simple and efficient. Furthermore, since integration of transforming DNA occurs very efficiently by homologous recombination, this technique provides an easy means for gene replacement or disruption. In 1996, the entire yeast genome was sequenced, making this the first eukaryotic genome available. The sequencing project identified over 6000 open reading frames (ORFs). The vast majority of genes (some 96% of the total) have been individually deleted by a consortium of laboratories

1

C. Guthrie and G. R. Fink, Methods Enzymol. 194.

METHODS IN ENZYMOLOGY, VOL. 366

Copyright ß 2003, Elsevier Inc. All rights reserved. 0076-6879/2003 $35.00

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in the Saccharomyces Genome Deletion project.2 A polymerase chain reaction (PCR) strategy was used for gene disruption, comprehensive details of which are available at the Saccharomyces Genome Deletion project’s website (http://www-sequence.stanford.edu/group/ yeast_deletion_project/deletion3.html). Briefly, application of the disruption protocol resulted in a start codon to stop codon replacement of each gene with a KanMX marker cassette. The PCR strategy was designed in such a way that each marker cassette was flanked by unique 20 base pair sequences, or ‘‘molecular bar codes.’’ PCR amplification of the bar codes allows unequivocal identification of a particular deletion strain. Furthermore, molecular bar codes have been utilized in a series of elegant growth rate experiments to identify genes that are important for growth under specific conditions.2,3 The deletion strains are available from a variety of sources (Table I) either as individual strains (which saves one the trouble of generating mutants in house) or as complete sets (MATa, MAT, homozygous diploid, heterozygous diploid). The particular series which we have used was obtained from Research Genetics (now part of Invitrogen) and is the first release of the MATa/MAT homozygous diploid collection. Research Genetics supplies the deletion set in a 96 well microtiter plate format, easily amenable to automation. Text files containing listings of the deletants present in each particular well on each particular plate were previously available for download from Research Genetics, but can now be obtained via the Saccharomyces Genome Deletion project’s website (http://www-sequence.stanford.edu/group/yeast_deletion_project/ ResGenftp.html). These text files can be read into a spreadsheet program for ease of manipulation.

2

E. A. Winzeler, D. D. Shoemaker, A. Astromoff, H. Liang, K. Anderson, B. Andre, R. Bangham, R. Benito, J. D. Boeke, H. Bussey, A. M. Chu, C. Connelly, K. Davis, F. Dietrich, S. W. Dow, M. El Bakkoury, F. Foury, S. H. Friend, E. Gentalen, G. Giaever, J. H. Hegemann, T. Jones, M. Laub, H. Liao, and R. W. Davis, Science 285, 901 (1999). 3 G. Giaever, A. M. Chu, L. Ni, C. Connelly, L. Riles, S. Veronneau, S. Dow, A. Lucau-Danila, K. Anderson, B. Andre, A. P. Arkin, A. Astromoff, M. El-Bakkoury, R. Bangham, R. Benito, S. Brachat, S. Campanaro, M. Curtiss, K. Davis, A. Deutschbauer, K. D. Entian, P. Flaherty, F. Foury, D. J. Garfinkel, M. Gerstein, D. Gotte, U. Guldener, J. H. Hegemann, S. Hempel, Z. Herman, D. F. Jaramillo, D. E. Kelly, S. L. Kelly, P. Kotter, D. LaBonte, D. C. Lamb, N. Lan, H. Liang, H. Liao, L. Liu, C. Luo, M. Lussier, R. Mao, P. Menard, S. L. Ooi, J. L. Revuelta, C. J. Roberts, M. Rose, P. Ross-Macdonald, B. Scherens, G. Schimmack, B. Shafer, D. D. Shoemaker, S. Sookhai-Mahadeo, R. K. Storms, J. N. Strathern, G. Valle, M. Voet, G. Volckaert, C. Y. Wang, T. R. Ward, J. Wilhelmy, E. A. Winzeler, Y. Yang, G. Yen, E. Youngman, K. Yu, H. Bussey, J. D. Boeke, M. Snyder, P. Philippsen, R. W. Davis, and M. Johnston, Nature 418, 387 (2002).

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TABLE I GENOTYPE OF PARENTAL STRAINS AND AVAILABILITY OF THE DELETION SERIES BY4741 BY4742 BY4743

MATa his3
1 leu2
0 met15
0 ura3
0 MAT his3
1 leu2
0 lys2
0 ura3
0 MATa/MAT his3
1/his3
1 leu2
0/leu2
0 met15
0/MET15 LYS2/lys2
0 ura3
0/ura3
0

American Type Culture Collection, http://www.atcc.org Open Biosystems, http://www.openbiosystems.com Research Genetics, http://www.resgen.com (In Europe, strains are available from EUROSCARF http://www.unifrankfurt.de/fb15/mikro/euroscarf/data/by.html)

Advantages and Disadvantages

Yeast have been exploited in an enormous number of different genetic screens by which strains with a phenotype of interest have been identified. The key advantage of the deletion series is that it permits a totally different type of screen to be performed. The most difficult part of a conventional genetic screen is the identification of the particular gene responsible for the observed phenotype. This information is available in advance when using an ordered array of defined yeast deletion strains. It is a simple case of recording the well number for the strain showing the phenotype of interest and referring back to the spreadsheet to determine which particular gene has been deleted in the strain of interest. A further advantage of the deletion series is that there are no biases—in essence, every gene has been deleted and a high level of saturation is ensured. Although extremely powerful, the use of the deletion series does have one major drawback. Because deletion mutants are used, only loss of function can be analyzed. This obviously means that essential genes are not represented in the series (except in the heterozygous diploid release). Equipment Required

The 96 well plate format of the deletion library would allow many procedures using this collection to be automated. However, the homozygous diploid collection from Research Genetics is supplied on just 54 microtiter plates. With such a small number, impressive results can be obtained with the minimum of equipment. Essential apparatus includes: A 12 channel automatic pipette which can deliver volumes up to 200 l. A 96 pin inoculating device, kept with the pins submerged in ethanol to maintain sterility.

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Sterile microtiter plates and lids, both standard flat bottomed plates and V-bottomed plates. Sterile adhesive foil covers for microtiter plates. It is also extremely useful to have both: A 96 well vacuum manifold. A centrifuge and rotor for centrifugation of 96 well microtiter plates. Initial Steps

Upon receiving the library, it is a good idea to make a duplicate collection in case of accidents. This is neither difficult nor time consuming. The microtiter plates are sent frozen on dry ice and should be stored at
80 C upon arrival. The wells in the plates contain 200 l of YPD medium (2% Bacto Peptone, 2% glucose, 1% yeast extract) supplemented with 200 g/ml of the antibiotic geneticin (GibcoBRL). 1.

2.

3.

4.

5.

Prepare a sufficient number of sterile 96 well microtiter plates to accommodate all of the strains in the collection by adding 200 l of YPD medium supplemented with 200 g/ml geneticin to each well. Remove one or two of the master plates from the freezer. Working next to a Bunsen burner flame (to create an updraft and minimize contamination problems) take the lid off of a master plate and swab the lid with 70% ethanol. Before the plate thaws, peel off the adhesive foil covering the wells and replace the plate lid. Use a 12 channel pipette to resuspend the yeast cells (which will mostly be on the bottom of the wells) by pipetting up and down gently. Great care must be taken to avoid splashing liquid between wells. Pass the 96 pin inoculation device through the Bunsen burner flame to remove ethanol and allow to cool briefly. Taking care not to splash, lower the 96 pin inoculation device gently in to the wells and then transfer to one of the freshly prepared plates of YPD/geneticin. Re-freeze the master plate at
80 C and, once frozen, apply a new sterile adhesive foil cover over the wells. Incubate the plates at 30 C until grown. Since the lids of the microtiter plates do not fit tightly, and since there is a small volume of liquid in each well, aeration is achieved by simple diffusion. However, if shaking is desired, microtiter plates which can hold 1 or 2 ml per well are available. Robust aeration can be obtained by inoculating 200–500 l of medium in such a plate, securing the lid with tape and shaking in a conventional shaking incubator.

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Add sterile glycerol to 15% (v/v) final concentration and freeze plates at
80 C. Once frozen, apply an adhesive foil cover over the wells.

The duplicate set of plates can be used for screening, the masters remaining in the freezer as a back up. Types of Study Possible

In broad terms, two types of study can be performed with the deletion series. First, there is a ‘genome wide’ screen whereby the entire deletion set is analyzed under particular conditions and a phenotype of interest scored. Examples of this type would include the work of Chan et al.4 who examined rapamycin sensitivity of around 2200 strains, the study from Snyder’s group5 that analyzed over 4000 strains in a study of polarized growth, and our own work investigating glycogen accumulation.6 Secondly, there is a targeted strategy, whereby subsets of the deletion collection are analyzed to answer such questions as, for example, which protein phosphatase catalytic subunits contribute to growth in high salt? As with any type of genetic screen, the only real requirement is that one has a phenotype that can be readily scored. A very good reference detailing around 100 different growth conditions that are particularly useful for genetic screens has been published and is an excellent starting point for anyone developing a screen.7 We will discuss two screens undertaken in our laboratory in order to illustrate these themes. First, we describe a genome wide screen for genes influencing glycogen storage. Second, we describe a targeted screen designed to identify the kinase and phosphatase catalytic and regulatory subunits responsible for control of glycogen synthase and phosphorylase. The simple methodology employed will be suitable for many other types of study. Genome Wide Screen for Genes Influencing Glycogen Storage

Glycogen serves as a reserve of glucose and its accumulation is initiated under conditions of nutrient limitation, such as the approach to stationary phase in liquid culture (reviewed in Ref. 8). Limitation for carbon, nitrogen, phosphorous or sulfur all act as triggers for increased glycogen synthesis.9 4

T. F. Chan, J. Carvalho, L. Riles, and X. F. Zheng, Proc. Natl. Acad. Sci. U.S.A. 97, 13227 (2000). 5 L. Ni and M. Snyder, Mol. Biol. Cell. 12, 2147 (2001). 6 W. A. Wilson, Z. Wang, and P. J. Roach, Mol. Cell. Proteomics 1, 232 (2002). 7 M. Hampsey, Yeast 13, 1099 (1997). 8 J. Francois and J. L. Parrou, FEMS Microbiol. Rev. 25, 125 (2001). 9 S. H. Lillie and J. R. Pringle, J. Bacteriol. 143, 1384 (1980).

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Our laboratory has been interested in glycogen as an example of a compound whose synthesis and utilization is under complex and intricate controls linked to the intracellular energy state as well as the nutritional status of the environment. Synthesis of glycogen requires the activities of glycogenin, a self-glucosylating initiator protein,10 glycogen synthase which catalyses bulk synthesis,11 and the branching enzyme.12 Glycogen breakdown requires glycogen phosphorylase13 and debranching enzyme14,15 or, under certain conditions, glucoamylase.16 However, the enzymes of glycogen metabolism are under a variety of transcriptional and posttranslational controls, and so genes encoding a number of other proteins and signaling pathways affect glycogen accumulation (see Ref. 8). Assessing glycogen accumulation in yeast is simple. When yeast colonies on plates are exposed to iodine vapor, they stain brown in proportion to the amount of glycogen present.17 Thus, wild type cells stain brown, glycogen hyperaccumulating mutants stain almost black and cells with low glycogen stain yellow. Using this technique, we and others had carried out a number of screens for mutants affecting glycogen storage but there was never any guarantee that our screens had been saturating and that we had identified all the genes affecting the glycogen accumulation process. A further complication is that cells which cannot utilize non-fermentable carbon sources make, but cannot maintain, glycogen stores and thus stain yellow.18 The high background of mitochondrial mutants obtained in genetic screens has made analysis of mutants which do not accumulate glycogen somewhat problematic. Since the deletion set is essentially a saturated collection of deletion mutants, and since it is ordered (and we know which mutants have respiratory defects), it is an ideal tool for this screen. Procedure

1.

10

Thaw out several plates from the deletion collection (we found that four plates at a time could be handled conveniently by one person)

C. Cheng, J. Mu, I. Farkas, D. Huang, M. G. Goebl, and P. J. Roach, Mol. Cell. Biol. 15, 6632 (1995). 11 I. Farkas, T. A. Hardy, M. G. Goebl, and P. J. Roach, J. Biol. Chem. 266, 15602 (1991). 12 D. W. Rowen, M. Meinke, and D. C. LaPorte, Mol. Cell. Biol. 12, 22 (1992). 13 P. K. Hwang and R. J. Fletterick, Nature 324, 80 (1986). 14 A. Nakayama, K. Yamamoto, and S. Tabata, Protein Expr. Purif. 19, 298 (2000). 15 M. A. Teste, B. Enjalbert, J. L. Parrou, and J. M. Francois, FEMS Micro. Lett. 193, 105 (2000). 16 I. Yamashita and S. Fukui, Mol. Cell. Biol. 5, 3069 (1985). 17 V. E. Chester, J. Gen. Microbiol. 51, 49 (1968). 18 B. Enjalbert, J. L. Parrou, O. Vincent, and J. Francois, Microbiology 146, 2685 (2000).

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409

FIG. 1. Cells were grown in 96-well microtiter plates as described. A 96-well vacuum manifold was used to filter the cells onto a nitrocellulose membrane. Glycogen accumulation was assessed by exposing the membrane to iodine vapor for 2 min. Cells staining more (ovals) or less (squares) intensely than wild type (arrow head) were scored. The empty keying well is marked with a circle.

2. 3.

4.

and transfer to fresh YPD medium in 96 well plates using a 96 pin inoculation device. Prepare duplicate plates for each incubation. A particular useful feature of the deletion series as supplied by Research Genetics is that on every plate, two wells are not inoculated with yeast but contain only growth medium (Fig. 1). These empty wells are used for keying purposes and identify both the library being used and the particular plate in the library. On the first plate in the library sequence of 54 plates, well A1 is empty whilst well A2 is empty on the second plate and so on. Furthermore, in the homozygous diploid deletion series, well H1 on each plate contains just medium. This feature makes the inclusion of internal controls straightforward and we inoculated well H1 of our experimental plates with the parental wild type strain in each case. The other empty well served both its intended keying function and as an internal check for contamination. Incubate at 30 C for 48 hr. Assess glycogen accumulation. As mentioned above, glycogen can be stained by exposing cells to iodine vapor. However, this can only be achieved using cells grown on plates or otherwise immobilized on a solid surface. Therefore, we transferred the cell suspensions to a 96 well vacuum manifold (from BioRad) and filtered the cells onto a nitrocellulose membrane. The membrane was exposed to iodine vapor for 2 min and then photographed with a digital camera (Fig. 1). As mentioned above, the screen was performed in duplicate and only wells that showed glycogen levels differing from wild type in both passes were scored. After scoring wells with high or low glycogen, the spreadsheet containing the information relating well and plate number to

410

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particular deletion strains was accessed and the mutants present in the wells of interest were identified. Using just a 12 channel pipette and a 96 well vacuum manifold, we were able to assess the glycogen accumulation phenotype of around 4600 strains in the course of a few weeks without any difficulty. Our screening methodology was validated by the fact that, in addition to many novel genes involved in the control of glycogen storage, we identified most of the genes already known to be important for this process (Tables II and III). Remarks

The most notable feature of this screen was the very large number of genes that we identified as being involved in the control of glycogen storage

BREAKDOWN

OF THE

TABLE II VARIOUS DIFFERENT GENES ISOLATED WHICH WERE FOUND TO PLAY A ROLE IN GLYCOGEN STORAGE Glycogen phenotype

Description Protein kinases Protein phosphatases (regulatory and catalytic subunits) Mitochondrial or respiratory Vesicular transport or vacuolar function Carbohydrate metabolism Amino acid metabolism Adenine metabolism Inositol metabolism Miscellaneous metabolism WD-40 repeat proteins Other signaling Ubiquitination Cytoskeleton Transport, pore proteins Sporulation Transcription, RNA processing Small ribosomal subunit Chromatin, DNA structure Known function, miscellaneous Unknown function, little or nothing known Total

Total number

Low

High

14 8

6 5

8 3

207 58 12 10 4 7 22 7 10 3 5 8 4 35 12 18 22 100 566

195 16 3 1 0 6 10 3 4 1 2 1 1 17 0 7 7 39 324

12 42 9 9 4 1 12 4 6 2 3 7 3 18 12 11 15 61 242

Adapted from W. A. Wilson, Z. Wang, and P. J. Roach, Mol. Cell. Proteomics 1, 232 (2002).

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411

YEAST DELETION SERIES

(566 or 12.4% of mutants studied). Had such a number been found in a conventional screen, we would have had to employ some secondary screening criterion to reduce the number of mutants to a level manageable for study. However, since using the deletion series is a ‘‘reverse genetic’’ approach, the identity of each mutant is known at the outset.

TABLE III KINASE and PHOSPHATASE CATALYTIC SUBUNITS, AND PHOSPHATASE REGULATORY SUBUNITS ISOLATED IN THE GENOME WIDE SCREEN Glycogen phenotype Protein kinases KIN1 GIN4 PTK2 RIM15 SNF1 CTK1 DBF2 NPR1 PSK2 TOR1 YDL025c YDR247w YNL099c YPL150w

Low Low Low Lowa Lowa High High High High Higha High High High High

Protein Phosphatase (catalytic subunits) YNL099c YCR079wb

Low High

Protein Phosphatase (regulatory subunits) GAC1 GLC8 PIG2 REG1 RTS1

Lowa Lowa Lowa Higha High

Adapted from W. A. Wilson, Z. Wang, and P. J. Roach, Mol. Cell. Proteomics 1, 232 (2002). a Known to be involved in glycogen accumulation from previous screens. b The protein encoded by YCR079w, which has similarity to type 2C protein phosphatases, was reported not to have phosphatase activity when expressed in E. coli as a GST-fusion protein [A. Cheng, K. E. Ross, P. Kaldis, and M. J. Solomon, Genes Dev. 13, 2946–2957 (1999)].

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In some cases, such as growth on different carbon sources or at different temperatures etc., it is more convenient to grow cells on solid media. Should this be the case, the 96 pin inoculating device can be used to transfer cells from microtiter plates to large diameter (15 cm) agar plates. Provided that the plates are sufficiently dry, there is little spreading of the transferred liquid.

Targeted Screens with Subsets of Strains

The ordered nature of the deletion series allows highly targeted screens to be carried out. For example, both glycogen synthase and glycogen phosphorylase are phosphoproteins and their activities are regulated by the cognate actions of specific protein kinases and phosphatases. We wanted to determine precisely which kinases and phosphatases were important for this control. By simply referring to the spreadsheet detailing strains found in each well, we constructed sub-collections that contained all kinase, all phosphatase catalytic subunit or all phosphatase targeting subunit mutants represented in the deletion series. We then measured glycogen synthase and phosphorylase activity and phosphorylation state in extracts from these strains. Although we chose to look at kinases and phosphatases, the targeted approach is obviously an ideal way to determine which members of a protein family contribute to any particular phenotype of interest.

Rationale for Glycogen Synthase Phosphatase/kinase Assay

Both glycogen synthase and glycogen phosphorylase are phosphoproteins. Phosphorylation of glycogen synthase converts the enzyme into a less active form that requires the presence of the allosteric activator glucose-6-P to elicit full activity.19 The activity of dephosphorylated glycogen synthase is essentially independent of this compound. Thus, the ratio of activity without and with glucose-6-P (
/ þ glucose-6-P activity ratio) is an index of the phosphorylation state of glycogen synthase, with high values indicating that dephosphorylated and active enzyme predominates. To assess the phosphorylation state of glycogen synthase in cell extracts (and hence the relative activity of glycogen synthase kinases and phosphatase), one needs to simply prepare cell lysates in the presence of kinase and phosphatase inhibitors and measure the
/ þ glucose-6-P activity ratio. The activity measured in the presence of glucose-6-P, referred to as the ‘‘total activity’’ is effectively a measure of the amount of glycogen synthase protein present. 19

L. B. Rothman-Denes and E. Cabib, Biochemistry 10, 1236 (1971).

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413

Therefore, measurement of total activity allows one to determine whether the expression levels of glycogen synthase vary between samples. Assay of Yeast Glycogen Synthase

The glycogen synthase assay follows the incorporation of [14C]glucose from UDP-[14C]glucose into glycogen.20 The procedure used with yeast glycogen synthase is essentially the same as that for the mammalian enzyme (discussed in DePaoli-Roach et al., this volume) but some yeast-specific details are noted below. GS homogenization buffer: 50 mM Tris–HCl pH 7.5, 100 mM NaF, 1 mM EDTA. Just prior to use, protease inhibitors (1 g/ml aprotinin, 0.5 g/ml leupeptin, 0.7 g/ml pepstatin A, 1 mM phenylmethylsulfonyl fluoride, 0.1 mM N--tosyl-L-lysine chloromethyl ketone) and reducing agent (3 mM dithiothreitol) are added. GS assay mixture without glucose-6-P: 50 mM Tris–HCl pH 7.8, 20 mM EDTA, 25 mM KF, 6.67 mM UDP-glucose, 1% (w/v) de-ionized rabbit liver glycogen. Sufficient UDP-glucose ([glucose-14C(U)]; Perkin Elmer Life Sciences) is added to give a specific activity of 200–300 dpm/nmol. GS assay mixture with glucose 6-P: As above but with glucose 6-P (added from a stock solution adjusted to pH 7.8) included at a concentration of 10.8 mM. Collection and Lysis of Cells Yeast cells are grown to late logarithmic/early stationary phase in 5 ml of the appropriate growth medium (e.g., YPD or synthetic medium) and collected by centrifugation in a bench top clinical centrifuge (1600g, 2 min, room temperature). The supernatant is decanted and the cell pellet frozen on dry ice. Ice cold GS homogenization buffer (300 l for cells grown in YPD medium or 200 l for cells grown in synthetic medium) is added, the cell pellet resuspended and transferred to a 1.5 ml microfuge tube. Acid washed glass beads (425–600 m diameter; Sigma) are added to the suspension to just below the level of the liquid and the mixture is vigorously shaken for 1 min using a vortex mixer. The cell suspension is cooled on ice for 2 min and the vortex mixing is repeated. After 5 passes of vortex mixing/cooling, the cell extract is separated from the glass beads by punching a hole in the bottom of the centrifuge tube with a syringe needle, standing the punctured tube inside another microfuge tube and then centrifuging as above. The resulting cell extract can be used directly 20

J. A. Thomas, K. K. Schlender, and J. Larner, Anal. Biochem. 25, 486 (1968).

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[30]

for glycogen synthase assay in many cases. However, certain yeast mutants have elevated levels of glucose 6-P or other metabolites which can interfere with the measurement of the
/ þ glucose 6-P activity ratio. Such metabolites can be removed by passage through a 1 ml spin column of G25 Sepharose equilibrated with homogenization buffer (see DePaoliRoach et al. this volume). Assay of Yeast Glycogen Synthase Aliquots (30 l) of undiluted yeast extract are added to 60 l of GS assay mix either with or without glucose 6-P. After 15 min incubation at 30 C, 75 l of the reaction mixture is removed, spotted to a square of 31ET chromatography paper (Whatman) and placed in ice cold 66% (v/v) ethanol. The paper squares are then washed and processed as described elsewhere in this volume (DePaoli-Roach et al.). The amount of [14C]glucose incorporated into glycogen is then determined and both the total activity (nanomole glucose incorporated per minute per milligram of protein) and the
/ þ glucose 6-P activity ratio are calculated. An example of the results obtained is shown in Fig. 2, where the parental strain and strains in which either the GIP1, GAC1 or PIG1 genes had been disrupted were analyzed. The
/ þ glucose 6-P activity ratio was similar to wild type in both the gip1::KanMX and pig1::KanMX mutants, but was substantially reduced in the gac1::KanMX mutant. Thus, the gac1 mutants are deficient in the ability to dephosphorylate and activate glycogen synthase. This is consistent with previous data demonstrating that Gac1p acts to target the Glc7p phosphatase catalytic subunit to glycogen synthase.21

Rationale for Glycogen Phosphorylase Phosphatase Assay

The situation is somewhat more complicated with yeast glycogen phosphorylase. Yeast phosphorylase is active only when phosphorylated22,23 and one cannot make a measure similar to the
/ þ glucose-6-P activity ratio. Therefore, by measuring the amount of phosphorylase activity in a sample, it is not possible to distinguish between changes in expression levels and changes in phosphorylation state. To address this problem, we devised a simple assay allowing us to quantify the ability of cell extracts to cause 21

J. M. Francois, S. Thompson-Jaeger, J. Skroch, U. Zellenka, W. Spevak, and K. Tatchell, EMBO J. 11, 87 (1992). 22 M. Fosset, L. W. Muir, L. D. Nielsen, and E. H. Fischer, Biochemistry 10, 4105 (1971). 23 K. Lerch and E. H. Fischer, Biochemistry 14, 2009 (1975).

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YEAST DELETION SERIES

415

FIG. 2. Representative data from the target screen for protein phosphatase regulatory subunits contributing to regulation of glycogen synthase. The indicated strains were grown overnight to late logarithmic/early stationary phase in YPD medium. Glycogen synthase activity was determined in the absence and presence of glucose 6-P. Data are the mean  standard error of the mean for three independent determinations, performed in duplicate.

dephosphorylation and inactivation of phosphorylase. Cell extracts were prepared in the absence of kinase and phosphatase inhibitors. The samples were split and one portion was incubated with MgATP to promote the action of endogenous phosphorylase kinase activity in the extracts whilst the other portion was incubated without MgATP to allow the action of endogenous phosphorylase phosphatase activities. At the end of the incubation period, the cell extracts were assayed for phosphorylase activity. The results are presented as the ratio of phosphorylase activity obtained without and with MgATP present in the preincubation (
/ þ MgATP activity ratio). Lower values indicate a greater dependence upon MgATP for maintenance of phosphorylase activity and hence higher phosphorylase phosphatase activity. Measurement of Endogenous Phosphorylase Phosphatase Activity in Yeast Lysates

Glycogen phosphorylase activity is measured in the reverse direction by monitoring the incorporation of [14C]glucose from [14C]glucose-1-P into

416

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glycogen. We use a modification of the methods described by Gilboe et al.24 and Hwang et al.25 Phosphorylase homogenization buffer: 50 mM Tris–HCl pH 7.5. Protease inhibitors (as for GS homogenization buffer) and reducing agent (1 mM dithiothreitol) are added just prior to use. Phosphorylase Assay Buffer: 150 mM sodium succinate pH 5.8, 150 mM glucose 1-P, 150 mM KF, 1.5% (w/v) deionized rabbit liver glycogen. [14C (U)]glucose 1-P (Perkin Elmer Life Sciences) is added to give a specific activity of 15–25 dpm/nmol. Collection and Homogenization of Yeast Yeast cells are grown and collected by centrifugation as described above for the glycogen synthase assay. The cell pellets are thawed by the addition of 200–300 l of ice cold phosphorylase homogenization buffer and lysed by vortex mixing in the presence of glass beads. The resulting cell extract is then passed through a spin column of G25 Sepharose to remove small molecules. Pre-incubation With or Without MgATP Aliquots (40 l) of undiluted yeast extract are incubated with or without the addition of 5 mM ATP and 30 mM MgCl2 in a final volume of 50 l. After 15 min incubation at 30 C, 30 l of the reaction mixture is removed and added to 60 l of phosphorylase assay buffer. After 20 min incubation at 30 C, 75 l of the reaction mixture is spotted to 31ET chromatography paper and processed as described for the glycogen synthase assay. An example of the type of data obtained is shown in Fig. 3, where the parental strain and strains in which either the GIP1, GLC8 or PIG1 genes had been disrupted were analyzed. In the parental strain, and in the gip1::KanMX and pig1::KanMX strains, the
/ þ MgATP activity ratio was low, indicating a high capacity to dephosphorylate and inactivate phosphorylase. However, the
/ þ MgATP activity ratio was increased in the glc8 mutant showing reduced phosphorylase phosphatase activity in this strain. Thus Glc8p is required for maximum phosphorylase phosphatase activity. Recently, Cannon’s group has also demonstrated a role for Glc8p in the control of phosphorylase phosphatase activity. In this case, an alternative assay was employed that measured the release of [32P] from labeled mammalian glycogen phosphorylase.26 The advantages and disadvantages of phosphatase assays based on release of [32P] from labeled substrates are discussed by DePaoli-Roach et al. (this volume). 24

D. P. Gilboe, K. L. Larson, and F. Q. Nuttall, Anal. Biochem. 47, 20 (1972). P. K. Hwang, S. Tugendreich, and R. J. Fletterick, Mol. Cell. Biol. 9, 1659 (1989). 26 S. S. Nigavekar, Y. S. Tan, and J. F. Cannon, Arch. Biochem. Biophys. 404, 71 (2002). 25

[30]

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417

FIG. 3. Representative data from the targeted screen for protein phosphatase regulatory subunits contributing to regulation of glycogen phosphorylase. The indicated strains were grown overnight to late logarithmic/early stationary phase in YPD medium. Glycogen phosphorylase assays were carried out on cell extracts that had been preincubated for 15 min either with or without the addition of Mg2 þ and ATP. Data shown are the mean  standard error of the mean for three independent determinations performed in duplicate.

Remarks

One disadvantage of these assays is that the need to homogenize samples using mechanical lysis with glass beads limits the number of samples that can be processed at one time. To address this issue, we have investigated the use of detergent lysis procedures in microtiter plates. Several vendors, including Pierce and Sigma, produce proprietary detergent mixtures for the lysis of yeast. These are marketed as being gentle enough that they do not cause extensive denaturation of proteins during the lysis procedure. We were able to assay glycogen synthase activity in cell extracts prepared using the reagent from Sigma (CelLytic-Y). Considerably less glycogen synthase activity was detected using the Pearce reagent (Y-PER) although both detergent solutions released comparable amounts of total protein.27 For the detergent lysis studies, we grew cells in microtiter plates and collected cell pellets in a centrifuge equipped for microtiter plate

27

C. Heyen and P. J. Roach, unpublished observations (2001).

418

KNOCKDOWN AND KNOCKOUT TECHNOLOGIES

[30]

centrifugation (Fisher Scientific Marathon 8K centrifuge with Microplate Carrier rotor). The use of V-bottomed plates, rather than conventional flat bottom plates, greatly facilitates removal of the supernatant following centrifugation. The cell pellets were resuspended with CelLytic-Y reagent to which protease inhibitors (see above), phosphatase inhibitor (100 mM NaF) and dithiothreitol (3 mM) had been added just prior to use. Approximately two times the pellet volume of CelLytic-Y was added to the wells and the pellet resuspended by gently pipetting up and down. After 40 min incubation at room temperature, the plates were centrifuged once again and the supernatant removed to a fresh microtiter plate. In this way, we could conveniently process 192 samples at once. Clearly the use of detergent lysis in the microplate format holds considerable promise. However, two points should be noted. First, the enzyme being studied must survive the detergent lysis process and, second, the assay must be sensitive enough such that the small amount of yeast cells available from one well of a microtiter plate provides sufficient material for measurement. Conclusion

Capitalizing on the experimental utility of yeast, investigators have for many years devised complicated screens. The same ingenuity and creativity used in these classic genetic screens can now be applied to the yeast deletion series, with all its associated advantages. Acknowledgments The work described in this chapter was supported by the National Institutes of Health Grant DK42576 (to P. J. R.), the Biomedical Research Committee of Indiana University School of Medicine (to W. A. W.), and the Indiana University Diabetes and Research Training Center (DK20542).

Author Index

Numbers in parentheses are footnote reference numbers and indicate that an author’s work is referred to although the name is not cited in the text.

A

Alexander, D. R., 203, 210, 216(8), 217(8), 219, 221, 221(8), 231 Alldridge, L., 221, 231 Allen, J. B., 176 Allen, P. B., 21, 158, 159(15), 243, 321, 322 Alms, G. R., 96, 97, 98, 145, 151(11), 152(11) Alphey, L., 364 Altmann, K. H., 376 Alto, N. M., 148, 170 Alvarez, L. J., 271 Amin, D., 8 An, H., 377 Ananth, U., 391 Andersen, H. S., 305 Andersen, J. A., 307, 317(10a,10b) Anderson, J. N., 305 Anderson, K., 404 Anderson, K. A., 176 Anderson, S. M., 126 Anderson, S. R., 4 Andracki, M. E., 376 Andrade, L. K., 376 Andre, B., 404 Ang, S. G., 18 Antonsson, B., 104 Appella, E., 338 Aragon, I. V., 373, 384 Aramburu, J., 289, 290(1) Aricescu, A. R., 224, 237 Arigoni, F., 7 Arin˜o, J., 347, 354, 358 Arisawa, M., 350, 351 Arkin, A. P., 404 Arkinstall, S., 104 Armstrong, C. A., 139 Armstrong, R. W., 275, 279(65) Arndt, K. T., 354 Arndt-Jovin, D. J., 211 Arnold, L. J., 376 Aronheim, A., 177, 178, 179(13,15), 183, 186

Aasland, R., 67 Abbott, W. M., 8, 342 Abell, C., 275, 280(66) Abo, A., 178 Abraham, R. T., 298, 304(24) Adams, J. A., 231 Adams, M. D., 103 Adams, P. A., 264 Adelstein, R. S., 123 Adler, E., 7 Aebersold, R., 54 Affar, E. B., 362, 371(15), 386 Agaisse, H., 363 Agami, R., 362, 371(11), 386 Aggen, J. B., 321 Agostinis, P., 5, 6(12,13), 9, 16(12), 189, 197(14), 198(14) Agrawal, S., 376, 378, 379 Aguzzi, A., 394 Ahmad, Z., 18 Ahn, N. G., 36 Ailles, L., 348 Aimoto, S., 265 Aitken, A., 18 Akamatsu, M., 265 Akbury, H., 66 Akerman, K., 363 Akhtar, S., 375 Akimaru, H., 363 al-Alawi, N., 177 Albar, J. P., 145, 149(10), 150(10) Alberts, A. S., 243 Aldea, M., 349, 350(8), 351, 351(7,8), 352(12), 354, 354(8,12), 357 Alemany, S., 160, 170(22), 171(22), 249 Alessi, D. R., 66, 67, 104, 145, 146(7), 158, 159(13), 164, 330, 331(11)

419

420

AUTHOR INDEX

Aschenbach, W. G., 18, 21 Asgedom, M., 262 Ashby, D. G., 202 Ashworth, A., 104, 305 Asouline, G., 275 Astromoff, A., 404 Atherton, E., 265 Attema, J., 126, 127(3) Auletta, C., 378 Ausubel, F. M., 341 Avaeva, S. M., 14, 342 Avigdor, A., 103 Avila, A., 363 Axton, J. M., 364 Ayllo´n, V., 145, 149(10), 150(10)

B Babu, K., 139 Bacon, T. A., 375 Bae, Y. S., 308 Baharians, Z., 202, 395, 402(18) Bahri, S. M., 139 Bai, G., 146, 158, 162(14), 249, 330 Baines, I. C., 308 Baker, P., 85 Balasubramanian, S., 104, 275, 280(66) Ballard, J. M., 89 Baltimore, D., 193 Banerjee, M., 178 Bangham, R., 404 Bannwarth, W., 5, 267 Banville, S. C., 264 Barany, G., 264, 266 Barban, M. A., 171 Barbanti-Brodano, G., 391 Barford, D., 3, 97, 135, 144, 152, 159, 188, 273, 306, 307, 317(10a,10b), 330, 331, 331(16), 337(16), 339 Barik, S., 152 Barker, H. M., 139, 361 Baron, U., 348 Barr, H. M., 175 Barrell, B. G., 96 Barrett, R. W., 261 Barrow, C. A., 362, 363(6), 364(6), 371(6), 403 Basilico, C., 338 Bass, M. A., 20, 146, 152(19), 156, 164, 165, 174(25)

Bastiaens, P. I. H., 213, 223 Batty, I. H., 76 Baudouin, E., 288 Bauman, P. A., 156 Baumann, G., 270 Baumgard, M. L., 43 Baumgartner, R. A., 115 Baumgartner, S., 363, 364 Baykov, A. A., 14, 342 Bayle, C., 275 Bayley, D., 379 Beattie, K. A., 145, 150 Beaven, M. A., 115, 122, 123 Beck, J. L., 301, 302(27) Beck, M., 79 Beck, T. A., 376 Beckman, S., 44 Becskei, A., 351 Beddington, R., 393 Begum, N., 123 Belisle, B., 178 Bellı´ , G., 349, 350(8), 351, 351(8), 352(12), 354(8,12) Bellosta, P., 338 Ben-Bassat, I., 103 Bender, G., 348 Bengtsson, M., 89 Benito, R., 404 Ben-Meir, D., 338, 339, 347 Bennett, C. F., 377, 378, 379 Bennett, J. M., 105 Bentin, T., 376 Berens, C., 340 Berg, R. H., 376 Berger, P., 44 Bergquist, K.-E., 268 Beri, R. K., 8, 342 Bernardi, G., 336 Bernards, R., 362, 371(11), 386 Berndt, N., 158, 174 Berry, M., 364 Bertin, S., 263 Besmond, C., 191, 192(16) Beullens, M., 145, 146, 147, 148, 148(15,22), 149, 150, 150(26), 151, 152(15), 153(22), 164, 243 Beverley, P., 221, 231 Beyer, A., 7, 9 Beythien, J., 267 Bhat, B., 379

AUTHOR INDEX

Bialojan, C., 173 Bigelow, J., 270 Bigner, S. H., 43 Bilbe, G., 363 Bilham, T., 18 Bilwes, A. M., 225 Biondi, A., 103 Birge, R. B., 276 Birrel, G. B., 285 Bisseling, T., 231 Bito, H., 304 Bittner, M. L., 104 Black, M. J., 271, 272(57) Blackburn, C., 266 Blanchetot, C., 221, 224, 226, 231(12,13), 233(12,13), 236, 236(12,13), 237, 237(13,28), 239(12), 240(13) Bleasby, A. J., 84 Blondeau, F., 44, 305 Bloom, G. S., 176 Bloomfield, C. D., 104 Bloor, C., 188, 202(9), 392(27), 398 Blumberg, P., 122 Blumberg, W. E., 206 Blumenthal, D. K., 291, 296(19) Blumer, K. J., 231 Bock, C., 20, 22(23), 32(23) Bodin, S., 44 Bodna´r, A., 214, 215(11), 221, 231 Bodnar, W. M., 275 Boeke, J. D., 176, 404 Bollen, M., 20, 135, 144, 145, 145(2), 146, 146(2), 147, 147(2), 148, 148(15,22), 149, 150, 150(26,31), 151, 151(31), 152(15,31), 153(22), 157, 164, 165, 243, 244(9), 321 Bonnard, M., 219 Bonneick, S., 44 Bonnin, A. L., 348 Borin, G., 5 Borkhardt, A., 103 Borthwick, E. B., 364, 374 Boschert, U., 104 Boschetti, M., 3 Bose, R., 354 Bose, S., 43 Boss, M. A., 391 Bossinger, C. D., 268 Botteri, F. M., 396 Boudrez, A., 149, 150, 151

421

Bourret, R. B., 58 Boutin, J. A., 263 Boutros, M., 363 Boyd, M. R., 341 Boyd, R. K., 89 Boyle, N. A., 263 Brachat, A., 350, 356 Brachat, S., 404 Brachmann, R. K., 176 Bradford, M. M., 28, 99 Brady, M. J., 20 Brand, L., 207, 231 Brander, S., 394 Brandt, H., 160 Brandt, R., 176 Brautigan, D. L., 22, 35, 38(6), 187, 243, 245, 248, 249, 249(17), 253, 253(12), 259, 259(28), 392(17), 395 Breeden, K., 18 Breit, S., 85, 90(10), 91(10) Bremnes, B., 67 Brenner, S., 275 Brent, R., 176, 341 Brew, K., 5, 330 Brewer, H. B., Jr., 23, 24(33) Brewis, N., 188, 202(9,10), 361, 364, 392(27), 398 Brey, P. T., 363 Briand, J. P., 148, 153(27) Broder, Y. C., 178, 179(15) Broeck, J. V., 363 Brown, G. E., 66 Brown, L., 361, 364, 374 Brown, N. E., 23 Brown, N. R., 3 Browne, G. J., 135, 139 Browner, M. F., 158, 162(14), 249, 330 Brummelkamp, T. R., 362, 371(11), 386 Buchardt, O., 376 Bueno, M., 384 Bujard, H., 348 Buj-Bello, A., 305 Buko, A. M., 89 Bumgarner, R., 371 Bunin, B. A., 264 Burakoff, S. J., 219 Burke, T. R., 265, 270 Burki, K., 396 Burlingame, A. L., 85, 89 Burridge, K., 308

422

AUTHOR INDEX

Bushkin, I., 314 Bussey, H., 96, 404 Butler, G., 178 Bylund, D. B., 8

C Cabib, E., 412 Calabresi, P., 336 Calderan, A., 5 Calero, F., 358 Caligiuri, M. A., 104 Calin, G. A., 391 Callahan, C., 391 Calne, D. B., 336 Calvo, S., 363 Camici, G., 307 Camici, M., 18 Campanaro, S., 404 Campbell, D. G., 67, 174 Campbell, J. M., 375 Campos, M., 145, 151(11), 152(11) Camps, M., 104 Candia, J. M., 36 Candia, O. A., 271 Cannon, J. F., 416 Cantley, L. C., 43, 65, 66, 276 Cao, T., 270 Caplen, N. J., 362 Caprini, E., 391 Capulong, Z. L., 160 Carafoli, E., 84, 88(1), 267 Carey, F., 8, 342 Cargill, J. F., 275, 279(65) Carling, D., 8, 342 Carlson, M., 96, 97, 98 Carmody, L. C., 156 Carnemolla, B., 269 Carr, D. W., 164 Carr, R., 307, 317(10a,10b) Carthew, R. W., 362 Carvalho, J., 407 Casals, N., 358 Caselli, A., 307 Casper, M. D., 376 Cassidy, R. A., 376 Catovsky, D., 105 Caudwell, F. B., 18, 20, 21, 147, 174 Caudy, A. A., 362, 371(14) Cavalieri, S., 348

Cayla, X., 145, 149(10), 150(10) Centonze, D., 336 Cepko, C., 193 Cervantes, F., 105 Cesareni, G., 270 Ceulemans, H., 20, 145, 147, 149, 150(31), 151(31), 152(31) Chabert, C., 104 Chait, B. T., 84 Chajut, A., 339 Chamberlin, R., 321 Chan, H., 379 Chan, T. F., 407 Chandhuri, M., 276 Chang, W., 265 Chao, D. M., 84 Chau, V., 17 Chawla, A., 66, 67 Chen, J., 44 Chen, M. S., 372 Chen, M. X., 147, 361 Chen, S. M., 178 Chen, W., 224 Chen, Y., 104 Chen, Y. H., 147, 361 Cheng, A., 8, 338, 339, 411 Cheng, C., 408 Cheng, N., 146, 152(19), 164, 174(25) Cheng, Q., 39 Chester, V. E., 408 Cheung, Y. W., 275, 280(66) Chia, W., 139 Chiang, M. Y., 379 Chidester, D., 379 Childs, J. E., 177 Ching, Y. P., 4, 7(9), 8(9) Chinkers, M., 372 Cho, H., 5 Chock, P. B., 17, 308 Choi, K. Y., 363 Choi, M., 298, 304(24) Chou, D., 188, 202(9), 392(27), 398 Chou, M. M., 276 Chu, A. M., 404 Chu, N. S., 336 Chu, Y., 162 Chu, Y. F., 333 Chun, K. T., 96 Chun, Y. S., 157 Chung, C. T., 191

423

AUTHOR INDEX

Cismasiu, V., 237 Clague, M. J., 44 Clarck-Lewis, I., 54 Clausen, H., 268 Clauser, K. R., 85 Clegg, R. M., 231 Clemens, J. C., 362, 363(7), 371(7) Clotet, J., 354, 358 Cloutier, J. F., 312 Cobb, M. H., 39, 41(10) Cocuzza, A. J., 379 Codd, G. A., 150 Cogan, E. B., 285 Cohen, A., 178 Cohen, P., 3, 6, 7(15), 8(15), 18, 19, 20, 21, 21(19), 23, 31, 60, 66, 135, 145, 146, 146(7), 147, 150, 151, 152, 156, 158, 159(13), 160, 164, 170(22), 171, 171(22), 174, 187, 189, 197, 197(13), 198(13,22), 243, 249, 282, 285(2), 290, 294(18), 295(18), 299(18), 300(18), 338, 339, 342, 361, 398 Cohen, P. T. W., 20, 97, 135, 138, 139, 144, 145, 145(3), 147, 152, 157, 158, 159, 159(13), 174, 187, 282, 285(2), 330, 331(11,16), 337(16), 339, 342, 361, 364, 374 Colbran, R. J., 146, 152(19), 156, 157, 159, 159(10), 160(10), 164, 165, 171, 174(10,25), 176 Colescott, R. L., 268 Collas, P., 148, 151(28), 155(28) Coller, H., 104 Collins, E., 122 Colomina, N., 357 Comer, F. I., 44 Condon, T. P., 378 Congreve, M., 307, 317(10a,10b) Connelly, C., 404 Connor, J. H., 147, 148(22), 152, 153(22), 162, 245, 249, 330, 333 Constantini, R., 393 Cook, P. D., 376, 377 Cook, P. F., 40 Cook, P. I., 268 Cooks, R. G., 89 Cooper, J. A., 271 Cooper, L. D., 20, 22(23), 32(23) Cooper, S. R., 377, 378 Copeland, T., 66 Corbett, A. H., 202

Cornelius, G., 363 Cossum, P. A., 379 Couderc, F., 275 Cox, C., 105 Coy, J. F., 44 Cozzone, A. J., 56 Crabtree, G. R., 290 Craft, C., 368 Craig, S. J., 379 Crameri, A., 392(22), 397, 399(22), 402(22) Cristina, N., 394 Croce, C. M., 391 Crooke, S. T., 377, 379 Cross, D. A., 20 Crovello, C. S., 57 Crowther, R. A., 396 Crumpton, M. J., 220 Culotta, V. C., 290, 291(15), 297(15), 298(15) Cummins, L. L., 376, 379 Cunningham, J., 20 Curran, M., 270 Currens, M. J., 341 Currie, R. A., 67 Curtiss, M., 404 Cutbertson, D. J., 20 Czech, M. P., 66, 67

D da Cruz e Silva, E. F., 157, 158, 159(15), 174, 321 da Cruz e Silva, O. B., 174 Dagle, J. M., 376 Dahan, D., 339 Dahl, N., 44 Daily, W. J., 376 Dale, R. E., 206 Damer, C. K., 85, 101, 145, 151(12) Damjanovich, S., 204, 207, 208(3,4,7) Damuni, Z., 391 Daniel, M. T., 105 D’Armiento, J., 149, 151(33) D’Arrigo, D., 67 Das, A. K., 144, 339 Davies, S. P., 339 Davis, A. J., 362, 363(6), 364(6), 371(6), 403 Davis, K., 404 Davis, P. W., 377 Davis, R. W., 96, 404 Davis, T., 43, 364

424

AUTHOR INDEX

Deak, M., 67 Dean, D. A., 384 Dean, N. M., 373, 376, 377, 378, 379, 382, 384 Deans-Zirattu, S., 158, 162(14), 249, 330 Debets, R., 210, 216(8), 217(8), 221(8) Debrunner, P. G., 301, 302(27) Decker, S. J., 273 Deisseroth, K., 304 de Jersey, J., 301, 302(27) Delibegovic, M., 135, 139 Dell, A., 89 Dellinger, D., 379 De Loof, A., 363 De Maeyer, M., 149, 150(31), 151(31), 152(31) DeMesmaeker, A., 376 den Hertog, J., 221, 224, 225, 226, 227(9), 231(12,13), 233(12,13), 236, 236(12,13), 237, 237(13,28), 239(11,12), 240(13) Dent, P., 21 Dente, L., 270 Denton, M. B., 89 Denu, J. M., 8, 9(21), 285, 307 DePaoli-Roach, A. A., 17, 18, 20, 20(3), 21, 22, 22(23), 31, 32(23), 152, 333 Derua, R., 189, 197(14), 198(14) Desai, D. M., 225, 230 Deuel, T. F., 224 Deutschbauer, A., 404 DeVine, R. J., 376 Diaz-Meco, M. T., 363 Dibo, G., 262 Di Como, C. J., 354 Dietrich, A. D., 19 Dietrich, F., 404 di Iasio, M. G., 391 Dillard, L. W., 275 Dillmann, W., 188, 202(9), 392(27), 398 Diltz, C. D., 307 Dimitrov, T., 224 Dinischiotu, A., 151 Disney, M. D., 376 Divinski, I., 339 Dixon, J. E., 43, 44, 47(5,13), 49, 51(19), 53, 271, 273, 305, 362, 363(7), 371(7) Dobbie, L., 139 Dobrowolska, G., 31 Dobrusin, E. M., 273 Doe, S., 221, 231

Doering, R., 266 Dombradi, V., 160 Donella-Deana, A., 3, 4(3), 5, 5(3), 6, 7, 7(15), 8(15), 9, 285, 398 Donner, D. B., 44 Donoghue, D. J., 372 Dornan, S., 221, 231 Dove, S. K., 44 Dow, S. W., 404 Dowd, S., 104 Dower, W. J., 261 Dowler, S., 67 Downes, C. P., 20, 44, 64, 65, 66, 67, 76, 77, 79, 84 Downing, J. R., 104 Drake, P. G., 305 Drexler, H. G., 103 Dror, R., 314 Duclos, B., 56 Dudai, Y., 364 Dufresne, S. D., 18 Dujon, B., 96 Dunican, L. K., 270

E Ebner, F. F., 157, 159, 159(10), 160(10), 174(10), 176 Edidin, M., 204, 208(2), 218 Edington, K., 363 Edman, P., 86 Edwards, M. C., 176 Egholm, M., 376 Egloff, M.-P., 97, 135, 144, 152, 159, 321, 330, 331(16), 337(16) Ehler, E., 392(14,15), 393, 394, 400(15) Eisinger, J., 206 Ek, P., 56, 62(5,6) Ekstrom, S., 89 El-Bakkoury, M., 404 Elbashir, S. M., 362, 371(13), 385 Elledge, S. J., 138, 145, 176, 230 Elliott, E., 245, 249(17) Ellman, J. A., 261 Ellmann, J. A., 264 Elson, A., 124, 125, 126(2) Embi, N., 20 Emr, S. D., 66 Endo, S., 162, 249, 331 Eng, J. K., 85

AUTHOR INDEX

Engel, M., 363 Engelberg, D., 177 Enjalbert, B., 408 Enomoto, S., 372 Ens, W., 89 Entian, K. D., 404 Epplen, J. T., 391 Erdo¨di, F., 152 Eriksson, C., 340 Eriksson, E., 340 Espanel, X., 271, 276(55) Esplin, E. D., 391 Estes, R., 368 Eto, M., 243, 244, 245, 245(11), 248, 248(11), 249, 249(11,17), 253(12), 257, 259 Etter, E. F., 245 Evagelidis, A., 339 Evangelopoulos, A. E., 14 Evans, D. R., 202, 392(22), 396, 397, 399(22), 402(19,22,23) Evans, G. A., 391 Evens, K., 151 Everett, A. D., 392(17), 395 Evtushenko, O. A., 14, 342

F Fabiato, A., 297(29), 304 Fabiato, F., 297(29), 304 Fadden, P., 145, 151(11), 152(11) Faier, S., 339 Fait, J., 188, 397, 398(25) Fajardo, J. E., 276 Fakata, K. L., 298 Fan, S., 338 Farkas, I., 408 Farmer, J. D., 290 Fauchere, J. L., 263 Feiken, E., 237 Felberg, J., 237 Feldmann, H., 96 Fellner, T., 187, 195 Fenn, J. B., 89 Fenyo, D., 84 Feramisco, J. R., 13, 243 Ferguson, K. M., 66, 67 Fernandez, A., 402 Ferna´ndez, R., 145, 149(10), 150(10) Ferrans, V. J., 314 Ferrell, J. E., 271

Ferrell, J. E., Jr., 246 Ferrigno, P., 197, 198(22) Field, S. J., 66 Fields, G. B., 264, 265, 268 Fields, K., 188, 392(27), 398 Fields, S., 176 Fienberg, A. A., 21 Figliozzi, G. M., 264 Finbloom, D. S., 363 Fink, G. R., 403 Finkel, T., 306, 314 Fiol, C. J., 31 Fire, A., 362 Fiscella, M., 338 Fischer, E. H., 5, 307, 414 Fitch, W. L., 275 Fjeld, C. C., 8, 9(21), 285 Fladd, C., 237 Flaherty, P., 404 Flandrin, G., 105 Fleenor, J., 362 Fleming, I. N., 84 Fletterick, R. J., 408, 416 Flint, A. J., 273 Flournoy, S., 377 Fodor, S. P. A., 261 Ford, H., 265 Forney, B., 162, 249 Forrester, W. C., 362, 371(15), 386 Fosset, M., 414 Fotsis, T., 85, 90(10), 91(10) Foulkes, J. G., 171, 249 Foury, F., 404 Fox, B. A., 220 Fox, C. A., 157 Francois, J., 407, 408, 408(8), 414 Frank, R., 263, 266, 266(12) Fransen, J. A., 126, 127(3) Franza, B. R., 230 Fraser, I. D., 148, 170 Frederick, D., 152 Freier, S. M., 376, 377, 378, 379(40) Freire, E., 218 Freitas, M., 271 Freundlieb, S., 348 Frey, U., 396 Friend, S. H., 404 Fritsch, A., 178 Fruman, D. A., 65 Frye, C., 43

425

426

AUTHOR INDEX

Fu, H., 271 Fujisawa, K., 363 Fukuda, N., 376 Fukuda, T., 308, 314(16), 315(16), 316(16) Fukui, S., 408 Fukumoto, T., 363 Fukunaga, K., 339 Fukunaga, R., 363 Fulga, T. A., 237 Furie, B., 57 Furie, B. C., 57 Furka, A., 262 Furuya, K., 244, 245(11), 248(11), 249(11)

G Gaasenbeek, M., 104 Gadella, T. W. J., 226, 231, 231(12), 233(12), 236(12), 239(12) Gadella, T. W. J., Jr., 221 Gaffney, P. R. J., 66 Galbraith, W. M., 378, 379 Galibert, F., 96 Gallagher, T., 151, 164 Gallego, C., 357 Gallop, M. A., 261 Galton, D. A., 105 Gamble, J., 221, 231 Ganapathi, M. K., 160 Gantalen, E., 404 Garcı´ a, A., 145, 149(10), 150(10) Garcia, P., 67, 72 Garfinkel, D. J., 404 Garı´ , E., 349, 350(8), 351, 351(7,8), 352(12), 354, 354(8,12), 357 Gaskell, S. J., 89 Gaullier, J. M., 67 Gay, F., 362, 371(15), 386 Gazdar, A., 391 Geary, R. S., 378, 379(40) Gebbink, M. F., 237 Gehring, W., 364 Geiss, G., 371 Geitz, D., 102 Geladopoulos, T. P., 14 Genoux, D., 243 Gergely, P., 152 Gerstein, M., 404 Gery, S., 339 Gesson, I., 263

Geuna, M., 348 Geysen, H. M., 262, 275 Ghosh, M. C., 289, 290, 298(16), 300, 301(26), 302(16), 303(16) Giaever, G., 404 Gibson, B. W., 18 Giclas, P. C., 378 Giddings, J., 379 Gietz, R. D., 353, 355 Gilboe, D. P., 416 Gillieron, C., 104 Giordani, A. B., 89 Giordano, A., 230 Giovanella, B. C., 43 Gish, G., 276 Glass, D. B, 13 Glinsmann, W. H., 243 Glish, G. L., 89 Glover, D. M., 363, 364 Goebl, M. G., 96, 408 Goedert, M., 396 Goff, L. K., 221, 231 Goffeau, A., 96 Golbeck, J., 290, 302(14), 303(14) Goldberg, J., 151, 159, 322, 330, 330(6), 331(6), 333(6), 337(15) Golden, T. A., 372, 373 Goldman, S. J., 219 Goldsmith, R., 264 Goldstein, B. J., 308 Golub, T. R., 104 Gomes, R., 364 Gong, C. X., 391 Gonnet, G., 84, 88(1) Gonzalez, C., 376 Goody, R. S., 237 Goodyear, L. J., 18, 21 Gordon, E. M., 261 Gordon, J. A., 149 Gorelick-Feldman, D. A., 391 Gorey-Feret, L. J., 379 Goris, J., 5, 6(12,13), 9, 16(12), 145, 187, 189, 197(14), 198(14), 390 Gossen, M., 348 Gotte, D., 404 Go¨tz, J., 390, 391, 392, 392(2,14,15,22,24), 393, 394, 396, 397, 399(22), 400, 400(15), 402, 402(22) Gralnick, H. R., 105 Gram, H., 270

AUTHOR INDEX

Gratama, J. W., 210, 216(8), 217(8), 221(8) Graves, P. R., 96 Gray, A., 64, 76, 77, 79 Gray, J. D., 66 Green, S. R., 339 Greengard, P., 21, 148, 151, 157, 158, 159, 159(15), 243, 321, 322, 330(6), 331(6), 333(6) Greenstein, D., 191, 192(16) Gresser, M. J., 280 Griesbeck, O., 235 Griffey, R. H., 376 Griffith, J. P., 290 Griffith, O. H., 285 Grill, E., 283 Groenen, P., 149 Groom, L. A., 104 Gross, M., 149 Groves, M. R., 188 Grundke-Iqbal, I., 391 Gruppuso, P. A., 248 Gryaznov, S. M., 376 Gschmeissner, S., 67 Gubellini, P., 336 Gudlener, U., 404 Guerrero, F., 363 Guilherme, A., 66, 67 Guo, J., 371 Gupta, R., 67, 72 Gustafson, E., 157 Guthridge, M. A., 338 Guthrie, C., 403 Guy, C. A., 268

H Haak, H. L., 105 Haddy, A., 290, 303(9) Haditsch, U., 243 Haj, F. G., 223 Halfter, W., 224 Hall, W. W., 84 Hallenbeck, J. M., 245 Hamma, T., 376 Hampsey, M., 407 Han, S. J., 363 Hanada, M., 338 Hanafusa, B., 276 Hancock, J. F., 177 Hanke, C., 7, 9

427

Hanlon, N., 3, 188 Hanneman, J., 8 Hannigan, G. E., 338 Hannon, G. J., 203, 362, 371(14) Hanrahan, J. W., 339 Harborth, J., 362, 371(13), 385 Harder, K. W., 54 Hardie, D. G., 4, 7(9), 8(9), 100, 339 Hardy, T. A., 408 Harlow, E., 139, 195 Harper, J. W., 230 Hartl, F. T., 40 Hartshorne, D. J., 145, 152 Hartson, S. D., 372 Hasan, M. T., 348 Haser, D. M., 276 Hashimoto, Y., 297 Hastewell, J. G., 379 Hastie, C. J., 361 Hatano, Y., 157 Hattier, T., 43 Hauer, C. R., 89 Haystead, T. A., 84, 85, 93(12), 95, 96, 97, 98, 100, 101, 145, 151(11,12), 152(11) Hazeki, O., 51 He, B., 149 He, K., 376 Heery, D. M., 270 Heesen, S., 356 Heffetz, D., 314 Hegemann, J. H., 404 Heim, R., 231, 235 Heitman, J., 289 Helminen, P., 103 Helps, N. R., 138, 139, 145, 152, 339, 364 Hemenway, C. S., 289 Hemmings, B. A., 4, 114, 160, 170(22), 171(22), 188, 202, 249, 362, 363, 363(7), 364, 371(7), 390, 392(2,14,22), 393, 396, 397, 399(22), 402, 402(19,22,23) Hemmings, H. C., 243 Hemmings-Mieszczak, M., 402 Hempel, S., 404 Henderson, P. J., 249 Hendrich, M. P., 301, 302(27) Hendriks, W. J., 126, 127(3) Henlin, J. M., 263 Hennighausen, L., 126, 127(3) Henry, S. P., 377, 378

428

AUTHOR INDEX

Hentz, M., 188, 397, 398(25) Herberg, F. W., 152 Herman, Z., 404 Hermans, J., 105 Hermesmeier, J., 56, 283, 286(10) Herrero, E., 347, 349, 350(8), 351, 351(7,8), 352(12), 354(8,12), 357 Hersh, E. M., 262 Hess, J. F., 58 Heyen, C., 417 Hibshoosh, H., 43 Hiken, J. F., 18, 20(3) Hillen, W., 340, 348 Hillenkamp, F., 89 Hinks, J. A., 307, 317(10a,10b) Hiraga, A., 145, 338 Hirano, K., 145 Hirshman, M. F., 18, 21 Hirt, H., 288 Hisahara, S., 363 Hishiya, A., 339 Hobbs, F. W., 379 Hobson, W. C., 378 Hofer, H. W., 290, 291(17), 297(17), 298(17), 302(17) Hoffmann, I., 305 Hofsteenge, J., 396, 402(19) Hogan, B., 393 Hogrefe, R. I., 376 Hoheisel, J. D., 96 Hojrup, P., 84 Holder, J. C., 20 Holik, J. J., 66, 67 Holmes, A. B., 66 Holmes, B., 221 Holmes, C. F., 18, 171, 361 Holmes, N., 231 Holmes, S. E., 391 Holst, J., 113, 114, 116(3), 117(3), 119(3), 121(3), 122(4), 123(4) Hong, M., 396 Honkanen, R. E., 152, 361, 372, 373, 382, 384 Hooft van Huijsduijnen, R., 271, 276(55) Horiuchi, A., 148, 158, 159(15), 321, 322, 330(6), 331(6), 333(6) Horva´th, G., 210, 216(8), 217(8), 221(8) Hou, D. X., 363 Houghten, R. A., 264 Houthaeve, T., 85, 90(10), 91(10)

Hoving, S., 267 Howard, E. F., 146, 152(19), 164, 174(25) Howard, R. B., 377 Hruby, V. J., 262 Hsieh-Wilson, L., 158, 159(15), 322 Hu, L. J., 44 Hu, R., 43 Hu, W. Y., 376 Huang, C. C., 336 Huang, C. Y., 290, 299(8) Huang, D., 96, 408 Huang, F. L., 243 Huang, H.-B., 148, 151, 152, 158, 159, 159(15), 321, 322, 330(6), 331(6), 332(7), 333(6,7) Huang, J., 57 Huang, L., 391 Huang, T. S., 8 Huang, X., 361 Huard, C., 104 Hubbard, M. J., 3, 20, 21(19), 146 Hudziak, R. M., 376 Huguenin-Reggiani, M., 271, 276(55) Hunt, D. F., 89 Hunter, C. P., 362 Hunter, T., 221, 225, 226, 227(9), 231(12), 233(12), 236(12), 239(11,12), 271 Hutvagner, G., 385 Huyer, G., 280 Hwang, P. K., 408, 416

I Ichijo, H., 372 Ikebe, M., 145, 146(7), 164, 257 Ikeda, S., 338 Ikura, M., 231 Imani, F., 362 Immanuel, D., 354 Ingebritsen, T. S., 162, 243, 249, 282, 339 Ingersoll-Ashworth, R. G., 391 Iqbal, K., 391 Irani, K., 314 Irino, S., 157 Ishihara, T., 396 Ishii, S., 363 Ishimoto, A., 363 Itaya, K., 14 Ito, M., 244 Itoh, M., 5

AUTHOR INDEX

Itoh, T., 66, 67 Ittmann, M., 43 Ivanetich, K. M., 263 Iversen, L. F., 305 Iversen, P. L., 376 Izuta, M., 351

J Jacq, C., 96 Jaeger, J. A., 376 Jagiello, I., 148, 149, 150, 150(26), 164 Jakes, R., 396 James, P., 84, 88(1) James, S. R., 66 Janda, K. D., 263, 275 Jansen, P. G., 305 Janssen, J. W., 103 Janssens, V., 145, 187, 390 Jaramillo, D. F., 404 Jasser, S. A., 43 Jaton, A. L., 396 Jensen, O. N., 84 Jhoti, H., 307, 317(10a,10b) Jia, Z., 280, 306 Jiang, G., 221, 225, 226, 227(9), 239(11) Jie Yang, J., 20, 22(23), 32(23) Jin, G., 371 Jing, S. L., 18 Jirik, F. R., 54 Johns, R. B., 267 Johnson, D. F., 97, 135, 147, 152 Johnson, L. N., 3 Johnson, P., 237 Johnston, M., 96, 404 Jones, M. E., 271, 272(57) Jones, T., 404 Jorcano, J. L., 178 Jovin, T. M., 211, 213 Juhl, H., 20 Jular, G., 67 Juliano, R. L., 375 Julius, M., 219 Jung, G., 262

K Kaiser, Kaiser, Kaiser, Kaldis,

E., 268 K., 364 R. E., 89 P., 8, 338, 339, 411

429

Kamada, K., 350 Kamibayashi, C., 176, 368, 392(17), 395 Kamps, M. P., 56 Kanai, F., 66 Kanamaru, R., 338 Kang, S. W., 308 Kanmatsuse, K., 376 Kansas, G. S., 105 Kanuka, H., 363 Karas, M., 89 Karby, S., 339 Karginov, A., 249 Karin, M., 177 Kariv-Inbal, Z., 339, 347 Karsh-Mizrachi, I., 363 Kass, R. S., 149, 151(33) Katada, T., 51 Kataoka, T., 377 Katoh, H., 372 Katsura, K., 338 Katz, S., 178, 179(15) Katze, M. G., 371 Kaufman, R. J., 339 Kawabe, T., 149 Kawasaki, A. M., 376 Kazanietz, M. G., 122 Kazmierski, W. M., 262 Kellner, R., 7, 9 Kelly, D. E., 404 Kelly, J., 280 Kelly, S. L., 404 Kemp, B. E., 13 Kempe, M., 266 Keng, Y.-F., 5, 16, 278, 279 Kennedy, A., 20, 22(23), 32(23) Kenworthy, A. K., 218 Keppens, S., 139, 150 Kerc, E., 283 Kerkhofs, H., 105 Keyse, S. M., 104 Khan, J., 104 Khangulov, S. V., 290 Kiener, P. A., 219 Kigawa, T., 66, 67 Kihlberg, J., 268 Kikuchi, K., 244 Kikuchu, A., 66, 67 Killea, S. D., 39 Kim, J.-H., 17, 18, 20, 21, 22(23), 32(23) Kim, J. L., 290

430

AUTHOR INDEX

Kim, S.-A., 44 Kim, S. R., 307 Kim, Y., 57 Kinder, D. S., 275 King, C. R., 270 King, F., 276 King, M. M., 290, 299(8) Kingston, R. E., 341 Kini, S., 157, 159(10), 160(10), 174(10) Kins, S., 390, 392(2,22), 397, 399(22), 402, 402(22) Kioschis, P., 44 Kishioka, H., 376 Kiss, E., 152 Kissinger, C. R., 290 Kitada, K., 350, 351 Kitagawa, Y., 157, 392 Kitas, E. A., 5, 267 Kitazawa, T., 245, 253(12), 257 Klabunde, T., 290 Klarlund, J., 66, 67 Klauck, S. M., 44 Klee, C. B., 6, 7, 289, 290, 290(1), 291(15), 294(18), 295(18), 296, 297(15,21), 298, 298(15,16,21), 299(18,21), 300, 300(18), 301(26), 302(16), 303(16,21) Kleeman, T., 152 Kleiderlein, J. J., 391 Kleinberger, T., 175, 176 Klem, R. E., 376 Klumpp, S., 7, 9, 56, 150, 282, 283, 284, 286(10) Knapp, R. J., 262 Knobloch, M., 243 Knowles, S. K., 376 Knuutila, S., 103 Kobayashi, R., 275 Kobayashi, T., 4, 7(9), 8(9), 283, 338 Koff, A., 230 Kole, H. K., 265 Kole, R., 375 Komaki, K., 338 Kondo, N. S., 376 Kooter, J. M., 385 Kornbrust, D. J., 378 Korsmeyer, S. J., 149 Koshiba, S., 66, 67 Kotter, P., 404 Kowluru, A., 56 Krachnak, V., 262

Krag, D., 270 Krainer, A. R., 149 Kraus, M. H., 363 Krebs, B., 290 Krebs, E. G., 31 Kretz, C., 44 Krieg, A. M., 377 Krieglstein, J., 56, 282, 284, 285(12) Krinks, M. H., 6, 7, 290, 294(18), 295(18), 299(18), 300(18) Krishnaraj, R., 5 Krueger, N. X., 14 Krutchinsky, A. N., 89 Kues, W., 392, 392(14), 393 Kular, G., 84 Kuramoto, E., 377 Kuret, J., 176 Kuriyan, J., 151, 159 Kurokawa, J., 149, 151(33) Kurosinski, P., 402 Kwak, N. G., 391 Kwon, K. S., 307 Kwon, Y.-G., 151, 159, 321, 322

L LaBonte, D., 404 Lacy, E., 393 Laemmli, U. K., 28 Lam, K. S., 262 Lamb, D. C., 404 Lamb, N. J., 402 Lambert, P. H., 263 Lamond, A. I., 146, 152(18), 222, 223(27) Lamont, D. J., 145 Lan, N., 404 Land, H., 193 Lander, E. S., 104 Lane, D., 139, 150, 195 Lanfrancone, L., 270 Langan, T. A., 189, 197, 197(13), 198(13,22) Langeberg, L. K., 148, 170 Langford, L. A., 43 Lanner, C., 20, 21, 22(23), 32(23) Lanzetta, P. A., 271 LaPorte, D. C., 408 Laporte, J., 44, 305 Larner, A. C., 363 Larner, J., 23, 24(33), 27, 29, 413

431

AUTHOR INDEX

Larson, H. J., 376 Larson, K. L., 416 Laub, M., 404 Lauer-Fields, J., 264 Laurell, T., 89 Laurent, J. P., 396 Lavi, S., 338, 339, 347 Lavoinne, A., 21 Lawrence, D. S., 5, 16, 18, 20(3), 35, 36(4), 278, 279 Lawrence, J. C., Jr., 18, 20, 21, 22(23), 32(23) Lazarovits, A. I., 220 Leach, C., 243, 253, 259, 259(28) Lebl, M., 262 Lechleider, R. J., 276 Leder, P., 125 Lee, E. Y., 5, 146, 157, 158, 160, 162, 162(14), 244, 249, 330, 331, 333 Lee, G., 176 Lee, J. I., 8 Lee, J. S., 363 Lee, S. R., 307 Lee, V. M., 391, 396 Lee, W. J., 363 Leeds, J., 378 Leeksma, C. H., 105 Le Feuvre, C. E., 220 Lehto, M. T., 222 Lemmon, M. A., 66, 67, 225, 227 Lendeckel, W., 362, 371(13), 385 Lerch, K., 414 Leslie, N. R., 44, 65 Lesnik, E. A., 376, 378, 379(40) Letsou, A., 363 Leung-Hagesteijn, C., 338 Levin, A. A., 376, 377, 378, 379(40) Levine, T. P., 80 Levy-Nissenbaum, O., 103 Ley, S. C., 220 Li, D., 31 Li, H.-C., 148 Li, J., 43 Li, J. L., 391 Li, L., 118 Li, N., 177 Li, S., 245, 289 Li, S. P., 300, 301(26) Li, X., 363 Li, Y.-C., 280 Li, Z. W., 394, 400

Liang, H., 280, 404 Liao, H., 404 Liaw, D., 43 Lickteig, R. L., 368 Ligon, A. H., 43 Lillie, S. H., 407 Lim, H. Y., 243 Lin, H., 43 Lin, J. W., 148, 170 Lineruth, K., 364 Lio, A., 275, 279(65) Lipp, H. P., 394 Liu, C., 290 Liu, H., 66 Liu, J., 290 Liu, L., 404 Liu, Q. R., 245 Liu, R.-Q., 264 Llopis, J., 231 Lloyd, H. G. E., 118 Loboda, A. V., 89 Lockett, S. J., 214, 215(11) Loew, R., 348 Loh, M. L., 104 Long, F. X., 331 Louis, E. J., 96 Lu, C. S., 336 Lu, M., 340 Lucau-Danila, A., 404 Lucocq, J. M., 64, 80, 81, 81(18), 84 Ludowyke, R. I., 113, 114, 116(3), 117(3), 118, 119(3), 121(3), 122(4), 123, 123(4) Luhrmann, R., 362 Lui, X., 275 Lung, F.-D., 270 Luo, C., 404 Luo, J., 339 Luo, X., 139 Lupo, A., 339 Lussier, M., 404 Luthi, A., 396

M Maaßen, A., 282 MacDougall, L. K., 145, 146(7), 164 MacGowan, C. H., 6, 7(15), 8(15), 398 Mackey, A. J., 85, 93(12), 101 MacKintosh, C., 57, 141, 150, 151, 164, 331 MacKintosh, R. W., 164

432

AUTHOR INDEX

MacLean, D., 271, 273 MacMillan, L. B., 146, 152(19), 164, 174(25) Maeda, T., 339 Maehama, T., 43, 44, 47(5,13), 53, 362, 363(7), 371(7) Magee, A. I., 177 Mahadev, K., 308 Mahendra, A., 338 Mahuren, J. D., 285 Maier, M. A., 377 Maizel, A. L., 219, 226 Majeti, R., 225 Mak, T., 44 Malencik, D. A., 4 Manalan, A. S., 290, 294(18), 295(18), 299(18), 300(18) Manao, G., 307 Mandel, J. L., 44, 305 Mandiyan, S., 280 Mann, M., 84, 85, 89, 90(10,18), 91(10) Manning, M. C., 36 Manoharan, M., 376, 377, 378, 379, 379(40) Mansour, S. J., 36 Mansuy, I. M., 243, 396 Mao, R., 404 Marcandier, S., 56 Marchiori, F., 5, 6, 6(12,13), 7(15), 8(15), 9, 16(12), 398 Marchler, G., 363 Marcusson, E. G., 379 Margolis, R. L., 391 Marin, O., 5, 14 Markham, P. M., 379 Marko-Varga, F., 89 Marks, A. R., 149, 151(33) Markworth, C. J., 275 Marley, A. E., 8, 342 Maroun, C. R., 219 Maroun, M., 186 Marshall, C. J., 177 Martı´ ez-A., C., 145, 149(10), 150(10) Martinez, J., 362 Martinou, I., 104 Martins, S. B., 148, 151(28), 155(28) Marx, S. O., 149, 151(33) Marzocchini, R., 307 Masaracchia, R. A., 13 Mason, T. J., 262 Matko´, J., 204, 208(2) Matsumoto, K., 338

Matsuo, T., 66 Matsuura, H., 372 Matsuura, J. E., 36 Matthews, H. R., 57 Matts, R. L., 372 Ma´tyus, L., 204, 207, 208(3,4,7) Matzke, A., 385 Matzke, M., 385 Mavila, N., 17 Mayer-Jaekel, R. E., 363, 364, 402 Mayo, L. D., 44 Mazumder, A., 104 McCaffery, J. M., 231 McCain, D. F., 14, 17(31), 271, 273(51) McCombie, R., 43 McCormack, A. L., 85 McDonald, W. H., 85 McDowell, R., 89 McGowan, C. H., 60, 282, 285(2) McInnis, M. G., 391 McKay, R. A., 376, 378, 379 McKeon, F., 3 McKinnell, I. W., 224 McLaughlin, S., 67, 72 McLean, T. K., 384 McLuckey, S. A., 89 McNall, S. J., 5 McNally, C. M., 123 McNamara, D. J., 273 McNeill, R. B., 165 McPartlin, A. E., 361 Means, A. R., 176 Meinhard, M., 283 Meinke, M., 408 Meins, M., 396 Meloen, R. H., 363 Meltzer, P. S., 104 Menard, P., 404 Mendelsohn, A. R., 176 Meng, C. K., 89 Meng, K., 224 Meng, T.-C., 304, 307, 308, 314(16), 315(16), 316(16), 317(10a,10b) Mercer, W. E., 338 Merlevede, W., 5, 6(12,13), 9, 16(12), 189, 197(14), 198(14) Merrifield, B., 262 Mertz, P., 289, 290(2) Mesirov, J. P., 104 Meskiene, I., 288

433

AUTHOR INDEX

Mewes, H. W., 96 Meyer, H. E., 5, 6, 6(12), 7, 7(15), 8(15), 9, 16(12), 398 Meyers, R. E., 65 Michalon, A., 243 Mieskes, G., 282 Milan, D., 3 Milarski, K. L., 273 Miles, W., 56 Miliaresis, C., 43 Miller, B., 384 Miller, P. E., 89 Miller, P. S., 376 Miller, R. H., 191 Mills, K. I., 104 Millward, T. A., 4, 114 Milton, R. C. d. L., 264 Milton, S. C. F., 264 Minna, J., 391 Mirabelli, C. K., 379 Miraglia, L. J., 376, 377 Mischak, H., 122 Mistl, C., 396 Mittler, R. S., 219 Miura, M., 363 Miyamoto, E., 339 Miyawaki, A., 231, 235 Mjalli, A. M. M., 275, 279(65) Moffat, J., 280 Moller, N. P., 305 Monard, D., 396 Moneti, G., 307 Monks, A., 341 Monteith, D. K., 377 Montminy, M., 243 Moolenaar, W. H., 237 Moore, D. D., 341 Moorhead, G., 97, 135, 141, 147, 151, 152, 164 Moos, M., Jr., 201 Moran, E. J., 275, 279(65) Moreno, C. S., 202 Morgan, D. O., 230 Morgan, R. A., 362 Morgenstern, J. P., 193 Mori, K., 372 Morita, F., 244, 245(11), 248(11), 249(11) Morita, K., 372 Morrice, N., 151, 164 Morris, A. J., 67, 72 Morris, H. R., 89

Mortensen, O. H., 305 Moscat, J., 363 Motoike, H., 149, 151(33) Mu, J., 408 Muda, M., 104, 362, 363(7), 371(7) Mudge, L.-M., 114, 116(3), 117(3), 119(3), 121(3) Muimo, R., 57 Muir, L. W., 414 Mukai, H., 245 Muller, D., 339 Muller, G., 348 Muller, S., 148, 153(27) Muller, U., 394, 400 Mumby, M. C., 114, 157, 176, 188, 361, 362, 363(6), 364(6), 368, 371(6), 397, 398(25), 403 Mun˜oz, I., 358 Munro, S., 20, 80, 135 Murakami, Y., 96 Murata, K., 187 Murphy, R. A., 245 Murray, M., 149 Mushinski, J. F., 122 Muslin, A. J., 149 Mustelin, T., 8 Myers, E. W., 103 Myers, K. J., 377, 378 Myers, M. P., 307, 317(10a,10b) Myles, T., 396, 402(19)

N Nachman, R. J., 363 Nagahashi, S., 350, 351 Nagao, M., 146, 157, 392 Nagata, S., 363 Nagy, P., 204, 208(4), 210, 214, 215(11), 216(8), 217(8), 221(8), 231 Nairn, A. C., 148, 151, 152, 158, 159, 159(15), 243, 321, 322, 330(6), 331(6), 333(6) Nakamura, F., 339 Nakano, T., 244 Nakayama, A., 408 Nakayama, H., 350, 351 Nakayama, M., 376 Nakielny, S., 21 Naldini, L., 348 Namgaladze, D., 290, 291(17), 297(17), 298(17), 302(17)

434

AUTHOR INDEX

Naruszewicz, I., 338 Narvin, W. B., 376 Natali, P. G., 391 Neel, B. G., 3, 223, 224, 276 Nefzi, A., 264 Negishi, M., 372 Negrini, M., 391 Neilsen, P. E., 376 Nelson, A., 308 Nemani, R., 160 Neri, D., 269 Neri, P., 269 Neugebauer, J., 77 Neville, M. E., Jr., 40 Ng, S. C., 264 Ni, L., 404, 407 Nicholas, E. J., 263 Nicklin, P. L., 379 Nicolaou, K. C., 275 Nielsen, J., 275 Nielsen, L. D., 414 Nielsen, P. E., 376 Niemela, S. L., 191 Nigavekar, S. S., 416 Nilsson, I., 305 Nilsson, J., 89 Nimmo, G. A., 171, 243 Nimmo, H. G., 23 Ninomiya-Tsuji, J., 338 Nishimoto, T., 230 Nishimura, M., 6 Nishitoh, H., 372 Nitsch, C., 392(22), 396, 397, 399(22), 402(22) Nitsch, R. M., 390, 392(2,15,24), 394, 397, 400(15), 402 Noble, N. E. M., 3 Noda, M., 224 Noel, J. P., 225, 227(9) Nofziger, T. H., 341 Noiman, E. S., 17 Nolan, G. P., 193 Nova, M. P., 275 Novotny, W., 378 Nunbhakdi-Craig, V., 176 Nuttall, F. Q., 416

O O’brien, R., 67 O’Connor, P. M., 338

Oerlemans, F. T., 126, 127(3) Ofek, P., 338, 347 Ogris, E., 187, 195, 202 O’Hearn, E. E., 391 Ohkura, H., 363, 364 Ohlmeyer, M. H. J., 275 Ohmori, T., 244, 245(11), 248(11), 249(11) Ohnishi, M., 338, 339 Ohst, K., 188, 202(9,10), 392(27), 398 Okada, M., 363 Okada, T., 51 Okano, H., 363 Olanow, W., 336 Oligino, L., 270 Oliver, S. G., 96 Ollendorff, V., 372 Olsen, O. H., 305 Olson, E. N., 290 Olsson, H., 76 Onnerfjord, P., 89 Ono, Y., 245 Ooi, S. L., 404 Oonk, H. B., 363 Oren, M., 339, 347 Orgad, S., 364 Osborne, S. L., 67 Osman, N., 220 Ostresh, J., 264 Ota, I., 8 Otaka, A., 265 Ouimet, C. C., 157 Overton, M. C., 231 Overvoorde, J., 221, 226, 231(12), 233(12), 236, 236(12), 237(28), 239(12) Overwin, H., 266 Owen, P., 54 Owens, S. R., 376, 379 Ozawa, K., 122

P Paddison, P. J., 203, 362, 371(14) Painter, G. F., 66 Pakkala, S., 103 Pallas, D. C., 202 Palva, T., 338 Panico, M., 89 Pappin, D. D. J., 84 Paramio, J. M., 178 Parandoosh, Z., 275

435

AUTHOR INDEX

Parent, C. A., 44 Parge, H. E., 290 Paris, H., 160 Park, I.-K., 333 Park, J. W., 204, 208(4) Parke, G. J., 275 Parker, P. J., 20 Parrish, S., 362 Parrou, J. L., 407, 408, 408(8) Parsons, R., 43 Partridge, J., 85, 101, 145, 151(12) Patel, J. C., 31 Paterson, B. M., 363 Patkaniowska, A., 362 Pato, M. D., 283 Paull, K. D., 341 Pavitt, G. D., 363 Pawson, T., 276 Paxton, T., 89 Payrastre, B., 44 Pear, W. S., 193 Pearl, C. G., 273 Pearson, W. R., 85, 93(12), 101, 145, 151(12) Pei, D., 271 Peleg, I., 123 Pelicci, G., 270 Pelicci, P. G., 270 Pellegrini, M. C., 280 Peltonen, L., 103 Perich, J. W., 5, 6(12), 9, 16(12), 267 Perrimon, N., 363 Perrino, B. A., 297 Pershouse, M. A., 43 Pesis, K. H., 57 Peters, G. H., 305 Petrenko, V. A., 269 Petrova, V., 67, 72 Petty, H. R., 220 Pham, H. T., 391 Philippsen, P., 96, 350, 356, 404 Phillips, J., 379 Picton, C., 18 Piedrafita, L., 349, 351, 351(7), 352(12), 354(12) Pieraccini, G., 307 Pini, A., 269 Pinna, L. A., 3, 4(3), 5, 5(3), 6, 6(12,13), 7, 7(15), 8(15), 9, 14, 16(12), 285, 398 Piribauer, P., 187 Pitcher, J., 18

Plunkett, M. J., 261, 264 Podsypanina, K., 43 Podtelejnikov, A. V., 84 Poels, J., 363 Po¨hlmann, R., 350 Pokorni, K., 356 Pollok, B. A., 231 Poo, H., 220 Porter, K., 376 Portet, C., 396 Poulter, L., 18 Poustka, A., 44 Prats, C., 18, 21 Pratt, W. B., 372 Prescott, A. R., 361 Price, E. R., 3 Price, N. E., 157 Prickett, T. D., 245, 249(17) Priganica, L., 76 Pringle, J. R., 407 Probst, A., 392(14,15), 393, 394, 396, 400(15) Proud, C. G., 14, 23, 31, 363 Puc, J., 43 Puttens, H., 396 Putz, H., 178

Q Qi, Y., 44 Qian, Z., 145, 151(11), 152(11) Qin, J., 84 Qin, K. F., 290 Quadroni, M., 84, 88(1) Quirion, R., 338

R Raanani, P., 103 Rait, V., 376 Ramachandra, C., 280 Rambaldi, A., 103 Ramos, J., 358 Ramponi, G., 307 Rankin, B. M., 219 Rao, A., 289, 290(1) Rapacciuolo, A., 188, 202(9), 392(27), 398 Raposo, G., 44 Ratcliffe, E., 114 Ratnofsky, S., 276 Ray, R., 84, 95 Reader, J. C., 275

436

AUTHOR INDEX

Rebecchi, M. J., 67, 72 Rebischung, C., 356 Rebollo, A., 145, 149(10), 150(10) Reese, C. B., 66 Reiken, S., 149, 151(33) Reinach, P. S., 271 Reinemer, P., 159, 330, 331(16), 337(16) Reiter, T. A., 298, 304(24) Resink, T. J., 160, 170(22), 171(22), 249 Reudiger, R., 188, 391 Revuelta, J. L., 404 Rew, D. A., 104 Reynolds, M. A., 376 Rhee, S. G., 307, 308 Ribera, J. M., 105 Richert, M. M., 126 Riess, O., 391 Riles, L., 404, 407 Riley, T. A., 376 Rink, H., 266 Roach, P. J., 17, 17(1,2), 18, 19, 20, 20(3), 22, 27, 31, 96, 403, 407, 408, 410, 417 Roberts, C. J., 404 Roberts, J. M., 230 Roberts, T., 276 Robinson, G. W., 126, 127(3) Rockman, H., 188, 202(9), 392(27), 398 Rodda, S. J., 262 Rodgers, L., 43 Rodriguez-Pena, A., 224 Roepstorff, P., 84 Roeske, R. W., 271 Roizman, B., 149 Roller, P. P., 265, 270 Roncal, F., 145, 149(10), 150(10) Rose, M., 404 Roskoski, R., Jr., 40 Ross, B. S., 377 Ross, C. A., 391 Ross, K. E., 118, 339, 411 Ross-Macdonald, P., 404 Rostas, J. A. P., 114 Rothman-Denes, L. B., 412 Rotin, D., 237 Rowen, D. W., 408 Roy, S., 268 Rozing, G., 275 Rozman, C., 105 Rudge, S. A., 67, 72 Ruediger, R., 188, 202(10), 397, 398, 398(25)

Ruiz, S., 178 Rulicke, T., 394, 400 Rusconi, S., 384 Rusnak, F., 289, 290, 290(2), 298, 302(14), 303(9,14), 304(24) Russo, G., 391 Rutter, W. J., 314 Ruutu, T., 103 Ruzza, P., 5 Ruzzene, M., 3, 5, 6, 7 Ryder, J. W., 126

S Saarinen, U. M., 103 Sagi-Assif, O., 103 Saito, H., 5, 14, 339 Saitoh, M., 372 Sakaguchi, K., 338 Sakamoto, K., 18 Sakashita, G., 244 Sales, M., 20 Salmeen, A., 307, 317(10a,10b) Salmon, S. E., 262 Saltiel, A. R., 20, 273 Samouilov, A., 290, 298(16), 302(16), 303(16) Sancar, A., 372 Sands, H., 379 Sansig, G., 396 Santi, D. V., 263 Santucci, A., 269 Sanz, P., 96, 97, 98 Sap, J., 225, 227(9) Sarno, S., 189, 197(14), 198(14) Sarshar, S., 275, 279(65) Sasaki, K., 157, 392 Sasmor, H., 376, 378, 379, 379(40) Sastry, L., 270 Sato, Y., 351 Satoh, C., 376 Sawyer, T., 273 Scammell, J. G., 373, 384 Scarlata, S., 67, 72 Schaapveld, R. Q., 126, 127(3) Schaffhausen, L. C., 276 Schaller, M. D., 312 Schechter, P. J., 378 Schelling, D. L., 338 Schepens, J. T., 126, 127(3) Scherens, B., 404

AUTHOR INDEX

Schiavo, G., 67 Schiestl, R. H., 102 Schild, A., 390, 392(2,24), 397 Schillace, R. V., 148, 151(29), 155(29) Schimmack, G., 404 Schlender, K. K., 29, 162, 333, 413 Schlessinger, J., 67, 177, 225, 227 Schmid, H., 267 Schmidt, K., 390, 392(2) Schmitz, R., 270 Schmutz, M., 396 Schneider, G., 312 Schnieder, I., 362 Schoenen, F. J., 275 Schonthal, A. H., 202, 395, 402(18) Schroder, M., 396 Schuck, T., 391 Schultz, J. E., 7, 9 Schuster, S. C., 58, 60(15) Schwartz, D. A., 376 Schweigerer, L., 85, 90(10), 91(10) Schwertfeger, K. L., 126 Scott, J. D., 113, 148, 151(29), 155(29), 164, 170 Scott, M. L., 193 Scrimgeour, A., 20, 21, 22(23), 32(23) Scuderi, A., 363 Scudiero, D. A., 341 Scurr, L. L., 123 Sebestye´n, Z., 204, 208(4), 210, 216(8), 217(8), 221, 221(8), 231 Sefton, B. M., 271 Segrelles, C., 178 Seidman, J. G., 341 Selke, D., 282, 283, 284, 285(12), 286(10) Sella, O., 339 Seltzer, W. K., 391 Selvin, P. R., 231 Sen, G. C., 371 Senba, S., 248 Seniff, D., 341 Senyei, A., 275 Seo, M. S., 308 Se´raphin, B., 351 Sergueev, D., 376 Sergueeva, Z., 376 Seroussi, E., 339 Se´rrano, L., 351 Sevenstyen, F., 262 Shabanowitz, J., 89 Shacter, E., 200

437

Shafer, B., 404 Shah, S., 67, 72 Shahbaz, M. M., 275, 279(65) Shaikh, S., 391 Shanahan, W. R., 378, 379 Shani, N., 339 Shao, J., 372 Sharf, R., 175 Sharom, F. J., 222 Sharp, A. H., 391 Shaw, B. R., 376 Shaw, M., 20 Shea, J. P., 379 Sheen, J., 338 Shen, K., 35, 36(4), 279 Shen, S. H., 338 Shen, Y., 312 Sheng, M., 148, 170 Shenk, T., 176 Shenolikar, S., 147, 148, 152, 153(22), 148(22), 157, 158, 159(15), 162, 243, 245, 249, 259, 321 Sheorain, V. S., 20 Sherr, M., 391 Shevchenko, A., 85, 89, 90(10,18), 91(10) Shi, Y., 362, 371(15), 386 Shibasaki, F., 3 Shih, S.-R., 148 Shima, H., 157, 244, 392 Shima, M., 146 Shirato, H., 244 Shoelson, S. E., 265, 276 Shoemaker, D. D., 404 Shoemaker, J. E. E., 379 Shoemaker, R. H., 341 Shoji, S., 363 Shriner, C. L., 22, 248 Sievers, E. L., 103 Sigler, P. B., 67 Sihta, E. Y., 351 Silberman, S. R., 160 Silverman, N., 363 Silverstein, A. M., 361, 362, 363(6), 364(6), 371(6), 372, 403 Sim, A. T. R., 113, 114, 116(3), 117(3), 119(3), 121(3), 122, 122(4), 123(4), 148 Simanis, V., 150 Simo´n, E., 358 Simon, M. I., 58, 60(15) Simonsen, A., 67

438

AUTHOR INDEX

Simonson-Leff, N., 362, 363(7), 371(7) Sioufi, N. B., 378, 379(40) Skroch, J., 414 Skurat, A. V., 17(1), 18, 19 Slama, J. T., 53, 305 Sleeman, J. E., 146, 152(18), 222, 223(27) Slonim, D. K., 104 Smith, A., 104 Smith, A. J. H., 139 Smith, C. H., 23 Smith, G. J., 8, 342 Smith, G. P., 269 Smith, J. A., 341 Smorodinsky, N. I., 339 Smyth, M. S., 265 Snavley, D. F., 271 Sneddon, A. A., 104 Snyder, B., 218 Snyder, M., 404, 407 Soderling, T. R., 20, 23, 297 Sohaskey, M. L., 271 Sola, M. M., 145, 146(7), 164, 189, 197(13), 198(13) So¨ling, H.-D., 282 Solomon, M. J., 8, 338, 339, 411 Sommer, D., 298 Song, H., 3 Song, O., 176 Songyang, Z., 276 Sontag, E., 176 Soohoo, C., 362, 371(15), 386 Sookhai-Mahadeo, S., 404 Sorokin, A., 227 Sotiroudis, T. G., 14 Sozzi, G., 391 Speth, M., 160 Spevak, W., 414 Spillantini, M. G., 396 Spitalny, G. L., 219 Squire, A., 223 St. Jean, A., 102 Stalmans, W., 20, 139, 145, 146, 147, 148, 148(15,22), 149, 150, 150(26,31), 151, 151(31), 152(15,31), 153(22), 164, 243 Stambolic, V., 44 Standing, K. G., 89 Stark, G., 400 Stark, M. J., 145, 338, 348 Starkova, N., 321 Steck, P. A., 43

Steele, M., 18, 20, 21, 22(23), 32(23) Steen, R. L., 148, 151(28), 155(28) Steiner, S., 356 Stemmer, P. M., 295, 298 Stenmark, H., 67 Stevens, L. R., 66 Stewart, A. A., 290, 294(18), 295(18), 299(18), 300(18) Stewart, M., 104 Still, W. C., 275 Stock, A. M., 57 Stoker, A. W., 224 Stokoe, D., 66 Stoolman, L. M., 105 Stoppini, L., 339 Storm, D., 243 Storms, R. K., 404 Strack, S., 146, 152(19), 157, 159, 159(10), 160(10), 164, 165, 171, 174(10,25), 176 Strada, S. J., 249 Stragier, P., 7 Stralfors, P., 145, 160, 170(22), 171(22), 249 Strater, N., 290 Strathern, J. N., 404 Street, A. J., 145, 158, 159(13), 330, 331(11), 361 Streuli, M., 14, 126, 127(3) Strovel, E. T., 339 Strub, M.-P., 298 Stryer, L., 204, 205(1), 230 Stull, J. T., 255 Su, J., 225, 227(9) Sugimura, T., 157 Sugino, A., 353 Sui, G., 362, 371(15), 386 Sullivan, J. E., 8, 342 Sultan, C., 105 Summers, J., 376 Summerton, J., 376 Sumner, A. J., 391 Sun, H., 36 Sundaresan, M., 314 Sunkel, C. E., 364 Superti-Furga, G., 44 Sussman, D. J., 339 Suter, U., 44 Sutton, A., 354 Suzuki, M., 244, 245(11), 248(11), 249(11) Suzuki, Y., 18, 20, 21, 22(23), 32(23) Svensjo, T., 340

AUTHOR INDEX

Swanson, R. N., 275 Swanson, R. V., 58, 60(15) Swanson, S. A., 298 Swedlund, B., 43 Sweitzer, S. M., 363 Swingle, M., 384 Swinnen, E., 363 Sytwu, T., 280 Szallasi, Z., 122 Szedlacsek, S. E., 237 Szo¨llo¨si, J., 203, 204, 207, 208(3,4,7), 210, 214, 215(11), 216(8), 217(8), 221, 221(8), 231

T Tabata, S., 408 Tahtiharju, S., 338 Takahashi, L. H., 275 Takahashi, T., 363 Takai, A., 173 Takeda, A., 219, 226 Takeda, Y., 23, 24(33), 27 Takekawa, M., 339 Takenawa, T., 66, 67 Takio, K., 291, 296(19) Takizawa, N., 257 Takle, E., 308 Tamayo, P., 104 Tamura, M., 146, 152(19), 164, 174(25) Tamura, S., 4, 7(9), 8(9), 338, 339 Tan, Y. S., 416 Tang, J., 379 Tanner, K. G., 307 Taske´n, K., 148, 151(28), 155(28) Tatchell, K., 149, 150(31), 151(31), 152, 152(31), 414 Tavalin, S. J., 148, 170 Tavoloni, N., 338 Tavtigian, S. V., 43 Taylor, D., 255 Taylor, G. S., 43, 44, 47(13), 49, 51(19), 53, 305 Taylor, I. W., 8, 342 Taylor, J. K., 377, 378 Tempest, P. A., 275 Temsamani, J., 379 Teng, D. H., 43 Tenza, D., 44 Terry-Lorenzo, R. T., 245

439

Tertoolen, L. G., 221, 224, 226, 231(12,13), 233(12,13), 236, 236(12,13), 237(13,28), 239(12), 240(13) Testa, S. M., 376 Teste, M. A., 408 Tettelin, H., 96 Thellmann, M., 348 Theodosiou, A., 305 Thieme-Sefler, A. M., 271, 273 Thomas, C. L., 67 Thomas, J. A., 29, 413 Thompson-Jaeger, S., 414 Tierney, S., 341 Tiganis, T., 273 Tiran, Z., 124 Tirasophon, W., 339 Tisi, D., 307, 317(10a,10b) Tivel, K. L., 376 Tobiume, K., 372 Toker, A., 43 Tokunaga, T., 377 Tolney, M., 396 Tong, Y., 338 Tonks, N. K., 3, 171, 224, 273, 304, 305, 306, 307, 308, 314(16), 315(16), 316(16), 317(10a,10b) Torfs, H., 363 Torgersen, K. M., 44 To´th, A., 152 Tountas, N. A., 243 Trainor, G. L., 379 Tregear, G. W., 267 Trent, J. M., 104 Trinkle-Mulcahy, L., 146, 152(18), 222, 223(27) Trojanowski, J. Q., 391, 396 Truong, L., 378, 379, 379(40) Tsai, A. Y., 14 Tsay, H.-J., 148 Tsien, R. W., 304 Tsien, R. Y., 231, 235 Tsou, C. L., 41 Tsukitani, Y., 361 Tu, J., 96 Tucker, P., 290 Tugendreich, S., 416 Tung, H. Y., 160, 170(22), 171(22) Turner, D. H., 376 Turowski, P., 188, 402 Tuschl, T., 362, 371, 371(13), 385 Tycko, B., 43

440

AUTHOR INDEX

U Uhl, G. R., 245 Ui, M., 14, 51 Ullrich, A., 227 Ullrich, V., 290, 291(17), 297(17), 298(17), 302(17) Urban, G., 373, 384 Urbe, S., 44 Urlaub, H., 362 Urlinger, S., 348 Uyeda, K., 6

Volgelberg-Ragaglia, V., 391 Volland, J. P., 263 Voltz, J. W., 148 Vorechovsky, I., 391 Vorherr, T., 267 Vorm, O., 89, 90(18) Vrana, K. E., 40 Vromirski, M., 89 Vulijanic, T., 268 Vulsteke, V., 147, 148, 148(22), 149, 150, 150(26,31), 151(31), 152(31), 153(22)

V

W

Vaghefi, M. M., 376 Valerio, R. M., 267 Valle, G., 404 Va´mosi, G., 210, 214, 215(11), 216(8), 217(8), 221(8) Vanden Broeck, J., 363 van der Kaay, J., 20, 77, 79 van der Krogt, G. N., 231 van der Wijk, T., 224 Vande Woude, G. F., 338 van Etten, I., 237 van Eynde, A., 144, 145, 147, 148, 148(22), 149, 150, 150(26), 153(22), 243 van Montfort, R. L. M., 307, 317(10a,10b) Van Poyer, W., 363 Van Regenmortel, M. H. V., 148, 153(27) Veillette, A., 312 Vemuri, B., 17 Venter, J. C., 103 Veronneau, S., 404 Verveer, P. J., 223 Vetrani, C., 270 Vetter, S. W., 5, 16, 260, 261, 267, 274, 278, 278(62) Vieria-Saecker, A. M., 391 Vigna, E., 348 Vilardo, P. G., 17, 18, 20, 21, 22(23), 32(23) Villa-Moruzzi, E., 114 Villar-Palasi, C., 33 Vincent, O., 408 Virbasius, J. V., 66, 67 Virshup, D. M., 157, 175, 187, 363 Viti, F., 269 Vlattas, I., 280 Voet, M., 404 Volckaert, G., 404

Wach, A., 350, 356 Wade, J. D., 267 Wadzinski, B. E., 146, 152(19), 156, 157, 159, 159(10), 160(10), 164, 165, 171, 174(10,25), 176 Waelkens, E., 5, 6(13), 151 Wagers, A. J., 105 Wagner, C. D., 275 Wagner, D. S., 275 Wakula, P., 147 Walberg, M. W., 176 Walder, J. A., 376 Waldner, A., 376 Walker, D. M., 44 Wallace, M. J., 237 Walsh, C. T., 5 Walsh, E. P., 145 Walter, G., 188, 202(9,10), 391, 392(27), 397, 398, 398(25) Walther, D., 245 Wang, C. Y., 404 Wang, H., 245, 338 Wang, J. Z., 391 Wang, K., 280 Wang, P., 271 Wang, Q. M., 31, 35, 40(5), 42(5) Wang, S. I., 43, 270 Wang, S. S., 266, 391 Wang, T., 377 Wang, X., 289, 290, 291(15), 295, 296, 297(15,21), 298(15,21), 299(21), 303(21) Wang, X. B., 245 Wang, Y., 19, 283 Wang, Z.-M., 35, 40(5), 42(5), 407, 410 Wang, Z.-X., 35, 38(6), 39 Ward, T. R., 404

441

AUTHOR INDEX

Wardle, R. L., 245 Warmka, J., 8 Watanabe, T. A., 148, 158, 159(15), 321, 322, 331, 336(21), 337(21), 378, 379(40) Watanabe-Fukunaga, R., 363 Waters, C. M., 105 Watson, A., 104 Watson, S. J., 157 Watt, P. W., 20, 21, 139 Watt, S. A., 64, 84 Webb, M. R., 38 Weber, K., 362, 371(13), 385 Wei, H., 202 Wei, Q., 363 Weiser, D. C., 245 Weiss, A., 219, 225, 226, 227(14) Weissmann, C., 394, 400 Welihinda, A. A., 339 Weller, D. D., 376 Wellings, D. A., 265 Welsh, G. I., 31 Wendeborn, S., 376 Wennogle, L. P., 280 Wera, S., 402 Werb, Z., 126 West, A. H., 57 West, R. W., Jr., 178 Westphal, R. S., 148, 159, 170, 176 White, C. L. III, 176 White, P. D., 265, 267 Whitehouse, C. M., 89 Wickstrom, E., 375 Wiederhold, K. H., 396 Wieringa, B., 126, 127(3) Wigler, M. H., 43, 275 Wilhelmy, J., 404 Willi, S., 44 Williams, D. D., 14, 363 Williams, D. H., 18 Williams, R. S., 290 Wilm, M., 85, 89, 90(10,18), 91(10) Wilsbacher, J. L., 39, 41(10) Wilson, C. J., 84 Wilson, W. A., 403, 407, 410 Winberg, G., 270 Winkler, T., 340 Winston, S., 89 Winzeler, E. A., 404 Wirtz, K. W. A., 213 Wiseman, B. S., 126

Wishart, M. J., 44, 305 Witz, I. P., 103 Witzel, H., 290 Wolf, G., 265 Wolfer, D. P., 394 Wong, L. K. H., 54 Wong, S. F., 89 Woods, R. A., 102, 355 Woodsome, T. P., 245, 253(12) Worby, C. A., 362, 363(7), 371(7) Wouters, F. S., 213 Wright, D. J., 17 Wu, D., 339 Wu, J. J., 187, 219, 226 Wu, L., 35, 36(4), 279 Wu, P., 207, 231 Wurmser, A. E., 66 Wyatt, J. R., 377 Wymann, M., 44

X Xiao, X. Y., 275 Xie, L., 273 Xu, Z., 219, 226, 227(14)

Y Yacyshyn, B. R., 379 Yalcin, A., 362, 371(13), 385 Yamaguchi, Y., 372 Yamamoto, K. R., 363, 408 Yamamoto, S., 377 Yamamoto, T., 377 Yamashita, I., 408 Yamashita, K., 230 Yamazaki, T., 351 Yamin, M., 224 Yan, X., 265 Yanagawa, Y., 338, 363 Yang, H., 350 Yang, J., 152 Yang, S.-A., 290 Yang, S.-I., 368 Yang, Y., 404 Yano, O., 377 Yao, F., 340 Yao, J., 290, 302(14), 303(14) Yates, J. R., 85 Yates, J. R., III, 85, 89 Yazawa, M., 245, 248

442

AUTHOR INDEX

Ye, B., 265 Yen, C., 43 Yen, G., 404 Yeong, F. M., 202 Yguerabide, J., 218 Yokoyama, S., 66, 67 Yoshikawa, S., 363 Yoshinaga, S. K., 363 Young, R. A., 84 Youngman, E., 404 Yu, J. W., 66 Yu, K., 404 Yu, L., 290, 302(14), 303(9,14) Yu, X. X., 188 Yu, Z. X., 314 Yuffe, M. B., 66 Yung, H. Y., 249 Yung, W. K., 43 Yuryev, A., 280

Z Zabarovsky, E. R., 270 Zaidi, T., 391 Zamboni, R., 280 Zamore, P. D., 385 Zander, N. F., 4 Zardi, L., 269 Zauberman, A., 339 Zaucha, J. A., 176 Zelenka, U., 414 Zerner, B., 301, 302(27) Zhang, A. J., 158, 162(14), 249 Zhang, H., 20, 22(23), 32(23), 338 Zhang, J., 5, 35, 36(4), 330 Zhang, L., 162, 249

Zhang, L. F., 331 Zhang, P. W., 245 Zhang, W., 22 Zhang, Y.-L., 273 Zhang, Z., 5, 146, 157, 330 Zhang, Z.-H., 39, 41(11) Zhang, Z. J., 330, 331 Zhang, Z.-Y., 5, 14, 16, 17(31), 34, 35, 36(3,4), 38(6), 40(5), 42(5), 260, 261, 271, 273, 273(51), 278, 279 Zhao, R., 44 Zhao, S., 146, 165, 330, 372 Zhao, Y., 35, 36(3), 38(6), 84 Zhao, Z., 3 Zhao, Z. J., 44 Zhen, Q., 245 Zheng, S., 339 Zheng, S. F., 407 Zhou, B., 34, 35, 36(4), 38(6), 39, 40(5), 41(11), 42(5) Zhu, G., 273 Zhu, L., 308 Zhu, T., 339 Zick, Y., 314 Zilbering, A., 308 Zirattu, S. D., 330 Zolnierowicz, S., 4, 114 Zondag, G. C., 237 Zounes, M. C., 376 Zuber, J. F., 270 Zucconi, A., 270 Zuckermann, R. N., 264 Zuo, Z., 382, 384 Zweier, J., 290, 298(16), 302(16), 303(16)

Subject Index

A Acute myelogenous leukemia, see PYST2 Alkaline phosphatase, see Glycophosphatidylinositol-anchored alkaline phosphatase AML, see Acute myelogenous leukemia Antisense knockdown, see Protein phosphatase-5 Atomic absorption spectroscopy, calcineurinassociated metals, 301

B B55 knockout effects in mice, 394 protein phosphatase-2A regulatory subunit, 175–176, 187–189 Bis[sulfosuccinimidyl]-suberate, receptor protein tyrosine phosphatase crosslinking, 226–227 Blastocyst, culture studies of phosphatase transgenic mice, 399–400

C Calcineurin activity assay anaerobic assays, 294–295 crude tissue extracts, 295–296 incubation conditions, 293–294 materials, 292–293 principls, 291 substrate preparation, 291–292 antibody preparation, 293 calcium/calmodulin-dependent inactivation, 296–297 calcium dependence, 290–291 metal cofactor studies atomic absorption spectroscopy, 301 electron paramagnetic resonance of iron oxidation state, 301–302 iron–zinc active center, 302–304 purification of enzyme, 299–301

oxidative regulation inactivation, 291 physiologic role, 304 superoxide dismutase protection against inactivation, 297–298 substrate specificity, 7, 290 therapeutic targeting, 289–290 CD45 dimerization studies, 225, 231, 237 fluorescence resonance energy transfer studies, 219–221 isoforms, 218–219 phosphotyrosine phosphatase activity, 218–219 CheA dephosphorylation, see Protein histidine phosphatase Combinatorial libraries, protein phosphatase characterization applications, 260–261 deconvolution of screening data conceptual road map, 276–277 examples, 276, 278–281 positional scanning, 275–276 tagging for compound identification, 274–275, 279 overview, 261 peptide synthesis activation reagents, 267 amino acid derivatives, 267 coupling reactions, 268 deprotection and cleavage, 268 Fmoc versus Boc chemistry, 265–266 Fmoc removal, 268 N-terminal modification, 269 phage display, 269–270 phosphopeptides, 269 postcleavage work-up, 269 reaction vessels, 266 solid supports, 266 solvents, 267 phosphatase activity assays fluorescence continuous assay, 271–272 inhibitor screening, 273

443

444

SUBJECT INDEX

overview, 270–271 quenched assay, 272–273 phosphatase binding assays, 273–274 synthesis of libraries guidelines, 281 mixture coupling, 262 one-bead one-compound approach, 262–265 parallel synthesis, 263 phosphopeptide libraries, 264–265 solid phase synthesis, 361–262 verification of results, 282 CPI-17, protein phosphatase-1 inhibition, 244–245

D DARPP-32, protein phosphatase-1 inhibition, 244, 321, 323–324 DNA microarray, PYST2 overexpression in acute myelogenous leukemia Atlas system, 104 bone marrow-derived leukocyte preparation, 105 considerations for clinical use, 111–113 differential gene expression analysis, 107–109 probe preparation, 106 verificaton with reverse transcriptase-polymerase chain reaction, 109–110 Donor photobleaching fluorescence resonance energy transfer microscopy, see Fluorescence resonance energy transfer

E Electron paramagnetic resonance, calcineurin iron oxidation state, 301–302 EPR, see Electron paramagnetic resonance ERK2, see Extracellular regulated kinase-2 Extracellular regulated kinase-2 protein phosphatase, see MKP3 stoichiometry of phosphorylation, 36

F FASTF, sequence analysis from mixed peptide sequencing, 85–88, 93–95

FASTS, sequence analysis from mixed peptide sequencing, 85–88, 93–95 Fluorescence microscopy donor photobleaching fluorescence resonance energy transfer microscopy data acquisition, 212–214 efficiency, 211, 213 intensity measuremement, 211–212 irreversibility, 212 intensity based fluorescence resonance energy transfer microscopy data acquisition, 215–216 principles, 214–215 limitations, 216–218 protein phosphatase-2A translocation studies using immunofluorescence microscopy, 121–123 Fluorescence resonance energy transfer donor photobleaching fluorescence resonance energy transfer microscopy data acquisition, 212–214 efficiency, 211, 213 intensity measuremement, 211–212 irreversibility, 212 efficiency, 204, 206 flow cytometry measurements autofluorescence, 209 correction factors, 208 data acquisition, 210–211 energy transfer efficiency, 207–209 intensity expression, 208–209 glycophosphatidylinositol-anchored alkaline phosphatase studies, 222 intensity based fluorescence resonance energy transfer microscopy data acquisition, 215–216 principles, 214–215 pleckstrin homology domain/lipid sensor complexes in phosphoinositide detection, 68–69 protein phosphatase-1 studies, 222–223 protein tyrosine phosphatase-1B studies, 223 receptor protein tyrosine phosphatase dimerization studies CD45 studies, 219–221 green fluorescent protein variants, 231, 233, 235–236 instrumentation, 234–235

445

SUBJECT INDEX

principles, 230–231, 233 receptor protein tyrosine phosphatase-, 221–222 single cell spectral microscopy studies, 236 receptor protein tyrosine phosphatase- dimerization studies, 221–222 theory, 204–207 FRET, see Fluorescence resonance energy transfer

G Glycogen phosphorylase assay in Saccharomyces cerevisiae gene deletion studies, 414–416 protein phosphatase assay substrate, 22 Glycogen synthase assay in Saccharomyces cerevisiae gene deletion studies, 413–414 kinases, 18, 20 phosphorylation sites, 18–20 purification from rabbit skeletal muscle anion-exchange chromatography, 23–24 dephosphorylated protein -amylase digestion, 26 anion-exchange chromatography, 25–26 dephosphorylation, 25 ethanol precipitation, 25–26 extraction, 24–25 Sepharose CL-4B chromatography, 26–27 yield, 27 materials, 23 overview, 23 phosphorylated protein, 27–28 Glycogen synthase phosphatase assays coupled dephosphorylation/activation assay, 32–34 radioactive assay, 31–32 Saccharomyces cerevisiae gene deletion studies, 412–413 substrate preparation glycogen synthase phosphorylation for coupled assay, 30 kinase preparation, 31 radiolabeling of glycogen synthase, 30–31

functions, 17–18, 20 isoforms, 20 Glycophosphatidylinositol-anchored alkaline phosphatase, fluorescence resonance energy transfer studies, 222 GS, see Glycogen synthase

H Histidine phosphatase, see Protein histidine phosphatase

I I-1, protein phosphatase-1 inhibition, 243, 245, 321 I-2, protein phosphatase-1 inhibition, 243, 245 I-4, protein phosphatase-1 inhibition 243–244 Immunoprecipitation protein phosphatase-1 interacting protein coimmunoprecipitation activity assay of complexes, 141–142 antibody incubation, 154–155 antibody selection, 136–138 applications, 142–144 buffers, 139 coupling of antibodies to protein GSepharose, 139–140 dissociating peptides release of enzyme activity, 141–142, 144 selection, 138 immunoadsorbed complex analysis with gel electrophoresis, 142 immunoprecipitation of protein complexes, 140–141 inhibitors, 139 isoform protein–protein interactions, 167–170 overview, 135–136 principles, 150 substrates for activity assay, 138–139 protein phosphatase-2A, coimmunoprecipitation for catalytic subunit–protein interaction characterization antibody coupling to protein A-Sepharose, 195–196 immunoprecipitation reaction, 196 overview, 192

446

SUBJECT INDEX

protein phosphatase-2A translocation assay, 123–124 receptor protein tyrosine phosphatase dimerization studies, 236–238 Iron–zinc active center, calcineurin, 302–304

K KEPI, protein phosphatase-1 inhibition, 245

L Lipid phosphatases, see Myotubularins; PTEN phosphatases

M Malachite green myotubularin assay advantages and limitations, 54–55 incubation conditions, 54–55 materials, 54 substrate specificity findings, 55–56 phosphothreonyl peptide substrate characterization with protein phosphatase assay incubation conditions, 15–16 materials, 15 principles, 14–15 protein phosphatase-2C activity assay, 342 PTEN assay advantages and limitations, 54–55 incubation conditions, 54–55 materials, 54 substrate specificity findings, 55–56 Mammary gland development in mouse, 126–128 receptor protein tyrosine phosphatase-" effects in transgenic mice, see Receptor protein tyrosine phosphatases MAPK, see Mitogen-activated protein kinase Mass spectrometry peptide mass fingerprinting, 84–85 peptide mass tag, 85 peptide sequencing using tandem mass spectrometry FASTF analysis, 93–95 FASTS analysis, 93–95

ionization techniques, 88–89 mass analysis, 89–90 sample preparation gel staining and in-gel digestion, 90–93 overview, 88 phosphorylation site analysis of protein phosphatase-1 phosphoprotein inhibitors, 258–260 Microcystin affinity chromatography, protein phosphatases and interacting proteins, 150–151, 155–156 Mitogen-activated protein kinase extracellular regulated kinase-2, see Extracellular regulated kinase-2 phosphatase, see PYST2 Mixed peptide sequencing FASTF/FASTS analysis of sequences overview, 85 sequence alignment, 86–88, 93–95 Reg1p regulatory protein effects on yeast phosphoproteome, 101 sample preparation, 87 sequencer, 86 MKP3 enzyme-coupled spectrophotometric assay calculations, 39–40 incubation conditions, 39 materials, 39 principles, 38–39 extracellular regulated kinase-2 activitycoupled assay calculations, 41–42 incubation conditions, 41 materials, 41 principles, 40–41 radioactive assay calculations, 37 extracellular regulated kinase-2 radiolabeling, 35–36 incubation conditions, 36–37 MS, see Mass spectrometry Myotubularins assay using fluorescent phosphoinositides incubation conditions, 50–51 materials, 50 principles, 49 substrate specificity findings, 51–52 thin-layer chromatography, 51 assay using malachite green

SUBJECT INDEX

advantages and limitations, 54–55 incubation conditions, 54–55 materials, 54 substrate specificity findings, 55–56 mutations in disease, 44 purification of recombinant proteins Escherichia coli expression cell growth and induction, 46 materials, 45–46 nickel affinity chromatography, 46–47 mammalian cell expression advantages and limitations, 49 FLAG-tagged proteins in human embryonic kidney cells, 48–49 materials, 47–48 substrates, 44

N NIPP-1, protein phosphatase-1 inhibition, 243–244

O Oxidative regulation, see Calcineurin; Protein tyrosine phosphatases

P Peptide synthesis, see Combinatorial libraries PHI-1, protein phosphatase-1 inhibition, 245 Phosphatidylinositol 3-kinase, assay using pleckstrin homology domain/lipid sensor complexes, 72–75 Phosphoinositides metabolism, 64–65 phosphatases, see Myotubularins; PTEN phosphatases pleckstrin homology domain/lipid sensor complexes in detection cell-based assays using green fluorescent protein chimeras controls, 81–82, 84 labeling, 80–81 rationale, 79–80 exogenous phosphoinositide detection, 70, 72 extracted lipid assay, 75–79 fluorescence resonance energy transfer, 68–69

447

formation of complex, 68 phosphatidylinositol 3-kinase assay, 72–75 principles, 68–69 PTEN assay, 75 protein binding domains, 66–67 signal transduction, 65–66 Phosphorylase a, preparation for DARRP-32 studies, 323 Phosphothreonyl peptide substrates malachite green assay for protein phosphatases incubation conditions, 15–16 materials, 15 principles, 14–15 protein phosphatase-2A versus protein phosphatase-2C substrate specificity, 6–9, 16–17 radioactive assays for protein phosphatases incubation conditions, 12–13 phase separation for radiophosphate determination, 12–13 phosphocellulose paper absorption assay, 13–14 phosphopeptide separation from nonphosphorylated form, 11–12 radiolabeling of peptides, 9–11 PHP, see Protein histidine phosphatase PI3K, see Phosphatidylinositol 3-kinase Pleckstrin homology domain, see Phosphoinositides PP1, see Protein phosphatase-1 PP2A, see Protein phosphatase-2A PP2B, see Calcineurin PP2C, see Protein phosphatase-2C PP4, see Protein phosphatase-4 PP5, see Protein phosphatase-5 Protein histidine phosphatase assays CheA assays autoradiography assay, 61 buffers, 58–60 principles, 57–58 substrate preparation, 60 trichloroacetic acid precipitation assay, 60–61 overview, 57–58 purification from rabbit liver ammonium sulfate precipitation, 62–63 anion-exchange chromatography, 62

448

SUBJECT INDEX

dye affinity chromatography, 64 gel filtration chromatography, 63–64 materials, 61–62 vertebrate features, 56–57 Protein phosphatase-1 classification, 144–145 fluorescence resonance energy transfer studies, 222–223 interacting protein abundance, 145 interacting protein identification activity inhibition, 146, 174 coimmunopecipitation of cell extracts antibody incubation, 154–155 isoform protein–protein interactions, 167–170 principles, 150 glutathione S-transferase pull-down assay binding conditions, 153–154 preblocking of glutathione-agarose, 153 principles, 149 pull-down and detection, 154 microcystin affinity chromatography elution and gel electrophoresis, 155–156 lysate preparation, 155 principles, 150–151 miscellaneous techniques, 151–152 overlay assay binding site analysis, 148, 153 blotting, 152 far-Western overlay assays of isoform protein–protein interactions, 164–167 principles, 147–148 washing and detection, 152–153 isoforms activity assay for binding protein characterization buffers, 170 detection, 172–173 incubation conditions, 171–172 substrate preparation, 170–171 antibody generation, 159–160, 174 immunoaffinity chromatography of catalytic subunits, 162–164 isolation from rat brain, 160–162 recombinant protein expression, 158–159, 173 tissue distribution, 157 types, 146, 156–157

phosphoprotein inhibitors assay of inhibitory activity, 249 CPI-17, 244–245 DARPP-32, 244, 321, 323–324 I-1, 243, 245, 321 I-2, 243, 245 I-4, 243–244 KEPI, 245 NIPP-1, 243–244 PHI-1, 245 phosphorylation assays antibody preparation, 250–252 glycerol polyacrylamide gel electrophoresis mobility shift assay, 255–256 lutidine polyacrylamide gel electrophoresis mobility shift assay, 257–258 mass spectrometry analysis of phosphorylation sites, 258–260 overview, 249–250 sodium dodecylsulfate-polyacrylamide gel electrophoresis mobility shift assay, 253–254 urea polyacrylamide gel electrophoresis mobility shift assay, 256–257 Western blot, 250–253 phosphorylationinenzymeregulation,246 phosphorylation of recombinant inhibitors, 247–248 recombinant inhibitor preparation, 246–247 protein–protein interaction immunoassays activity assay of complexes, 141–142 antibody selection, 136–138 applications, 142–144 buffers, 139 coupling of antibodies to protein G-Sepharose, 139–140 dissociating peptides release of enzyme activity, 141–142, 144 selection, 138 immunoadsorbed complex analysis with gel electrophoresis, 142 immunoprecipitation of protein complexes, 140–141 inhibitors, 139 overview, 135–136

SUBJECT INDEX

substrates for activity assay, 138–139 recombinant protein expression and purification activity assays, 324 affinity chromatography glutathione S-transferase fusion proteins, 328 nickel affinity chromatography, 329 PNUTS resin, 328–329 baculovirus–Sf9 cell system for expression and purification characterization of purified enzymes, 334–338 histidine-tagged enzyme, 327–328, 336–337 rat PP1C, 326–327, 334–336 Escherichia coli system for expression and purification characterization of purified enzymes, 330–334 histidine-tagged enzyme, 325–326, 333–334 rabbit PP1C, 324–325, 330–333 isoform expression, 158–159, 173 materials, 322 Western blotting, 329 Reg1p regulatory protein, see Reg1p RVxF sequence in protein–protein interactions, 135, 147, 321 Protein phosphatase-2A B55 regulatory subunit, 175–176, 187–189, 394 B subunit families, 390 knockdown, see RNA interference, protein phosphatase knockdown in Drosophila phosphothreonyl peptide substrate assays, see Phosphothreonyl peptide substrates Ras recruitment system in protein–protein interaction analysis advantages, 186–187 bait construction, 179–180 testing, 182 controls, 182 library screening, 183–184 materials, 178 media, 178–179 plasmid isolation of yeast and analysis, 185–186

449

principles, 176–178 validation of interactions, 186 Western blot, 182 yeast transformation, 180–181 regulating proteins, 113–114 sit4 hal3 conditional mutant construction using tetracycline-regulable promoter functional analysis of induced cells, 356–358 promoter substitution, 354–355 rationale, 354 transformation and selection, 355–356 verification of correct integration, 356 site-directed mutagenesis A–C subunit interaction studies, 188–189 activity assay for catalytic subunit–protein interaction characterization controls, 198–199 incubation conditions, 199–200 overview, 192 substrate preparation, 196–198 applications in holoenzyme reconstitution, 202–203 coimmunoprecipitation for catalytic subunit–protein interaction characterization antibody coupling to protein A-Sepharose, 195–196 immunoprecipitation reaction, 196 overview, 192 extraction of mutant proteins from cells, 194–195 polymerase chain reaction mutagenesis commercial system, 190 mutagenesis reaction, 191 primer design, 190 restriction digestion, 191 transformation of bacteria, 191–192 stable expression of mutants in BOSC 23 cells, 193–194 Western blot analysis of complex formation, 201–202 substrate specificity, 4, 6–7 subunits, 175, 187–189, 390–391 transgenic mouse studies activity assays, 398–399 advantages and limitations, 400–403 approaches in functional studies, 391–392

450

SUBJECT INDEX

blastocyst culture studies, 399–400 B0 /PR56 regulatory subunit expression, 394–395 C dominant negative mutant mice Dom1 line, 396–397 Dom5 line, 397 overview, 395 knockout mice C gene knockout, 392–394 phenotypes, 392 regulatory subunit knockouts, 394 PR65 dominant negative mutant mice, 397–398 translocation assays in rbl-2H3 mast cells cell activation, 117–118 cell culture, 115–116 cell sensitization, 116–117 coimmunoprecipitation of phosphatases and myosin, 123–124 -hexosaminidase secretion assay, 118–119 immunofluorescence microscopy, 121–123 overview, 114–115 subcellular fractionation, 119–120 translocation inhibitor studies, 121 Protein phosphatase-2B, see Calcineurin Protein phosphatase-2C assays divalent cation dependence differences by substrates, 286 lipophilic compound activation, 286–288 substrates BAD, 284 casein, 284 phosphopeptides, 283–285 storage, 286 classification, 338 divalent cation dependence, 282–283, 286 domains, 338–339 inducible expression of PP2C in human embryonic kidney cells cell viability assay, 341–342 clone isolation, 343 induction effects on cells apoptosis, 346–347 cell cycle arrest, 346–347 colony formation inhibition, 345–346 proliferation inhibition, 345–346

malachite green activity assay, 342 plasmids, 339–340 plating efficiency, 340 tetracycline-dependent induction characteristics, 343–345 T-Rex tetracycline-regulated expression system, 340 Western blotting, 341 isoforms, 282 monoclonal antibody preparation, 341 phosphothreonyl peptide substrate assays, see Phosphothreonyl peptide substrates substrate specificity, 4, 6–7, 338 Protein phosphatase-4, knockdown, see RNA interference, protein phosphatase knockdown in Drosophila Protein phosphatase-5 activation, 372–373 antisense knockdown cellular uptake and retention, 378–379 controls, 379–380 modified oligodeoxynucleotides, 374–378 principles, 374 screening, 379–380, 382, 384 sequence design, 377–378, 383, 389–390 domains, 372 protein–protein interactions, 372 RNA interference knockdown, see also RNA interference, protein phosphatase knockdown in Drosophila controls, 385 principles, 384–385 RNA synthesis, 385–386, 388 sequence selection, 383, 385–386, 389–390 vectors, 388 Protein tyrosine phosphatases, see also Myotubularins; PTEN catalytic mechanism, 306–308 classification, 304–305 fluorescence resonance energy transfer studies of PTP1B, 223 mouse mammary gland studies, see Receptor protein tyrosine phosphatases oxidative regulation assay anaerobic work station, 310–311 buffers, 309–310 cell lysate preparation, 311

451

SUBJECT INDEX

in-gel assay, 313–314, 317 principles, 308–309 substrate radiolabeling, 312–313 overview, 305–306 prospects for study, 317–318 reversibility, 308 stimulus-induced oxidation, 314–316 phosphopeptide substrates, 5–6 signal transduction, 305–306 substrate specificity, 4–5 PTEN assay using fluorescent phosphoinositides incubation conditions, 50–51 materials, 50 principles, 49 substrate specificity findings, 51–52 thin-layer chromatography, 51 assay using malachite green advantages and limitations, 54–55 incubation conditions, 54–55 materials, 54 substrate specificity findings, 55–56 assay using pleckstrin homology domain/ lipid sensor complexes, 75 mutation in cancer, 44 purification of recombinant proteins Escherichia coli expression cell growth and induction, 46 materials, 45–46 nickel affinity chromatography, 46–47 mammalian cell expression advantages and limitations, 49 FLAG-tagged proteins in human embryonic kidney cells, 48–49 materials, 47–48 substrates, 43–44 PYST2 acute myelogenous leukemia overexpression, 103–104 DNA microarray analysis Atlas system, 104 bone marrow-derived leukocyte preparation, 105 considerations for clinical use, 111–113 differential gene expression analysis, 107–109 probe preparation, 106 verificaton with reverse transcriptase–polymerase chain reaction, 109–110

R Ras recruitment system, see Protein phosphatase-2A Receptor protein tyrosine phosphatases, see also CD45 dimerization chemical cross-linking bis[sulfosuccinimidyl]-suberate crosslinking, 226–227 overview, 225–226 coimmunoprecipitation studies, 236–238 comparison of techniques for study, 238–240 disulfide bond formation, 225 fluorescence resonance energy transfer studies green fluorescent protein variants, 231, 233, 235–236 instrumentation, 234–235 principles, 230–231, 233 receptor protein tyrosine phosphatase, 221–222 single cell spectral microscopy studies, 236 genetic cross-linking cysteine mutants, 227 detection with nonreducing polyacrylamide gel electrophoresis, 228–230 induction, 225 domains, 224 ligand regulation, 224–225 receptor protein tyrosine phosphatase-", transgenic mouse studies of mammary gland structure overview, 124–126 materials, 129 sample preparation, 129–131 staining of samples, 131–132 slide storage, 132 transparency of samples, 132 tumor promotion role, 125–126 Reg1p effects on yeast phosphoproteome extract preparation, 99–100 media, 98 mixed peptide sequencing, 101 mutant yeast strain, 96–97

452

SUBJECT INDEX

phosphatase assay anion-exchange chromatography, 101–102 automated assay, 102 substrate preparation, 101 phosphate radiolabeling of cell culture, 98–99 site-directed mutagenesis studies, 102 specific activity of ATP -phosphate determination, 100 two-dimensional gel electrophoresis, 100–101 protein phosphatase-1 interactions, 96–97 RNA interference, protein phosphatase knockdown in Drosophila advantages over isoform-specific inhibitors, 361, 371 applications, 371–372 complementary DNA preparation, 364–366 double-stranded RNA cell treatment, 367 production, 366 expression analysis reverse transcriptase-polymerase chain reaction, 369–371 Western blot, 368–369 posttranscriptional gene silencing, 362 protein phosphatase homolog identification, 364 regulatory subunit targeting, 363–364, 371 S2 cell line characteristics, 362–363 culture, 367 extract preparation, 367–368

S Saccharomyces cerevisiae gene deletion advantages in phosphatase genetic screening, 403, 405 deletion strain types, 404–405 duplication of libraries, 406–407 equipment for analysis, 405–406 Genome Deletion project, 404 genome-wide versus targeted screens, 407 glycogen storage gene screening genome-wide screening gene identification, 410–411

induction of glycogen synthesis, 407–409, 412 iodine staining, 408–409 kinase and phosphatase catalytic subunits, 410–411 glycogen phosphorylase phosphatase assay, 414–416 glycogen synthase assay, 413–414 glycogen synthase phosphatase/kinase assay, 412–413 lysate preparation for enzyme assays, 416–418 targeted screens with subsets of strains, 412 Superoxide dismutase, protection against calcineurin inactivation, 297–298

T Tetracycline-regulable promoters inducible expression of protein phosphatase-2C in human embryonic kidney cells cell viability assay, 341–342 clone isolation, 343 induction effects on cells apoptosis, 346–347 cell cycle arrest, 346–347 colony formation inhibition, 345–346 proliferation inhibition, 345–346 malachite green activity assay, 342 plasmids, 339–340 plating efficiency, 340 tetracycline-dependent induction characteristics, 343–345 T-Rex tetracycline-regulated expression system, 340 Western blotting, 341 yeast expression systems direct Tet system, 348–351 dual activator–repressor system, 351–353 overview, 347–348 plasmids for protein tagging, 353–354 reverse Tet system, 351 sit4 hal3 conditional mutant construction functional analysis of induced cells, 356–358 promoter substitution, 354–355 rationale, 354

SUBJECT INDEX

transformation and selection, 355–356 verification of correct integration, 356

W Western blot protein phosphatase-1 far-Western overlay assays of isoform protein–protein interactions, 164–167

453

phosphoprotein inhibitor analysis, 250–253 recombinant protein expression analysis, 329 protein phosphatase-2A complex formation analysis, 182, 201–202 protein phosphatase-2C, 341 RNA interference of protein phosphatase knockdown in Drosophila, 368–369