Dopamine-Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY™

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For further volumes: http://www.springer.com/series/7651

Dopamine Methods and Protocols

Edited by

Nadine Kabbani Department of Molecular Neuroscience, Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA

Editor Nadine Kabbani Department of Molecular Neuroscience Krasnow Institute for Advanced Study George Mason University Fairfax, VA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-62703-250-6 ISBN 978-1-62703-251-3 (eBook) DOI 10.1007/978-1-62703-251-3 Springer New York Heidelberg Dordrecht London Library of Congress Control Number: 2012950334 © Springer Science+Business Media, LLC 2013 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher’s location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover illustration: The image depicts a whole mount adult Drosophila brain triple-labeled with rabbit anti-GFP antibody (green), mouse anti-FasII (1D4) antibody (red) and DAPI (blue) which mark the dopaminergic neurons (revealed by genetically labeling with ple-GAL4,UAS-mCD8::GFP), axon tracts of the mushroom bodies and the central complex, and all brain cell nuclei, respectively. Note in the merged image the projections of dopamine neurons to areas of the mushroom body, the Drosophila center for learning and memory, and to the central complex, which contributes to the regulation of locomotion. (See Chapter 13.) Printed on acid-free paper Humana Press is a brand of Springer Springer is part of Springer Science+Business Media (www.springer.com)

Preface Dopamine, a catecholamine transmitter, plays a number of important physiological roles in the brain and body. Clues to dopamine’s role in motivation and learning have come from over 50 years of studies in laboratory animals, which have included rodents and nonhuman primates. In more recent years, studies on the role of dopamine in disease have opened new avenues of research and discovery. Genetic cloning has further enabled studies of dopamine in other species such as Drosophila melanogaster and Danio rerio. This edition of Methods in Molecular Biology brings together and provides detailed protocols on leading approaches in the study of dopamine within biological systems. In the brain, dopamine functions as a key neurotransmitter in regions such as the cortex and striatum. Dopamine is also an important modulator of ion balance in the kidney and adaptation to light in the retina. The many effects of dopamine on physiological systems and organs are dependent on a class of receptors, which are coupled to heterotrimeric G proteins. In mammals, five dopamine receptors (D1–D5) have been identified. A fundamental aspect of dopamine function is the localization of these receptors at the membrane, their interaction with signaling and regulatory molecules, and their ability to assemble into higherorder receptor oligomers (with dopamine and non-dopamine receptors) within cells. In many species, dopamine plays a major role in reward-driven learning. Indeed, almost every type of reward that has been studied increases dopamine transmission in the brain, and a variety of highly addictive drugs, including stimulants such as cocaine and methamphetamine, act directly on the dopamine system. Several prominent diseases of the nervous system are associated with dopamine. In particular, alterations in dopamine levels are intimately linked with the onset and progression of Parkinson’s disease, which results from the death of dopaminergic neurons within the substantia nigra. Schizophrenia, a disease of multiple genes and origins, has long been linked to dopamine imbalances within the striatum and cortex with the majority of classical antipsychotic drugs acting as antagonists at D2 receptors and many newer generation antipsychotics maintaining an effect on D4 receptors. This book is of interest to a range of scientists including cellular and molecular biologists, electrophysiologists, and pharmacologists. The chapters are intended for students and experts alike and for anyone interested in exploring the vast field of dopamine research. The book is divided into four parts based on methods: cellular/biochemical, imaging, genetics, and electrophysiology. Presented are chapters with step-by-step, clear, and precise instructions for various research procedures. This includes protocols for bioluminescence and fluorescence imaging, receptor immunoprecipitation and proteomic analysis, creation and characterization of a mouse model of Parkinson’s disease, real-time measurement of dopamine in the brain, and modeling signal transduction in silico. This volume is the product of contributions from experts and key figures within the field. I would like to thank the authors for their outstanding work and cooperation during the preparation of the volume. Specifically, I would like to thank the series editor, Professor John Walker, for his support during the assembly of this book. Nadine Kabbani, Ph.D.

v

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

BIOCHEMICAL, PROTEOMIC, AND COMPUTATIONAL TOOLS

1 Detection of Cell Surface Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . Jiping Xiao and Clare Bergson 2 Methods for the Study of Dopamine Receptors Within Lipid Rafts of Kidney Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peiying Yu, Van Anthony Villar, and Pedro A. Jose 3 Methods of Dopamine Research in Retina Cells . . . . . . . . . . . . . . . . . . . . . . . . . Ana Lucia Marques Ventura, Fernando Garcia de Mello, and Ricardo Augusto de Melo Reis 4 Capture of D2 Dopamine Receptor Signaling Complexes in Striatal Cells for Mass Spectrometry Proteomic Analysis . . . . . . . . . . . . . . . . . Nadine Kabbani and Jacob C. Nordman 5 Modeling Spatial Aspects of Intracellular Dopamine Signaling . . . . . . . . . . . . Kim T. Blackwell, Lane J. Wallace, BoHung Kim, Rodrigo F. Oliveira, and Wonryull Koh

PART II

v ix

3

15 25

43 61

CELLULAR IMAGING

6 A Biophysical Approach for the Study of Dopamine Receptor Oligomerization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sylwia Lukasiewicz, Agata Faron-Górecka, and Marta Dziedzicka-Wasylewska 7 Detection of Receptor Heteromers Involving Dopamine Receptors by the Sequential BRET-FRET Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gemma Navarro, Peter J. McCormick, Josefa Mallol, Carme Lluís, Rafael Franco, Antoni Cortés, Vicent Casadó, Enric I. Canela, and Sergi Ferré 8 BRET Approaches to Characterize Dopamine and TAAR1 Receptor Pharmacology and Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stefano Espinoza, Bernard Masri, Ali Salahpour, and Raul R. Gainetdinov 9 Dopaminergic Regulation of Dendritic Calcium: Fast Multisite Calcium Imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wen-Liang Zhou, Katerina D. Oikonomou, Shaina M. Short, and Srdjan D. Antic

vii

79

95

107

123

viii

Contents

PART III

GENETIC MANIPULATION IN CELLS AND ORGANISMS

10 Functional Analysis of Human D1 and D5 Dopaminergic G Protein-Coupled Receptors: Lessons from Mutagenesis of a Conserved Serine Residue in the Cytosolic End of Transmembrane Region 6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bianca Plouffe and Mario Tiberi 11 A Molecular Genetic Approach to Uncovering the Differential Functions of Dopamine D2 Receptor Isoforms . . . . . . . . . . . . . . . . . . . . . . . Yanyan Wang, Toshikuni Sasaoka, and Mai T. Dang 12 Genomic Strategies for the Identification of Dopamine Receptor Genes in Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wendy Boehmler, Jessica Petko, Victor A. Canfield, and Robert Levenson 13 Application of Cell-Specific Isolation to the Study of Dopamine Signaling in Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eswar Prasad R. Iyer, Srividya Chandramouli Iyer, and Daniel N. Cox

PART IV

141

181

201

215

ELECTROCHEMICAL, PHYSIOLOGICAL, AND BEHAVIORAL ANALYSIS

14 Regulation of Dopamine Transporter Expression by Neuronal Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shalini Padmanabhan, Thach Pham, and Balakrishna M. Prasad 15 Monitoring Axonal and Somatodendritic Dopamine Release Using Fast-Scan Cyclic Voltammetry in Brain Slices . . . . . . . . . . . . . . . . . . . . . . Jyoti C. Patel and Margaret E. Rice 16 Real-Time Chemical Measurements of Dopamine Release in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . James G. Roberts, Leyda Z. Lugo-Morales, Philip L. Loziuk, and Leslie A. Sombers 17 The MPTP/Probenecid Model of Progressive Parkinson’s Disease . . . . . . . . . Anna R. Carta, Ezio Carboni, and Saturnino Spiga Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

229

243

275

295 309

Contributors SRDJAN D. ANTIC • Department of Neuroscience, University of Connecticut Health Center, Farmington, CT, USA RICARDO AUGUSTO DE MELO REIS • Laboratory of Neurochemistry, Program in Neurobiology IBCCF, UFRJ, Rio de Janeiro, Brazil KIM T. BLACKWELL • The Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA CLARE BERGSON • Department of Pharmacology and Toxicology, Georgia Health Sciences University, Augusta, GA, USA WENDY BOEHMLER • Department of Biological Sciences, York College of Pennsylvania, York, PA, USA ENRIC I. CANELA • Department of Biochemistry and Molecular Biology, Faculty of Biology, University of Barcelona, Barcelona, Spain VICTOR A. CANFIELD • Department of Pharmacology, Penn State College of Medicine, Hershey, PA, USA EZIO CARBONI • Department of Biomedical Sciences, University of Cagliari, Cagliari, Italy ANNA R. CARTA • Department of Biomedical Sciences, University of Cagliari, Cagliari, Italy VICENT CASADÓ • Department of Biochemistry and Molecular Biology, Faculty of Biology, University of Barcelona, Barcelona, Spain ANTONI CORTÉS • Department of Biochemistry and Molecular Biology, Faculty of Biology, University of Barcelona, Barcelona, Spain DANIEL N. COX • School of Systems Biology, Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA MAI T. DANG • Department of Neurology, Hospital of University of Pennsylvania, Philadelphia, PA, USA MARTA DZIEDZICKA-WASYLEWSKA • Institute of Pharmacology, Polish Academy of Sciences, Kraków, Poland; Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, Krakow, Poland STEFANO ESPINOZA • Department of Neuroscience and Brain Technologies, Italian Institute of Technology, Genoa, Italy AGATA FARON-GÓRECKA • Institute of Pharmacology, Polish Academy of Sciences, Krakow, Poland SERGI FERRÉ • Department of Health and Human Services, Intramural Research Program, National Institute on Drug Abuse, National Institutes of Health, Baltimore, MD, USA RAFAEL FRANCO • Department of Biochemistry and Molecular Biology, University of Barcelona, Barcelona, Spain ix

x

Contributors

RAUL R. GAINETDINOV • Department of Neuroscience and Brain Technologies, Italian Institute of Technology, Genoa, Italy FERNANDO GARCIA DE MELLO • Laboratory of Neurochemistry, Program in Neurobiology IBCCF, UFRJ, Rio de Janeiro, Brazil ESWAR PRASAD R. IYER • School of Systems Biology, George Mason University, Manassas, VA, USA SRIVIDYA CHANDRAMOULI IYER • School of Systems Biology, George Mason University, Manassas, VA, USA PEDRO A. JOSE • Department of Pediatrics, Center for Molecular Physiology Research, Children’s National Medical Center, and School of Medicine, George Washington University, Washington, DC, USA NADINE KABBANI • Department of Molecular Neuroscience, Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA BOHUNG KIM • The Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA WONRYULL KOH • The Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA ROBERT LEVENSON • Department of Pharmacology, Penn State College of Medicine, Hershey, PA, USA CARME LLUÍS • Department of Biochemistry and Molecular Biology, University of Barcelona, Barcelona, Spain PHILIP L. LOZIUK • Department of Chemistry, North Carolina State University, Raleigh, NC, USA LEYDA Z. LUGO-MORALES • Department of Chemistry, North Carolina State University, Raleigh, NC, USA SYLWIA LUKASIEWICZ • Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, Krakow, Poland JOSEFA MALLOL • Department of Biochemistry and Molecular Biology, University of Barcelona, Barcelona, Spain BERNARD MASRI • Cancer Research Center of Toulouse, INSERM U1037 Université Paul Sabatier Toulouse III, CHU Rangueil, Toulouse, France PETER J. MCCORMICK • Department of Biochemistry and Molecular Biology, University of Barcelona, Barcelona, Spain GEMMA NAVARRO • Department of Biochemistry and Molecular Biology, University of Barcelona, Barcelona, Spain JACOB C. NORDMAN • Department of Molecular Neuroscience, Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA KATERINA D. OIKONOMOU • Department of Neuroscience, University of Connecticut Health Center, Farmington, CT, USA RODRIGO F. OLIVEIRA • The Krasnow Institute for Advanced Study, George Mason University, Fairfax, VA, USA SHALINI PADMANABHAN • Department of Pharmacology, Medical College of Georgia, Augusta, GA, USA JYOTI C. PATEL • Departments of Neurosurgery and Physiology & Neuroscience, New York University School of Medicine, New York, NY, USA

Contributors

xi

JESSICA PETKO • Department of Pharmacology, Penn State College of Medicine, Hershey, PA, USA THACH PHAM • General Surgery, Dwight D. Eisenhower Army Medical Center, Fort Gordon, GA, USA BIANCA PLOUFFE • Departments of Medicine/Cellular and Molecular Medicine/ Psychiatry, Ottawa Hospital Research Institute (Neuroscience Program), University of Ottawa, Ottawa, ON, Canada BALAKRISHNA M. PRASAD • Department of Pharmacology, Medical College of Georgia, Augusta, GA, USA;Clinical Investigation, Dwight D. Eisenhower Army Medical Center, Fort Gordon, GA, USA MARGARET E. RICE • Departments of Neurosurgery and Physiology & Neuroscience, New York University School of Medicine, New York, NY, USA JAMES G. ROBERTS • Department of Chemistry, North Carolina State University, Raleigh, NC, USA ALI SALAHPOUR • Department of Pharmacology and Toxicology, University of Toronto, Toronto, ON, Canada TOSHIKUNI SASAOKA • Department of Laboratory Animal Science, Kitasato University School of Medicine, Kanagawa, Japan SHAINA M. SHORT • Department of Neuroscience, University of Connecticut Health Center, Farmington, CT, USA LESLIE A. SOMBERS • Department of Chemistry, North Carolina State University, Raleigh, NC, USA SATURNINO SPIGA • Department of Life and Environmental Sciences, University of Cagliari, Cagliari, Italy MARIO TIBERI • Departments of Medicine/Cellular and Molecular Medicine/ Psychiatry, Ottawa Hospital Research Institute (Neuroscience Program), University of Ottawa, Ottawa, ON, Canada ANA LUCIA MARQUES VENTURA • Department of Neurobiology, Program in Neurosciences, Universidade Federal Fluminense, Niterói, Brazil VAN ANTHONY VILLAR • Department of Pediatrics, Center for Molecular Physiology Research, Children’s National Medical Center, and School of Medicine, George Washington University, Washington, DC, USA LANE J. WALLACE • College of Pharmacy, Ohio State University, Columbus, OH, USA YANYAN WANG • Department of Pharmacology, College of Medicine, Beckman Institute for Advanced Science and Technology, University of Illinois at UrbanaChampaign, Urbana, IL, USA JIPING XIAO • Cardiovascular Institute, University of Pennsylvania, Philadelphia, PA, USA PEIYING YU • Department of Pediatrics, Center for Molecular Physiology Research, Children’s National Medical Center, School of Medicine, George Washington University, Washington, DC, USA WEN-LIANG ZHOU • Department of Neuroscience, University of Connecticut Health Center, Farmington, CT, USA

Part I Biochemical, Proteomic, and Computational Tools

Chapter 1 Detection of Cell Surface Dopamine Receptors Jiping Xiao and Clare Bergson Abstract Dopamine receptors are a class of metabotropic G protein-coupled receptors. Plasma membrane expression is a key determinant of receptor signaling, and one that is regulated both by extra and intracellular cues. Abnormal dopamine receptor signaling is implicated in several neuropsychiatric disorders, including schizophrenia and attention deficit hyperactivity disorder, as well as drug abuse. Here, we describe in detail the application of two complementary applications of protein biotinylation and enzyme-linked immunoabsorbent assay (ELISA) for detecting and quantifying levels of dopamine receptors expressed on the cell surface. In the biotinylation method, cell surface receptors are labeled with Sulfo-NHS-biotin. The charge on the sulfonyl facilitates water solubility of the reactive biotin compound and prevents its diffusion across the plasma membrane. In the ELISA method, surface labeling is achieved with antibodies specific to extracellular epitopes on the receptors, and by fixing the cells without detergent such that the plasma membrane remains intact. Key words: Schizophrenia, ADHD, DAPI, Biotinylation, ELISA, Plasma membrane

1. Introduction Dopamine (DA) regulates movement, endocrine function, reward behavior, and memory processes by stimulating a family of five subtypes of G protein-coupled receptors (GPCRs) designated the D1 to D5 receptors (D1R-D5R). Disorders involving DA transmission include Parkinson’s disease (PD), as well as a number of neuropsychiatric illnesses including attention deficit hyperactivity disorder (ADHD) and schizophrenia. Several lines of evidence suggest that plasma membrane levels of D1Rs, in particular, are critically linked to working memory, an executive function impaired in schizophrenia (1–5). Since deficits in working memory and related executive functions are currently treatment-resistant, reagents which manipulate D1R cell surface expression could represent an effective therapeutic strategy. Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_1, © Springer Science+Business Media, LLC 2013

3

4

J. Xiao and C. Bergson

A number of factors have been discovered in the past 15 years or so which can regulate surface levels of D1Rs. For example, both hyper-and hypo-dopaminergic states produce alterations in surface D1Rs in vivo (6), and similar effects are observed in cells in culture with D1R agonists and antagonists (7, 8). Further, activation of glutamatergic N-methyl-D-aspartic acid (NMDA) receptors in neurons stimulates accumulation of D1Rs on synaptic membranes (9). This effect is regulated by physical interaction of NR1 NMDA receptor subunits with D1Rs (10). In addition, a variety of other mechanisms regulate D1R surface levels including endocytic recycling (11), receptor phosphorylation (12–14), as well as physical association with cytoskeletal proteins (15). Biotinylation and enzyme-linked immunoabsorbent assay (ELISA) offer a number of advantages for detecting and quantifying cell surface receptors. With either method, it is possible avoid the use of radioisotopes as is typically required in receptor ligand binding assays. Both methods are inherently quantitative. While immunofluorescent detection of DA receptor subtypes is also straightforward, quantification of surface levels by this method is not. The isolation of receptors on the cell surface devoid of contamination from other membrane compartments is troublesome with subcellular fractionation methods involving gradient centrifugation. However, the tools currently available for cell surface ELISA and biotinylation permit unambiguous assessment of receptors residing specifically on the plasma membrane. We provide detailed protocols for biotinylation and ELISA based-methods to quantify the cell surface levels of DA receptors under basal conditions and agonist stimulation. We use D1Rs to illustrate application of these approaches. However, these tools can be easily adapted for other DA receptor subtypes. In the biotinylation method, cell surface receptors are labeled with non-cleavable Sulfo-NHS-biotin. At neutral pH, the sulfo-NHS ester reacts rapidly with any primary amine-containing protein such that the biotin label is attached via a stable amide bond. As the sulfonyl group is charged, the compound shows good water-solubility, and poor ability to cross intact plasma membranes. As a result, labeling is restricted to the extracellular domains of proteins spanning the plasma membrane. The sulfo-NHS-biotin compound can also be used to study endogenous receptors in primary culture or in brain slices (16, 17). Cleavable biotinylation reagents such as sulfo-NHSS-S-biotin include a disulfide group positioned such that biotin label can be removed by treatment with reducing agents. These compounds are useful for quantifying agonist-stimulated receptor internalization as the biotin label can be stripped from receptors remaining on the cell surface prior to cell lysis (18). In the cells surface ELISA method, labeling is achieved by fixing the cells without detergent such that the plasma membrane remains intact.

1

Detecting Surface DA Receptors

5

Receptors can be detected with epitope or subtype specific primary antibodies, followed by enzyme-linked secondary antibodies, and exposure to chromogenic substrates.

2. Materials 2.1. Biotinylation of Cell Surface DA Receptors

1. HEK293 cells. 2. FLAG-D1R cells: this is an HEK293 cell line which stably expresses human D1Rs carrying a FLAG epitope tag inserted at the N-terminus of the receptor coding sequence. 3. HEK293 culture medium: Dulbecco’s Modified Eagle’s Medium (DMEM) (Sigma-Aldrich, St. Louis, MO) supplemented with 10% fetal bovine serum (FBS) (Sigma-Aldrich), 1% penicillin-streptomycin (Roche Diagnostics, Indianapolis, IN). 4. FLAG-D1R stable cell line medium: Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin, 450 μg/mL G418 (Invitrogen Life Technologies, Grand Island, NY). 5. PBS: 8.5 mM sodium phosphate, 1.5 mM potassium phosphate, 137 mM NaCl, pH 7.4. 6. Non-cleavable sulfo-NHS-Biotin (Pierce, Thermo Fisher Scientific, Rockford, IL). 7. 10 mM glycine in PBS. 8. Lysis buffer: 150 mM NaCl, 20 mM Tris–HCl, pH 7.5, 0.5% NP-40, 10% glycerol containing protease inhibitor cocktail (1 tablet/10 mL). 9. Protease inhibitor cocktail (Roche Diagnostics). 10. Sonic dismembrator (Fisher Scientific). 11. Streptavidin slurry (Pierce Biotechnology). 12. 1.5 M guanidine HCl. 13. 1× SDS loading buffer: 63 mM Tris–HCl, 10% glycerol, 2% SDS, 0.0025% Bromophenol Blue (Sigma), pH 6.8. 14. 7.5% SDS-PAGE gels. 15. Electrophoresis power supply, SDS-PAGE and protein gel transfer equipment (Bio-Rad, Hercules, CA). 16. 10× transfer buffer: 0.25 M Tris base, 2 M glycine. Dilute with double distilled H2O and add methanol to 20% for use. 17. PVDF membrane (Protran, GE Healthcare, Waukesha, WI). 18. Whatman 3 M paper.

6

J. Xiao and C. Bergson

19. TBS-T buffer: 250 mM Tris–HCl, pH 7.5, 1.5 M NaCl, 1% (v/v) Tween-20. Dilute from 10× TBS stock with ddH2O and add Tween-20. 20. Blocking buffer: 5% (w/v) nonfat dry milk in TBS-T. 21. Anti-FLAG M2 monoclonal antibody (mab). 22. Goat anti mouse-HRP antibody (Jackson ImmunoResearch, West Grove, PA). 23. ECL plus detection kit (Amersham, GE Healthcare, Waukesha, WI). 24. Kodak X-ray film (Kodak, Rochester, NY). 2.2. Detection of Cell Surface DA Receptors by ELISA

1. 24-well tissue culture plates. 2. 1 μg/mL of laminin: (BD bioscience, San Jose, CA). 3. 4% paraformaldehyde solution in PBS. 4. Non-permeabilizing blocking buffer: Tris-buffered saline (TBS) containing 5% nonfat dry milk and 5% normal goat serum. 5. Anti-FLAG monoclonal antibody M2 (Sigma-Aldrich). 6. Goat anti mouse antibody conjugated with horseradish peroxidase (HRP) (Jackson ImmunoResearch). 7. Tetramethylbenzidine (TMB) substrate (Pierce). 8. Stop buffer: 2 M H2SO4. 9. 20 nM DAPI solution (Invitrogen). 10. Standard fluorescence and absorbance multi-well plate reader or spectrophotometer.

3. Methods 3.1. Biotinylation of Cell Surface DA Receptors

1. Culture 1 × 106 FLAG-D1R cells and untransfected HEK293 cells in the appropriate medium in 10 cm dishes for 2 days at 37°C in a humidified 5% CO2 incubator (see Note 1). 2. Place dishes on ice. All of the following steps are performed on ice except if noted otherwise. 3. Wash cells three times briefly with 10 mL ice-cold PBS (see Note 2). 4. Add 5 mL 0.5 mg/mL cell-impermeable, non-cleavable sulfoNHS-Biotin in PBS to cells and incubate for 30 min on ice (see Note 3). 5. Wash cells three times with 10 mL glycine (10 mM) in PBS to quench the unbound biotin reagent (see Note 4).

1

Detecting Surface DA Receptors

7

6. After the last wash, wash cells once with ice-cold PBS. 7. Harvest cells in 1.5 mL of ice-cold PBS using a cell scraper, and collect cells by spinning the suspension at 800 × g for 5 min in a microcentrifuge. 8. Retain the pellet, and resuspend cells in 500 μL of lysis buffer. 9. Sonicate cells for 10 s on ice (see Note 5). 10. Incubate cells on ice for 30 min. 11. Centrifuge samples at 18,000 × g for 30 min at 4°C. 12. Keep the supernatant. 13. Add the Streptavidin slurry (100 μL) to the supernatant, and mix by end-over-end rotation for 2 h at 4°C. 14. Pellet the biotinylated protein bound streptavidin resin by centrifugation at 10,500 × g for 2 min (see Note 6). 15. Retain and wash the resin twice with lysis buffer, twice with 1.5 mL guanidine HCl, followed by two additional washes with lysis buffer (see Note 7). 16. Elute the bound proteins with 50 μL SDS-PAGE loading buffer by boiling the beads for 3–5 min at 100°C (see Note 8). 17. Load 20 μL of the samples and separate on a 7.5% SDS-PAGE gel by gel electrophoresis (see Note 9). 18. Transfer proteins to PVDF membrane in a gel transfer apparatus (see Note 10). 19. After transfer, wash the membrane three times (5 min each) with TBS-T on a rocking platform. 20. Incubate the membrane with 5% milk in TBS-T for 1–3 h at RT on rocking platform. 21. Discard block and add anti-FLAG M2 mab diluted 1:2,000 in blocking solution. Incubate the membrane at 4°C overnight (or for 2 h at RT) on a nutator (see Note 11). 22. Discard the primary antibody; wash the membrane three times (10 min each) with TBS-T at RT on rocking platform. 23. Incubate with the secondary antibody, rabbit anti mouse-HRP antibody (1:10,000) for 1 h at RT on a nutator or rocking platform. 24. Discard the secondary antibody and wash the membrane three times (10 min each) with TBS-T. 25. After the final wash, add 1 mL ECL reagent to cover the membrane, incubate for 1 min at RT. Wrap the membrane with saran wrap sheet, place in a developing cassette, and expose to X-ray film for a suitable time (typically, from 10 s to several minutes). Develop film in dark room (see Note 12) (Fig. 1).

8

J. Xiao and C. Bergson

Fig. 1. Detection of cell surface D1Rs by biotinylation. 48 h after plating untransfected HEK293 cells and FLAG-D1R cells expressing human D1Rs, cell surface proteins were labeled with sulfo-NHS-biotin. Biotinylated proteins were recovered by streptavidin resin. Cell surface Flag-D1Rs were detected by immunoblotting biotinylated proteins with HRP conjugated FLAG M2 antibody.

3.2. Detection of Cell Surface DA Receptors by ELISA

1. Coat a 24-well plate by incubating plate with 1 μg/mL of laminin at least 2 h (see Note 13). 2. Discard laminin solution and leave the plate in the hood for 30 min to dry. 3. Plate FLAG-D1R and untransfected HEK293 cells at 2 × 104 cells/well and culture cells for 2 days. 4. Wash the cells three times briefly with PBS; and then add 4% paraformaldehyde and incubate for 20 min at RT to fix cells (see Note 14). 5. Discard 4% paraformaldehyde and wash cells three times (5 min each) with PBS. 6. Block cells under non-permeabilizing conditions (PBS containing 5% nonfat dry milk, and 5% normal goat serum) for 1 h at RT (see Notes 15 and 16). 7. Discard blocking buffer and incubate cells with mouse antiFLAG M2 mab (1:250) in blocking buffer (PBS containing 5% nonfat dry milk, and 5% normal goat serum) for 2 h at RT (see Note 17). 8. Discard the primary antibody, and wash cells with PBS on a rocking platform (see Note 18). 9. Incubate cells with the HRP-conjugate secondary antibody (1:5,000) in blocking buffer for 1 h at RT (see Note 19).

1

Detecting Surface DA Receptors

9

Fig. 2. Determination of cell density using DAPI. HEK293 cells were plated at varying densities in a 24-well plate. After washing with PBS four times, 100 μL of DAPI (300 nM in PBS) was added to each well, and the plate was incubated for 5 min at RT. After the incubation, wells were rinsed several times with PBS. Samples were excited in 358 nm and the emission at 461 nm was recorded.

10. Discard the secondary antibody, and wash cells four times (10 min each) with PBS on a rocking platform (see Note 20). 11. Add 500 μL of TMB to each well, and incubate the plate for 15 min at RT (see Note 21). 12. Stop the reaction by adding 50 μL H2SO4 (2 M). The color will turn from blue into yellow. 13. Transfer 400 μL of the solution and measure the OD at 450 nm (see Note 22). 14. After HRP detection, add 100 μL DAPI (300 nM) for 5 min. Measure the DAPI fluorescence by exciting at 350 nm, and detecting at 470 nm. Cell number can be inferred from a standard curve of cells plated versus DAPI intensity as shown in Fig. 2.

4. Notes 1. Cells stably transfected with plasmids containing neomycin resistant markers such as the FLAG-D1R cells can also be maintained in HEK293 cell medium containing 250 μg/ mL G418. However, the purity of the cell line is better maintained with higher concentrations of G418 (e.g., 450 μg/mL). 2. The cells density should be about 90% confluence. Higher confluence will decrease the efficiency of cell surface protein biotinylation.

10

J. Xiao and C. Bergson

3. Biotinylation reagents are susceptible to hydrolysis so the biotin compound should be prepared just prior to use. Optimal results are obtained when the cell labeling solution is prepared from newly opened bottles. Alternatively, a stock biotinylation solution (100–200 mg/mL) could be prepared in DMSO, and aliquots stored at −20°C until use. 4. This step washes out protein in the culture medium which can be biotinylated as well as free sulfo-NHS-Biotin. Wash the cells gently since the plates are not coated with laminin, and HEK293 cells detach easily. 5. Keep the sonication probe moving slowly in the solution to avoid local fluxes in temperature, while keeping it submerged to avoid foaming. 6. Carefully pipette off supernatant. Alternatively, use a spin filter to retain the streptavidin resin in the upper reservoir. 7. The guanidine HCl wash helps reduce nonspecific binding to streptavidin. This step does not affect recovery of the avidin– biotin complexes as the high (10−15 M) affinity of avidin and biotin renders them fairly insensitive to extremes of pH, detergent, solvents, and temperature. 8. Boiling the samples is necessary to disrupt the non-covalent association of biotinylated proteins with the streptavidin beads. 9. Run mini-gels at 100 V for 10 min through stacking portion, and at 200 V for 40 min through the separating region of the gels. 10. Transfer can be carried out at 100 V for 1 h, or overnight at 25 V, both at 4°C. 11. The FLAG M2 mab is used to specifically detect D1Rs tagged with the FLAG epitope among all the biotinylated cell surface proteins eluted from the streptavidin slurry. If the DA receptor is not tagged, an alternative approach would be to immunoprecipitate with polyclonal receptor specific antibodies, and probe blots of material subsequently eluted from protein A/G resin with streptavidin-conjugated HRP. Resin washing conditions would need to be adjusted accordingly. 12. Biotinylation efficiency will vary from protein to protein. If labeling efficiency seems low as gauged from the intensity of bands in streptavidin recovered lanes versus lysate lanes, consider performing a second round of biotinylation before lysing the cells. Alternatively, increase the pH of the biotinylation solution (pH 8–9) to improve labeling by increasing the proportion of lysine ε-amino groups conjugated (19). 13. The laminin coating step helps decrease cell loss as the ELISA protocol involves several steps with extensive washes.

1

Detecting Surface DA Receptors

11

14. This step can also be carried out following treatment with agonists such as shown in Fig. 3a. Agonist-induced receptor internalization can be inferred from the ratio of receptor surface levels detected before and after agonist treatment (Fig. 3b). 15. This condition assures that anti-FLAG antibodies only bind the D1R on the cell surface. Nonspecific binding of antibody to either the plates or the cells will increase background signal. Blocking buffer composition and volume or blocking time might need to be adjusted to reduce background noise. We also suggest plating HEK293 cells which do not express FLAGD1Rs. Additionally, include FLAG-D1R negative control wells where the primary antibody is omitted and only the HRPsecondary is added; or where FLAG mab is followed by unconjugated secondary antibodies. Negligible TMB signals coming from such negative control samples are necessary to validate the results from the experimental samples. 16. An alternative strategy could be the use of D1R subtype selective antibodies directed at an extracellular epitope.

HRP (a.u./cell number)

a

15

Vehicle SKF81297

10 5 0

HEK293

FLAG-D1R

b Internaliztion Ratio (%)

75 50 25 0 –25 HEK293

FLAG-D1R

Fig. 3. Cells surface D1Rs measured 15 min after addition of D1R agonist SKF81297 (10 nM) or vehicle using the ELISA method. (a) Surface D1Rs by cell surface ELISA assay. Cells were fixed under non-permeabilizing conditions, and cell surface D1Rs detected with anti-FLAG and HRP conjugated secondary antibodies, followed by ELISA using the TMB substrate for HRP. Cell numbers were determined by DAPI staining. (b) The “endocytosis ratio” was determined by the (surface D1Rs in vehicle treated cells- surface D1Rs treated with SKF for 15 min/surface D1Rs in vehicle treated cells). The bar graphs show the mean ± SEM of three independent experiments each including four replicates per group.

12

J. Xiao and C. Bergson

17. Be cautious, no detergent! 18. If background signal is high (e.g., non-transfected HEK293 and FLAG-D1R cells give equivalent signals), wash cells six times, or decrease the incubation time with the anti-FLAG mab from 2 h to 1 h. 19. Be cautious! No detergent! 20. If background is high, wash cells six times. 21. Blue color should appear after 15 min. TMB is a chromogenic HRP substrate which absorbs at 450 nm. However, chemiluminescent and fluorescent HRP substrates are also available. 22. Remember to keep the plate for cell number counting. References 1. Vijayraghavan S, Wang M, Birnbaum SG, Williams GV, Arnsten AFT (2007) Inverted-U dopamine D1 receptor actions on prefrontal neurons engaged in working memory. Nat Neurosci 10:376–384 2. Zahry J, Taylor JR, Mathew RG, Arnsten AF (1997) Supranormal stimulation of D1 dopamine receptors in the rodent prefrontal cortex impairs spatial working memory performance. J Neurosci 17:8528–8535 3. McNab F, Varrone A, Farde L, Jucaite A, Bystritsky P, Forssberg H, Klingberg T (2009) Changes in cortical dopamine D1 receptor binding associated with cognitive training. Science 323:800–802 4. Abi-Dargham A, Mawlawi O, Lombardo I, Gil R, Martinez D, Huang Y, Hwang DR, Keilp J, Kochan L, Van Heertum R, Gorman JM, Laruelle M (2002) Prefrontal dopamine D1 receptors and working memory in schizophrenia. J Neurosci 22:3708–3719 5. Castner SA, Williams GV, Goldman-Rakic PS (2000) Reversal of anti-psychotic-induced working memory deficits by short-term dopamine D1 receptor stimulation. Science 287:2020–2022 6. Dumartin B, Jaber M, Gonon F, Caron MG, Giros B, Bloch B (2000) Dopamine tone regulates D1 receptor trafficking and delivery in striatal neurons in dopamine transporterdeficient mice. Proc Natl Acad Sci U S A 97:1879–1884 7. Martin-Negrier M, Charron G, Bloch B (2000) Agonist stimulation provokes dendritic and axonal dopamine D(1) receptor redistribution in primary cultures of striatal neurons. Neuroscience 99:257–266

8. Brismar H, Asghar M, Carey RM, Greengard P, Aperia A (1998) Dopamine-induced recruitment of dopamine D1 receptors to the plasma membrane. Proc Natl Acad Sci U S A 95:5573–5578 9. Scott L, Kruse MS, Forssberg H, Brismar H, Greengard P, Aperia A (2002) Selective upregulation of dopamine D1 receptors in dendritic spines by NMDA receptor activation. Proc Natl Acad Sci U S A 99:1661–1664 10. Pei L, Lee FJS, Moszczynska A, Vukusic B, Liu F (2004) Regulation of dopamine D1 receptor function by physical interaction with the NMDA receptors. J Neurosci 24:1149–1158 11. Vargas GA, Von Zastrow M (2004) Identification of a novel endocytic recycling signal in the D1 dopamine receptor. J Biol Chem 279:37461–37469 12. Yu P, Asico LD, Luo Y, Andrews P, Eisner GM, Hopfer U, Felder RA, Jose PA (2006) D1 dopamine receptor hyperphosphorylation in renal proximal tubules in hypertension. Kidney Int 70:1072–1079 13. Kim OJ, Gardner BR, Williams DB, Marinec PS, Cabrera DM, Peters JD, Mak CC, Kim KM, Sibley DR (2004) The role of phosphorylation in D1 dopamine receptor desensitization: evidence for a novel mechanism of arrestin association. J Biol Chem 279:7999–8010 14. Lamey M, Thompson M, Varghese G, Chi H, Sawzdargo M, George SR, O’Dowd BF (2002) Distinct residues in the carboxyl tail mediate agonist-induced desensitization and internalization of the human dopamine D1 receptor. J Biol Chem 277:9415–9421 15. Kim OJ, Ariano MA, Lazzarini RA, Levine MS, Sibley DR (2002) Neurofilament-M interacts

1 with the D1 dopamine receptor to regulate cell surface expression and desensitization. J Neurosci 22:5920–5930 16. Holman D, Henley JM (2007) A novel method for monitoring the cell surface expression of heteromeric protein complexes in dispersed neurons and acute hippocampal slices. J Neurosci Methods 160:302–308 17. Mao SC, Hsiao YH, Gean PW (2006) Extinction training in conjunction with a partial agonist of the glycine site on the NMDA

Detecting Surface DA Receptors

13

receptor erases memory trace. J Neurosci 26:8892–8899 18. Ali MK, Bergson C (2003) Elevated intracellular calcium triggers recruitment of the receptor cross-talk accessory protein calcyon to the plasma membrane. J Biol Chem 278: 51654–51663 19. Gottardi CJ, Dunbar LA, Caplan MJ (1995) Biotinylation and assessment of membrane polarity: caveats and methodological concerns. Am J Physiol 268:F285–F295

Chapter 2 Methods for the Study of Dopamine Receptors Within Lipid Rafts of Kidney Cells Peiying Yu, Van Anthony Villar, and Pedro A. Jose Abstract There is increasing evidence that G protein-coupled receptor (GPCR) signaling is regulated in lipid raft microdomains. GPCRs and GPCR-signaling molecules, including G proteins and protein kinases, have been reported to compartmentalize in these microdomains. Dopamine D1-like receptors (D1R and D5R) belong to a family of GPCRs that are important in the regulation of renal function. These receptors are not only localized and regulated in caveolae that contains caveolin-1 but are also distributed in noncaveolar lipid rafts which do not contain caveolin-1. This chapter describes detergent- and non-detergentbased methods to obtain lipid raft fractions from renal proximal tubule cells. Key words: Lipid rafts, Caveolae, Membrane microdomains, Dopamine receptor

1. Introduction Dopamine receptors belong to the α group of the rhodopsin-like family of G protein-coupled receptors (GPCRs) and are classified into two subfamilies depending on their effect on adenylyl cyclase activity. The D1-like receptors (D1R and D5R) stimulate while the D2-like receptors (D2R, D3R and D4R) inhibit adenylyl cyclase activity (1). Dopamine D1-like receptors have been implicated in the modulation of various neural processes, including learning, memory, reward, and motor activity (2, 3), and in the regulation of blood pressure by actions on the adrenergic nervous system, hormone secretion, and epithelial ion transport (1). Lipid rafts are membrane microdomains composed of cholesterol, sphingolipids, glycosylphosphatidylinositol-linked (GPIlinked) proteins, and other proteins such as caveolin (4–6).

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_2, © Springer Science+Business Media, LLC 2013

15

16

P. Yu et al.

Caveolae and lipid rafts have been implicated to play a role in cellular processes like membrane sorting, receptor trafficking, signal transduction, and cell adhesion. Lipid rafts serve as signaling platforms for several signaling molecules such as G protein subunits, enzymes, and adaptor proteins that play important roles in signal transduction in a variety of mammalian cells (4–7). Lipid rafts are characterized by their relative insolubility in nonionic detergents at 4°C and light buoyant density on sucrose gradient (4, 7). Among the lipid rafts, caveolae are the best characterized, are localized on cell surface invaginations, and are formed by polymerization of caveolin proteins with cholesterol (5–9). Three caveolin genes encode the caveolin proteins, namely, caveolin-1, caveolin-2, and caveolin-3. Caveolin-1 has been used as a marker protein for caveolae (5–9). There are several other markers for lipid rafts, such as flotillin-1, CD55, and alkaline phosphatase (10, 11). Flotillin-1 has been used as a lipid raft marker protein in cells that do not contain caveolae, i.e., blood cells (11), neural cells (12), and rat kidney tubule cells (13, 14). We have reported that there are non-caveolar lipid rafts in human embryonic kidney cells since these cells are devoid of measurable caveolin-1 (13). There are several ways to prepare lipid rafts using detergent or detergent-free methods. Detergent-free methods have been developed to isolate lipid rafts (7–9). Schnitzer et al. described a detergent-free method to isolate lipid rafts from rat lung vasculature by perfusion with a suspension of cationic colloidal silica particles, which is a good method for in vivo studies (7). The methods reported by Smart et al. and Song et al. are also detergent-free and have been extensively used to isolate lipid rafts membranes from a variety of cells (8, 9). The method by Smart et al. allows the isolation of a more purified fraction of lipid rafts because it uses purified plasma membranes rather than total cell lysates (8), in contrast to the method by Song et al. which uses total cell lysates (9). The results obtained using non-detergent extracted rafts are more reproducible and generate a greater fraction of inner leaflet-membrane lipids than detergent-extracted rafts (15). The samples obtained by detergent methods have been termed detergent-resistant membranes (DRMs) or detergent-insoluble fraction (7, 10, 15). The nonionic detergents, e.g., Triton X-100, are commonly used to purify lipid raft fractions (7, 10, 16–18). However, different detergents may yield different lipid raft components because different types of raft proteins have varying degrees of resistance to extraction by specific detergents (10, 15). Differences between detergent and non-detergent methods for the preparation of lipid rafts may be responsible for the observed variability in the lipid composition of the isolated rafts (7, 10, 15). We now describe non-detergent and detergent methods to isolate lipid raft membranes.

2

Dopamine Receptors in Lipid Microdomains

17

2. Materials 2.1. Cell Culture

1. HEK-293 cells heterologously expressing human D1 receptor (HEK-hD1) that have been previously characterized (13). 2. Prepare complete medium for cell culture by adding 5 mL of Pen/Strep and 50 mL of FBS to 500 mL of DMEM/F12 medium. 3. Grow cells in 150-mm dishes with complete medium in a humidified incubator in 5% CO2 and 95% air.

2.2. Sucrose Gradient Centrifugation

All stock solutions are prepared in distilled water at room temperature and stored at 4°C. 1. 250 mM 2-N-morpholino ethanesulfonic acid (Mes) stock solution, pH » 6.7–6.8. 2. 1.5 M sodium chloride (NaCl) stock solution. 3. Mes-buffered saline (MBS) solution: 25 mM Mes and 150 mM NaCl, pH » 6.7–6.8. 4. 500 mM sodium carbonate, pH 11 (pH need not be adjusted). 5. 5%, 35%, and 80% sucrose solutions in MBS buffer (see Note 1). 6. Add protease inhibitor cocktail to the sodium carbonate and sucrose solutions. 7. Protein assay using BCA kit (Pierce Thermo Scientific (Rockford, IL)). 8. Phosphate-buffered saline (PBS). 9. D1-like receptor agonist fenoldopam (1 mM) (Sigma-Aldrich, St. Louis, MO) stock solution, aliquoted into small volumes (50 μL/aliquot), protected from light, and stored at −20°C. Antioxidants are needed for prolonged incubation of dopamine and dopamine agonists. 10. Prepare fresh solution of methyl-β-cyclodextrin (βCD) (Sigma) (2%) in DMEM/F12 serum-free medium (SFM) at room temperature. 11. Cholesterol-βCD solution (Sigma) for cholesterol repletion experiment: (a) Dissolve cholesterol (20 mg/mL) in ethanol by sonication. (b) Dissolve βCD (2%) in DMEM/F12 SFM. (c) Prepare cholesterol-βCD solution by adding 20 μL of cholesterol solution to 10 mL cyclodextrin solution, mix by vortexing, and incubating the cholesterol-βCD solution at 40°C for 30 min (see Note 2).

18

P. Yu et al.

Table 1 Prepare varying concentrations of OptiPrep solutions Solutions (total 5 mL)

30%

20%

10%

5%

50% OptiPrep (mL)

3.0

2.0

1.0

0.5

MBSTS (mL)

2.0

3.0

4.0

4.5

12. 50% OptiPrep stock solution: 45 mL of 60% OptiPrep mixed with 9 mL of OptiPrep diluent. 13. MBSTS buffer: MBS with 0.5% (v/v), Triton X-100 in 10% sucrose, or other nonionic detergents, e.g., β-octyl glucoside, CHAPS, deoxycholate, Lubrol WX, Lubrol PX, Brij 58, Brij 96, Brij 98 (Sigma), as needed (see Note 3). 14. Prepare 5% and 30% gradient OptiPrep solutions according to Table 1 using 50% OptiPrep stock solution and MBSTS buffer. 15. 6× sample buffer: 7.5 mL of 0.5 M Tris–HCl, pH 6.8, 1 g of SDS powder, 3.6 mL of 100% glycerol, 2 mg of bromphenol blue, 1 g of dithiothreitol in a total 10 mL volume with distilled water. 2.3. Western Blot for Lipid Raft Proteins

1. Nitrocellulose membranes (0.2 μm pore size) (Invitrogen Life Technologies, Grand Island, NY). 2. Pre-stained molecular weight markers (Invitrogen Life Technologies). 3. Vertical midi-format electrophoresis cell, which should include a buffer tank and lid with power cables. 4. Criterion Precast Gels: 4–20% polyacrylamide gel, 26-well gel (Bio-Rad, Hercules, CA) or 8–16% polyacrylamide gel, 15-well gel (Invitrogen). 5. 10× Tris/Glycine/SDS stock buffer, to make 1× running buffer. 6. 10× Tris/Glycine buffer, to make 1× transfer buffer containing 20% methanol. 7. 10× PBS-tween-20 buffer, to make 1× washing buffer. 8. 0.1% Amido Black, 45% MEOH, 10% acetic acid in distilled water. 9. 0.1% Ponceau S solution in 5% acetic acid (remove the dye from the membrane by several washes with distilled water). 10. Primary antibodies and secondary antibodies conjugated to horseradish peroxidase. 11. Enhanced chemiluminescence (ECL) Western blotting detection reagents (GE Healthcare, Waukesha, WI).

2

Dopamine Receptors in Lipid Microdomains

19

3. Methods 3.1. Preparation of Lipid Raft Fraction with Non-detergent Method

Caveolae and lipid raft proteins are resistant to the solubilizing actions of detergents and some non-detergent reagents, such as sodium carbonate. Therefore, the raft proteins and membranes can be prepared using these detergents or reagents for sucrose gradient centrifugation. To prepare caveolar-enriched or non-caveolar lipid rafts, one can use the detergent-free sucrose gradient centrifugation protocol according to Song et al. (9) with slight modifications (13). This method can be adopted for all mammalian cells and tissues, including those that do not express caveolin-1 (13, 14), i.e., HEK-293 cells. For example, rat renal proximal tubule cells, used as an example, do not express caveolin-1 and therefore do not have caveolae (13, 14). We suggest using at least two 150-mm dishes for a single preparation. All experiments are carried out at 4°C except for cell culture and cell treatments. 1. Collect cell pellets. Culture cells in 150-mm dishes with DMEM/F12 complete medium at 37°C until the cells reach 95% confluence. Remove the cell culture medium and wash the cells twice with PBS. Then, starve the cells in DMEM/F12SFM for 1–2 h at 37°C. Treat the cells with vehicle or drugs (e.g., fenoldopam, 2% βCD, cholesterol–cyclodextrin solution) at 37°C for 1 h. Wash the cells once with cold PBS or cold DMEM/F12-SFM. Scrape the cells into a 15 mL tube containing cold PBS. Pellet the cells by centrifugation at 2,000 × g for 5 min. Discard the supernatant to obtain the cell pellet. 2. Prepare cell homogenates. Add 1.5 mL of 500 mM sodium carbonate to the cell pellet and mix by vortexing. Place the 15 mL tube containing the cells on ice and homogenize the cell suspension using a Dounce homogenizer (10 strokes), a Teflon polytron (three 10-s bursts), and a tip sonicator (three 30-s bursts). The homogenization steps are carried out on ice (see Note 4). Add 1.5 mL of 80% sucrose (final volume 3 mL, sucrose concentration, 40%) and mix the homogenate by vortexing (three 30 s bursts) and sonicating (three 30 s bursts) on ice. Determine the protein concentrations by BCA kit (OD 570). 3. Prepare a discontinuous sucrose gradient. Place equal amounts of cell homogenates (3 mL) into the bottom of each precooled 12 mL ultracentrifuge tubes and overlay 4.5 mL of 35% sucrose and 4.5 mL of 5% sucrose to each tube. The ultracentrifuge tubes should be balanced when placed and positioned in SW-41 buckets. 4. Centrifuge the tubes containing the cell homogenates at 180,000 × g (38,000 rpm) for 16 h at 4°C in a Beckman SW41 rotor (see Note 5).

20

P. Yu et al.

5. Remove the tubes from the bucket at the end of the ultracentrifugation step. A light-scattering band that contains caveolae-enriched lipid raft membranes is seen at the interface of the 5–35% sucrose gradient. Carefully collect twelve 1 mL fractions by pipetting 1 mL starting from the top of the ultracentrifuge tube and transfer the fractions into the pre-labeled 1.5 mL microcentrifuge tubes (see Note 6). The light-scattering band is located at the 3rd to 5th fractions from the top, with the peak at the 4th fraction. 6. Prepare samples for immunoblotting. Transfer 0.5 mL aliquots from each fraction into other pre-labeled 1.5 mL microcentrifuge tubes. Add 0.1 mL of 6× sample buffer to each sample. Vortex each tube until dye and samples are mixed well and put the tubes in boiling water for 5 min. The samples for immunoblotting can be saved at −20°C until use. The rest of the fractionated samples not mixed with the 6× sample buffer are saved at −80°C (see Note 7). 3.2. Preparation of Lipid Raft Fraction with Detergent Method

Detergent resistance or detergent insolubility results from the segregation of integral or membrane-associated proteins into cholesterol- and glycosphingolipid-enriched membrane microdomains termed lipid rafts. The nonionic detergents such as Triton X-100, β-octyl glucoside, CHAPS, deoxycholate, Lubrol WX, Lubrol PX, Brij 58, Brij 96 and Brij 98 have been used to purify lipid raft fractions (7, 10, 16–18). However, different detergents may yield different lipid raft components because different types of raft proteins have varying degrees of resistance to different detergents (10, 15). 1. Collect cell pellets (see Subheading 3.1, step 1) (One 150-mm dish for one preparation). 2. Prepare cell extract on ice for 30 min in 0.3 mL cold MBSTS (0.5% Triton X-100 and protease inhibitors) by pushing the cell suspension through a 25-gauge needle, ten times (cell pellet volume is about 0.1 mL/dish and the total cell lysate volume is about 0.4 mL). Adjust the cell extract (0.4 mL) to 40% OptiPrep by adding 0.8 mL of cold 60% OptiPrep, mix the cell extract by vortexing. Determine the protein concentrations using a BCA kit (OD 570). The total protein amount should be the same for all centrifuge tubes with the same volume (1 mL). 3. Prepare a discontinuous OptiPrep gradient. Load 1 mL of the cell extract into the bottom of precooled 5 mL ultracentrifuge tubes. Overlay with 1 mL of each 30%, 25%, 20%, and 0% OptiPrep solutions in MBSTS buffer, as prepared in Table 1 (see Note 8). 4. Ultracentrifuge the OptiPrep gradient solutions at 175,000 × g (42,000 rpm) at 4°C for 4 h in Beckman SW 50.1 rotor. Other

2

Dopamine Receptors in Lipid Microdomains

21

rotors can be used such as SW 55 (4 h at 170,000 × g), TLS55 rotor (2.5 h at 250,000 × g). However, the equivalent g-force and centrifugation time should be adjusted according to the rotor type. Label 1.5 mL microcentrifuge tubes for the next step. 5. Carefully remove the ultracentrifuge tubes. Collect 0.5 mL fractions from top to bottom and prepare for immunoblotting, as in Subheading 3.1. 3.3. Immunoblotting to Analyze Lipid Raft Proteins

Western blot allows the identification and analysis of the lipid raft proteins. In general, one should first identify where the peak of lipid raft fractions is located using lipid raft marker proteins such as caveolin-1, caveolin-3, or flotillin-1. To compare the effect of drugs on lipid raft protein expression, 4–20% Criterion Precast Gradient Gel with 26 wells per gel is recommended. All the steps are carried out at room temperature. 1. Run gels. Pre-warm the sucrose gradient samples in a water bath at 37°C. Mix the samples completely by vortexing (there should be no precipitate at the bottom of the tubes). Load the samples and molecular weight marker into a 4–20% Criterion Precast gradient gel. Run the gel with running buffer at 120 V for about 2 h. Stop the electrophoresis when the dye migrates to 0.5–1.0 cm above the bottom edge of the gel. 2. Transfer the proteins from gels onto nitrocellulose membranes. Prepare the sandwich of gels and nitrocellulose membranes in transfer buffer and place the sandwich into semidry transfer equipment and start the transfer at 0.24 mA at constant current for 60–90 min. 3. Block the membranes. Rinse the membranes twice with distilled water after transfer. Verify the protein loading by staining the membranes with 0.1% Ponceau S solution or 0.1% Amido Black solution for 10 s and washing the sheets with distilled water. The stained sheets can be scanned to record the protein loading information. Block the membranes for 1 h in blocking buffer (5% nonfat dry milk in PBST washing buffer). 4. Perform the immunoblotting. Incubate the blocked membranes overnight at 4°C with primary antibody diluted in blocking buffer (see Note 9). Remove the primary antibody following the overnight incubation and wash the membranes 3× with wash buffer. Incubate the membranes for 1 h with secondary antibody diluted in blocking buffer. 5. Develop the film. Incubate the membranes for 1 min with ECL reagent after washing with wash buffer 3×. Visualize the immunoreactive bands by autoradiography (see Note 10).

22

P. Yu et al.

4. Notes 1. The sucrose solutions (5%, 35%, and 80%) are prepared in MBS buffer, pH 6.8 (13) rather then in sodium carbonate solution (pH 11) (9). This results in a sample pH near 7.0 instead of pH 11. This may be beneficial to most of the enzyme proteins. 2. For cholesterol depletion experiment, the cells are incubated with methyl-β-cyclodextrin (βCD) (2%) for 1 h at 37°C. However, methyl-α-cyclodextrin has been recommended as a negative control (19). For cholesterol repletion experiments, βCD and cholesterol complex is used (Subheading 3.1, step 3). There is a commercially available cholesterol-cyclodextrin complex (SIGMA #C4951). However, the complex can also be prepared as described above (Subheading 2.2, item 11). An inactive analog of cholesterol (cholestane-3β, 5α, 6β-triol) has been suggested as a control (20). 3. Extraction using nonionic detergents. In general, Triton X-100 or CHAPS can solubilize the membranes that are extremely enriched in cholesterol and sphingolipids (15). Different raft proteins have different sensitivities to the different detergents. For example, even in the same cell type, different GPI-anchored proteins which associate with lipid rafts can be distinguished based on their sensitivity to solubilization in nonionic detergents. A good example of this is the prion protein, a GPIlinked protein, which was found only in non-raft fractions after solubilization in 0.5%Brij 96, but was distributed evenly between the raft and non-raft fractions when 0.5% Triton X-100 was used (21). 4. To avoid loss of cell samples during homogenization, we use a tip sonicator (five 20-s bursts, with a 2-min interval after each burst) instead of using Dounce homogenizer and Teflon polytron. All sonication steps should be performed with the test tubes on ice. This homogenization procedure can be used for cells but not for tissues. 5. The SW40 or SW41 rotors can be used for sucrose gradient centrifugation. The speed of the centrifugation is specific for each rotor. In general, the rotor speeds are 38,000 rpm (18,000 × g)/16 h for SW41 rotor and 36,000 rpm (16,000 × g)/18 h for SW40 rotor. 6. Collect twelve 1 mL fractions starting from the bottom by inserting a fine plastic tube in to bottom of the centrifuge tube and withdrawing one 1 mL each time using a 2 mL syringe, or a peristaltic pump.

2

Dopamine Receptors in Lipid Microdomains

23

7. The sucrose gradient samples with pH 6.8–7.0 can be stored at −80°C for enzyme assays, e.g., adenylyl cyclase assay. There are many ways to concentrate the fraction samples such as speedvac concentrator or by precipitation with 10% trichloroacetic acid (TCA). The membranes from lower sucrose gradient fractions can also be concentrated by three-fold dilution of the samples with MBS and pelleted by centrifugation at 20,000 × g for 30 min. 8. The OptiPrep discontinuous gradient can be made by overlaying 3 mL of 30% and 0.5 mL of 5% Optiprep solutions (16), or by overlaying 1 mL of 30%, 1 mL of 25%, 1 mL of 20%, and 1 mL of 0% OptiPrep solutions (17). However, it is best to prepare an OptiPrep continuous gradient using a machine for preparing gradients (Bio-Rad) or by overlaying 0.8 mL of each 30%, 25%, 20%, 15%, and 0% OptiPrep solutions and precentrifugation at 175,000 × g (42,000 rpm) at 4°C for 2 h in Beckman SW 50.1 rotor. Subsequently, load the protein samples at the bottom of the continuous OptiPrep gradient tube. 9. The primary antibody can be diluted in an antibody diluting solution (Invitrogen), and the diluted primary antibody can be collected and saved at −20°C for subsequent usage. The primary antibody diluted in 5% milk buffer is not recommended for storage. 10. To visualize the immunoreactive bands, the use of Licor (Odyssey) is recommended. When using Licor, the membranes should be blocked using a special blocking solution, such as casein or BSA blocking buffers (Bio-Rad), and the appropriate secondary antibody conjugated to IRDye® infrared dyes (in PBS casein buffer). The immunoreactive bands are visualized by scanning the membrane using Licor.

Acknowledgments These studies were supported in part by grants from the National Institutes of Health (HL023081, HL074940, DK039308, HL068686, and HL092196). References 1. Jose PA, Eisner GM, Felder RA (2002) Role of dopamine receptors in the kidney in the regulation of blood pressure. Curr Opin Nephrol Hypertens 11:87–92 2. Holmes A, Lachowicz JE, Sibley DR (2004) Phenotypic analysis of dopamine receptor knockout mice; recent insights into the

functional specificity of dopamine receptor subtypes. Neuropharmacology 47: 1117–1134 3. Kong MMC, Hasbi A, Mattocks M, Fan T, O’Dowd BF, George SR (2007) Regulation of D1 dopamine receptor trafficking and signaling by caveolin-1. Mol Pharmacol 72:1157–1170

24

P. Yu et al.

4. Simons K, Ikonen E (1997) Functional rafts in cell membranes. Nature 387:569–572 5. Insel PA, Head BP, Ostrom RS, Patel HH, Swaney JS, Tang CM, Roth DM (2005) Caveolae and lipid rafts: G protein-coupled receptor signaling microdomains in cardiac myocytes. Ann N Y Acad Sci 1047:166–172 6. Lingwood D, Simons K (2010) Lipid rafts as a membrane-organizing principle. Science 327:46–50 7. Schnitzer JE, McIntosh DP, Dvorak AM, Liu J, Oh P (1995) Separation of caveolae from associated microdomains of GPI-anchored proteins. Science 269:1435–1439 8. Smart EJ, Ying YS, Mineo C, Anderson RG (1995) A detergent-free method for purifying caveolae membrane from tissue culture cells. Proc Natl Acad Sci U S A 92:10104–10108 9. Song KS, Li S, Okamoto T, Quilliam LA, Sargiacomo M, Lisanti MP (1996) Co-purification and direct interaction of Ras with caveolin, an integral membrane protein of caveolae microdomains. Detergent-free purification of caveolae microdomains. J Biol Chem 271:9690–9697 10. Foster LJ, De Hoog CL, Mann M (2003) Unbiased quantitative proteomics of lipid rafts reveals high specificity for signaling factors. Proc Natl Acad Sci U S A 100:5813–5818 11. Salzer U, Prohaska R (2001) Stomatin, flotillin-1, and flotillin-2 are major integral proteins of erythrocyte lipid rafts. Blood 97:1141–1143 12. Huang P, Xu W, Yoon SI, Chen C, Chong PL, Liu-Chen LY (2007) Cholesterol reduction by methyl-beta-cyclodextrin attenuates the delta opioid receptor-mediated signaling in neuronal cells but enhances it in non-neuronal cells. Biochem Pharmacol 73:534–549

13. Yu P, Yang Z, Jones JE, Wang Z, Owens SA, Mueller SC, Felder RA, Jose PA (2004) D1 dopamine receptor signaling involves caveolin-2 in HEK-293 cells. Kidney Int 66:2167–2180 14. Breton S, Lisanti MP, Tyszkowski R, McLaughlin M, Brown D (1998) Basolateral distribution of caveolin-1 in the kidney. Absence from H+-ATPase-coated endocytic vesicles in intercalated cells. J Histochem Cytochem 46:205–214 15. Pike LJ (2004) Lipid rafts: heterogeneity on the high seas. Biochem J 378:281–292 16. Waheed AA, Jones TL (2002) Hsp90 interactions and acylation target the G protein Gα12 but not Gα13 to lipid rafts. J Biol Chem 277:32409–32412 17. Verkade P, Harder T, Lafont F, Simons K (2000) Induction of caveolae in the apical plasma membrane of Madin-Darby canine kidney cells. J Cell Biol 148:727–739 18. Macdonald JL, Pike LJ (2005) A simplified method for the preparation of detergent-free lipid rafts. J Lipid Res 46:1061–1067 19. Vial C, Evans RJ (2005) Disruption of lipid rafts inhibits P2X1 receptor-mediated currents and arterial vasoconstriction. J Biol Chem 280:30705–30711 20. Murtazina R, Kovbasnjuk O, Donowitz M, Li X (2006) Na+/H+ exchanger NHE3 activity and trafficking are lipid raft-dependent. J Biol Chem 281:17845–17855 21. Madre N, Smith KL, Graham CH, Jen A, Brady K, Hall S, Morris R (1999) Functionally different GPI proteins are organized in different domains on the neuronal surface. EMBO J 18:6917–6926

Chapter 3 Methods of Dopamine Research in Retina Cells Ana Lucia Marques Ventura, Fernando Garcia de Mello, and Ricardo Augusto de Melo Reis Abstract Dopamine is the main catecholamine found in the retina of most species, being synthesized from the L-amino acid tyrosine. Its effects are mediated by G protein coupled receptors subfamilies that are commonly coupled to adenylyl cyclase in opposite manners. There is evidence that this amine works as a developmental signal in the embryonic retina and several distinct roles have been attributed to dopamine in the retina such as proliferation, synaptogenesis, neuroprotection, increased signal transmission in cone, gap junction modulation, neuronal–pigmented epithelium–glial communication, and neuron–glia interaction. Here we describe methods that have been used in the study of the dopaminergic function in the retina in the last 40 years. We emphasize the approaches used in the studies on the development of the avian and rodent retina. The dopaminergic system is one of the first phenotypes to appear in the developing vertebrate retina. Key words: Retina, Dopamine, Cyclic AMP, Müller glia, Amacrine, Tyrosine hydroxylase, Development

1. Introduction Dopamine is a key catecholamine found in the vertebrate retina, present mainly in a subtype of amacrine cell where most of the machinery for synthesis (tyrosine hydroxylase—TH, dopamine decarboxylase—DDC) and release (vesicular monoamine—VMAT and membrane—DAT transporters) are found in most vertebrate species (1). The best way to study the functional response of dopamine in retinal cells is to characterize its receptors, since most of the effects mediated by dopamine are through two basic types of G protein-coupled receptor, D1 and D2, which stimulate and inhibit, respectively, the enzyme adenylyl cyclase (2). Dopamine-mediated cyclic AMP (cAMP) accumulation, via D1-like receptors, is observed very early during retina ontogeny, before synaptogenesis and, in Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_3, © Springer Science+Business Media, LLC 2013

25

26

A.L.M. Ventura et al.

some species, before the expression of TH, the enzyme that characterizes the neuronal dopaminergic phenotype (3, 4). D2-like receptors appear in the tissue days after D1-like activity is detected (5). In the embryonic avian retina, before the tissue is capable of synthesizing its own dopamine via TH, dopamine synthesis is observed from l-DOPA supplied to the neuroretina from the pigmented epithelium which results in dopaminergic communication in the embryonic tissue before TH expression (6). Recently, Müller glial cells have also been shown to be able to synthesize and release dopamine, at least in culture conditions (7, 8). Mixed neuron–glia cultures obtained from embryonic chick express D1A and D1B receptors mRNA, but not D1D, as detected by RT-PCR (9). Müller glia cell also expresses the D1 receptor (10). The binding of [3H]-SCH 23390 revealed a significant amount of expressed receptors and released dopamine was detected in cell extracts of cultured Müller cells exposed to the DA precursor, l-DOPA (11). Here we describe practical procedures related to dopamine research in retina cells, including signaling (cAMP accumulation), immunocytochemistry (for dopaminergic markers), mRNA quantification and receptor binding (for D1A, D1B, and D1D receptors, as well as D2) and assays in neuroprotection, synaptogenesis and dopamine release in early postnatal retina.

2. Materials 2.1. Cell Culture and Western Blotting

1. Dulbecco’s Modified Eagle’s Medium (DMEM), 10% fetal calf serum (FCS), and gentamicin (Invitrogen, Life Technologies, Rockford, IL). 2. Solution of trypsin (Worthington) stored in single use aliquots (0.1% at 0.5 mL) at −70°C. 3. Epidermal growth factor (1 mg/mL EGF) and B27 supplement (Invitrogen) prepared in single use aliquots of 25 μL 50 μg/mL and 0.5 mL stock, respectively. Both are added to 50 mL DMEM for the preparation of neurosphere retina culture. 4. CMF (Ca2+ and Mg2+ free solution): 76.55 g/L NaCl, 3.05 g/L KCl, 1.65 g/L Na2HPO4, 0.610 g/L KH2PO4, 21.95 g/L glucose, and 7.90 g/L NaHCO3. 5. Plastic dishes: 35 or 60 mm 4-well dishes (Falcon or Nunc. Int.). 6. Modified Laemmli buffer for cell lysis: 1 mL of 0.5 M Tris– HCl, pH 6.8 + 1.6 mL of 10% (w/v) sodium dodecyl sulfate (SDS) + 0.8 mL of glycerol + 0.4 mL of β-mercaptoethanol; prepare also a 10× solution of bromophenol blue (0.2%).

3

Dopamine in Retina

27

7. Running buffer (1 L): 3 g Trizma base, 14.4 g glycine, 10 mL 10% SDS. 8. Transfer buffer (1 L): 3 g Trizma base, 14.4 g glycine, 1 mL 10% SDS, 100 mL methanol. 9. TBS (1 L): 2.42 g Trizma base, 8 g NaCl; pH 7.6. 10. 9% polyacrylamide mini-gels: mix 2.25 mL of 1.5 M Tris–HCl buffer, pH 8.8, 2.7 mL of 30% acrylamide/0.8% bis solution, 4.05 mL H2O, 15 μL of TEMED, 15 μL of 10% ammonium persulfate solution, and 15 μL of 10% SDS. Pour in the gel apparatus, leaving space for a stacking gel and overlay with a 0.1% SDS solution. Polymerize for 30 min; when the polymerization line appears, pour off SDS solution and make stacking gel by mixing 625 μL of 0.5 M Tris–HCl, pH 6.8, 375 μL acrylamide/bis solution, 1.45 mL of H2O, 12.5 μL of TEMED, 25 μL of ammonium persulfate, and 25 μL of 10% SDS. Pour on the top of the running gel, insert comb, and let polymerize; remove comb and wash the wells with running buffer. 2.2. mRNA Preparation

1. Trizol and DNAse (Gibco, Life Technologies). Standard protocols in this section should be done according to kit instructions or according to ref. 12. 2. First strand synthesis of cDNA is performed using the First– Strand cDNA Synthesis Kit (Amersham Biosciences, GE Healthcare, Piscataway, NJ). 3. The following oligonucleotides are specific to amplify retinal cDNA preparations for avian dopamine D1A and D1B receptors, as well as for the L27 ribosomal protein (see Fig. 1 and ref. 9. D1A (GenBank sequence no. L36877): 5¢-CCAAGGGAGCAGAAGCTTTC-3¢ (base position 908)

Fig. 1. PCR products corresponding to D1A (372 bp), D1B (296 bp) and the ribosomal protein L27 (235 bp, internal control) mRNA separated on a 2% agarose gel and visualized after staining with 0.5 g/mL ethidium bromide.

28

A.L.M. Ventura et al.

5¢-TACCCGACAATGCTGGAGAC-3¢ (base position 1,279) PCR product = 372 bp D1B (GenBank sequence no. L36878): 5¢-GAGGACATGAGCACCACATG-3¢ (base position 610) 5¢-GTGTGATGGTGGCAGTCAAC-3¢ (base position 905) PCR product = 296 bp Ribosomal protein L27 as internal control (GeneBank sequence no. X56852): 5¢-AAGCCGGGGAAGGTGGTG-3¢ (base position 42) 5¢-GGGTGGGCATCAGGTGGT-3¢ (base position 276) PCR product = 235 bp 4. 50 μL reaction mixture: 50 pmol of specific oligonucleotides, 200 μM dNTPs, 2.5 U of Taq polymerase, enzyme buffer (20 mM Tris–HCl, 50 mM KCl, pH 8.4), and 1.5 mM MgCl2. 2.3. Binding Assays

1. Ice-cold lysis buffer: 5 mM Tris–HCl, 5 mM MgCl2, pH 7.4. 2. Incubation buffer: 50 mM Tris–HCl, pH 7.4, containing 120 mM NaCl, 5 mM KCl, 1.5 mM CaCl2, 4 mM MgCl2, and 1 mM EDTA. 3. [3H]-SCH 23390 (~70.3 Ci/mmol, Perkin-Elmer); 20 μM (+)-Butaclamol (Sigma). 4. GF/C glass fiber filters immersed in a 0.3% solution of polyethyleneimine (Sigma). 5. 10% Trichloroacetic acid (~50 mL).

2.4. Immunofluorescence

1. Clean spherical microscope coverslips (0.15 mm). 2. Phosphate buffered saline (PBS) in g/L: Mix 0.1 CaCl2 (0.680 mM), 0.2 KCl (2.7 mM), 0.2 KH2PO4 (1.47 mM), 0.12 MgSO4 (0.4896 mM), 8 NaCl (136 mM), and 1.15 Na2HPO4 (8 mM) (adjust to pH 7.4 with HCl if necessary), and pass through 0.22 μm filters. 3. Paraformaldehyde (Sigma): Prepare a 4% (w/v) solution in PBS fresh for each experiment. Dissolve in solution in a stirring hot-plate in a fume hood and then cool to room temperature for use. 4. Permeabilization solution: 0.25% (v/v) Triton X-100 or tween20 in PBS. 5. Antibody dilution buffer: 3% (w/v) BSA in PBS. 6. Secondary antibody: Anti-mouse or anti-rabbit IgG conjugated to Cy3 (Jackson Immunoresearch, West Grove, PA).

3

Dopamine in Retina

29

7. Nuclear stain: 300 nM DAPI (4,6-diamidino-2-phenylindole) in water. 8. Mounting medium: Antifade (Molecular Probes, Eugene, OR). 2.5. Dopamine Extraction

1. Tris–HCl solution, pH 8.8. 2. Dihydroxybenzylamine (DHBA, the internal standard of extraction). 3. Alumina 4. 100 mM perchloric acid. 5. Reverse phase column: LC-18 column (4.6 mm × 250 mm, Supelco). 6. Mobile phase: 20 mM sodium dibasic phosphate, 20 mM citric acid, pH 2.64, containing 10% methanol, 0.12 mM Na2EDTA, and 566 mg/L heptanesulfonic acid.

3. Methods 3.1. Retinal Cultures for Dopamine Assays

1. Remove the eyes, dissect the retinas free of the pigmented epithelium in Dulbecco’s modified Eagle’s medium (DMEM). 2. Transfer the retinal pieces and wash twice in Ca2+- and Mg2+free solution (CMF). 3. Dissociate the tissue using trypsin (Worthington) for 10 min (37°C). 4. At this point, the experimenter should decide in a number of choices as listed below.

3.1.1. Mixed Neuronal-Glial Cultures (See FIG. 2A)

Dissociate and plate 1/2 retina per dish (1–1.5 × 107 cells or more). It is not necessary to treat plastic dishes with substrates. Ideal for functional assays measuring receptor mediated second messenger shifts, binding or Western blot analysis for specific proteins mediated by dopamine.

3.1.2. Enriched Neuronal Cultures (See Fig. 2b)

Low density neuronal cultures, where approximately 2 × 106 cells or less are seeded onto treated poly L-Lysine (10 μg/mL) plastic dishes in DMEM medium plus 1% FCS (see Note 1).

3.1.3. Müller Glia Cell Cultures from Embryonic or Postnatal Retina (See Fig. 2c)

1. Use the amount of 5 × 106 cells over culture dishes in DMEM containing 10% FCS. 2. Medium should be changed every 3 days. 3. After approximately 10 days, cell cultures are treated with 4 mM ascorbic acid for 2 h to eliminate neurons (13).

30

A.L.M. Ventura et al.

Fig. 2. Dopamine has been investigated as a developmental signal in the retinal tissue or cultures prepared in many different ways. (a) Mixed neuron–glial cells (prepared in high density, with ~20 × 106 cells) is ideal for functional assays measuring receptor mediated second messenger shifts, binding or western blot analysis. (b) Enriched neuronal cells prepared in low density, with ~2 × 106 cells or less, seeded on treated poly L-Lysine (10 μg/mL) plastic dishes. (c) Müller glia culture from embryonic or postnatal retina prepared from progenitors (5 × 106 cells) in DMEM containing 10% FCS and cultured for 10 days, when neurons are eliminated (13). (d) Neurospheres retinal cultures prepared in the presence of EGF in an untreated culture dish. On day 5, neurospheres are plated under differentiating conditions that allows the emergence of all retinal neurons and Müller glia. (e) Photomicrograph (Dr. Marilia Guimarães) of a TH-positive cell in E10C3 chick retina cell culture (15).

4. Purified glial cultures can be used 3 days later and maintained for up to 3 weeks. These cultures can be used for different purposes such as signaling, binding, immunocytochemistry. 3.1.4. Neurospheres Retinal Cultures (See Fig. 2d)

3.2. Western Blotting for Detection of Dopamine Markers (TH, L-Dopa Decarboxylase, VMAT, DAT, nurr-1) 3.2.1. Preparation of Sample Extracts

Retinal cells are plated in DMEM supplemented with gentamicin, 1% B27 supplement, and 20 ng/mL epidermal growth factor (EGF) and then placed in an untreated 35 mm culture dish (Corning). On day 5, neurospheres are plated under differentiating conditions onto a poly-D-Lysine matrix (Invitrogen) coverslips with different substrates such as laminin or fibronectin (Sigma). 1. Incubate retinas or retinal cells in culture with drugs of interest in DMEM medium 2. Transfer tissues to ~70 μL of sample buffer without bromophenol blue. Mix well with vortex. 3. For retinal cells in culture, remove medium, wash cells with medium without serum, and add ~70 μL of sample buffer. Scrape cells with a large bore pipette tip and transfer viscous material to tubes. 4. Boil extracts in boiling water for 10 min and centrifuged at 27,000 × g for 10 min to remove non-soluble material.

3

Dopamine in Retina

31

5. Estimate protein content in 2 μL samples of extracts by the Bradford protein assay, using a BSA solution containing 2 μL of sample buffer as standard; make in duplicate or triplicate. 6. After protein content determination, add 0.1 vol. of a 10× bromophenol blue solution to the remaining volume of samples. 7. Prepare 9% polyacrylamide Subheading 2.1, item 10.

mini-gels

according

to

8. Mount gels in the running apparatus and add running buffer in the two chambers. 9. Add retinal extract samples (50 μg/lane) to the lanes of the SDS polyacrylamide gels and run at ~20 mA for 1–1.5 h, until dye reaches the end of the gel. 10. Soak PVDF membranes (GE Healthcare) for 10 s in pure methanol, 5 min in H2O, and 10 min in transfer buffer. Transfer proteins to PVDF membranes for 1 h, at 100 V. 11. Disassemble the transfer unit and stain membrane with a 0.2% Ponceau rouge solution in 3% TCA; check the transfer, mark molecular weight bands; remove Ponceau rouge solution with Tris-buffered saline (TBS). 3.2.2. Immunodetection

1. Block membranes with 5% nonfat milk in Tris-buffered saline (pH 7.6) with 0.1% Tween-20 (TBS-T), for 1 h, at room temperature. 2. Incubate membranes with diluted primary antibodies (1:2,000, TH; 1:1,500, DDC; 1:1,000, VMAT), overnight, at 4°C. 3. Wash membranes 3× with TBS-T. 4. Incubate with secondary antiserum conjugated to horseradish peroxidase (Bio-Rad Labs. Inc.) for 1 h at room temperature. 5. Wash membrane 2× with TBS-T for 5 min and 1× with TBS for 10 min. Results are detailed elsewhere (6, 7, 10, 14). 6. Develop blots using enhanced chemiluminescence (ECL), according to the manufacturer’s protocol (ECL plus, GE Healthcare). 7. Strip membranes in glycine 0.2 M, pH 2.2, for 30–40 min, at room temperature. 8. Re-probe membranes with anti-ERK 2 (Cell Signaling) or anti-actin (Santa Cruz Biotechnology, Santa Cruz, CA), at 4°C, followed by incubation with the secondary antibody and detection as described above (7, 10, 14).

3.3. Immunocytochemistry 3.3.1. Culture Fixation and Staining

Cultures should be prepared in coverslips for the purpose of saving antibody. 1. Briefly rinse retinal cell cultures with PBS and fix in 4% paraformaldehyde (PA) in 0.16 M Phosphate buffer (pH 7.2) for 5 min.

32

A.L.M. Ventura et al.

2. Rinse cultures extensively with sodium phosphate-buffered saline (PBS). 3. Rinse cultures with PBS plus Triton X-100 (0.25%). 4. Pre-incubate cultures with 5% bovine serum albumin (BSA) in PBS plus Triton X-100 (0.25%). 5. Incubate culture coverslips with primary antibody (1:500, TH; 1:700, DDC; 1:300, VMAT; 1:300 Nurr-1). 6. After several rinses in PBS, incubate cultures with appropriate secondary Alexa fluor 488 conjugated (Molecular Probes) or 598 conjugated (use 1:500 dilution). Alternatively, use secondary biotinylated antibody (Vector Labs, 1:200) for 2 h, followed by the avidin–biotin complex (ABC, Vector Labs) for an additional 2 h (see Subheading 3.3.2). 7. Wash in PBS and counterstain culture cells with DAPI. Mount slides in sodium n-propyl-gallate 0.2 M (pH 7.2) in glycerol. 8. Control cultures of immunohistochemistry reactions should be incubated with PBS in the absence of primary antibody. 9. Acquire photomicrographs with standard fluorescence microscopy. Example of TH detection in retina cells in culture or tissue is shown in (15, 16) (Fig. 3). 3.3.2. Retinal Tissue

1. Use alternate radial retina sections for performing immunohistochemistry. DDC, TH, VMAT or DAT are commonly markers used for dopaminergic immunohistochemistry (see Note 2). 2. Sections are rinsed in PBS and pre-incubated in 5% bovine serum albumin (BSA) and 3% normal goat serum in PBS, for 1 h. Then, sections are incubated in an antibody against DDC

Fig. 3. Photomicrograph of TH-positive amacrine cell located in the inner nuclear layer (INL) in a radial section of chick retina. Scale bar: 20 μm; ONL outer nuclear layer, GCL ganglion cell layer, IPL interplexiform layer. The image is from Dr. Patricia Gardino.

3

Dopamine in Retina

33

(Chemicon anti-rabbit 1:200 in PBS plus 0.25% Triton X-100, overnight). 3. Controls are performed by omission of the primary antibody. Next day, the retinal sections are rinsed in PBS and incubated in the secondary biotinylated antibody against rabbit IgG (Vector Labs, 1:200) for 2 h, followed by the avidin–biotin complex (ABC, Vector Labs) for an additional 2 h. The binding of the antibody to the sections is revealed by the addition of a substrate for peroxidase (DAB or SG, Vector Labs (Southfield, MI)). 4. Sections are coverslipped with 40% glycerol in PBS. Alternate sections are processed for TH immunohistochemistry however in this case, there was no need to heat the tissue to improve antibody penetration. The sections are pre-incubated in 5% BSA and 3% normal goat serum in PBS for 1 h followed by the incubation in rabbit antibody against TH (Eugene Tech, AB1569) at the dilution of 1:3,000 in PBS plus 0.25% Triton X-100, overnight. As for DDC the same procedure is applied to visualize the immunohistochemistry reaction with peroxidase. Sections are also coverslipped with 40% glycerol in PBS (4, 17, 18). 3.4. Detection of mRNA for Dopamine D1A and D1B Receptors from Chick Retina by RT-PCR (See Fig. 1) 3.4.1. Extraction of Retinal RNA

1. Remove eyes, transfer to cold Ca2+, Mg2+ free solution (CMF) and dissect retinal tissue free of pigmented epithelium under environmental light. 2. Transfer each retina to a new 1.5 mL tube containing 0.5 mL Trizol reagent to extract total RNA; mix well in a vortex (see Note 3). 3. Centrifuge for 10 min, at 18,000 × g, at 4°C. Transfer supernatant to a new tube and add 0.2 mL pure, high quality chloroform. Mix in vortex for 15 s and leave for 2–15 min at room temperature. 4. Centrifuge for 15 min, at 11,000 × g, at 4°C. Transfer the aqueous phase to a new tube and add 0.5 mL of high quality isopropanol; leave for 5–10 min at room temperature. 5. Centrifuge for 10 min, at 15,300 × g, at 4°C. Remove supernatant and add carefully 1 mL of high quality 75% ethanol in DEPC-treated water to wash pellet. 6. Remove supernatant and add 0.5 mL of 100% ethanol. 7. Store at −70°C until use.

3.4.2. DNaseI Treatment

1. Centrifuge total RNA samples for 5 min, at 15,300 × g, at 4°C. 2. Leave the tubes opened on the bench at room temperature until last traces of fluid have evaporated; add 25 μL of DEPCtreated water (see Note 4).

34

A.L.M. Ventura et al.

3. Incubate for 10 min at 55–60°C; keep at 4°C. 4. Dilute 1 μL of RNA preparations in 200 μL to determine the concentration and purity of RNA in a spectrophotometer by reading the OD at 260 and 280 nm (see Note 5). 5. Incubate 30 μg of RNA with RNAase free DNase I (0.5 U/μg RNA) in a 100 μL incubation mixture, containing enzyme buffer (20 mM Tris–HCl, 50 mM KCl, pH 8.4,) and 2 mM MgCl2; incubate for 10 min at 37°C. Keep at 4°C. 6. Extract RNA by adding 1 vol. of buffered phenol and mix. 7. Centrifuge at 54,300 × g for 10 min at 4°C. Remove supernatant and add 1 vol. of chloroform. 8. Centrifuge at 54,300 × g for 10 min at 4°C. Remove supernatant and add 1/10 vol. of 3 M NaOAc. 9. Add 2 vol. of 100% ethanol and incubate for 20 min at −70°C. 10. Centrifuge at 54,300 × g at 4°C. 11. Wash pellet with 70% ethanol. 12. Discard supernatant and add 20 μL DEPC-treated water. 13. Determine the concentration of RNA in samples using a spectrophotometer. 3.4.3. First Strand Synthesis of cDNA

1. Incubate RNA preparation for 10 min at 65°C to denature eventual double strand segments in RNA. 2. Dilute RNA to 1 μg/20 μL with DEPC-treated water. 3. Perform first strand synthesis of cDNA following the procedure described in the First strand cDNA synthesis kit (GE Healthcare). In brief, incubate 1 μg of RNA in a 33 μL reaction mixture containing Moloney murine leukemia virus reverse transcriptase, 0.2 μg of random hexamers, and 6 mM dithiothreitol for 60 min at 37°C. 4. Store at 4–8°C until use.

3.4.4. Amplification of cDNA by PCR

1. Amplify directly 5 μL of cDNA preparation in a 50 μL reaction mixture prepared according to Subheading 2.2 (see Note 6). 2. Run PCR by initially denaturing samples at 94°C for 4 min and submitting them to 25–27 cycles of 1 min at 94°C (denaturation), 1 min at 60°C (annealing) and 1 min at 72°C (extension), followed by a final extension of 5 min at 72°C. 3. Prepare a 2% agarose gel without Ethidium Bromide; add 2 μL of gel loading buffer (0.25% bromophenol blue, 0.25% xylene cyanol, 30% glycerol) to 10 μL samples. 4. Run samples and ladder in TAE buffer at 75 V for 40 min. 5. Stain gel with Ethidium bromide solution (0.5 μg/mL) ~40 min.

3

Dopamine in Retina

35

6. Destain in water for 20 min at room temperature; visualize under U.V. light and photograph with a digital camera using a yellow filter. 7. To compare the relative amount of the different PCR products, amplify the same amount of cDNA with specific oligonucleotides, photograph gels and determine the density of each gel band using the 1D Gel Analysis Software (Kodak) or a similar program. Relate the intensities of the bands corresponding to receptor mRNA with the intensity of the band corresponding to the internal control L27 ribosomal protein (see Note 7). Results are detailed (9, 10). 3.5. Dopamine D1 Receptor Binding Assays in Chick Retinal Membranes (See Fig. 4)

1. Remove eyes, transfer to cold Ca2+, Mg2+ free solution (CMF) and dissect retinal tissue free of pigmented epithelium under environmental light (see Note 8).

3.5.1. Membrane Preparation

3. Disrupt cells (using 10–13 stroke movements).

2. Transfer tissues to a Dounce homogenizer (type B) containing ~3 mL of ice-cold lysis buffer prepared in Subheading 2.3. 4. Transfer material to centrifuge tubes and wash the homogenizer with 1–2 mL of ice-cold incubation buffer (see Subheading 2.3). 5. Transfer to tubes and centrifuge at 27,000 × g, for 30 min, at 4°C

100

800 Bound/Free

Specific [3H]-SCH23390 binding (fmol/mg protein)

150

50

600 400 200 0

0 50 100 150 Bound (fmol/mg protein)

0 0.0

0.5

[3H]-SCH

1.0

1.5

2.0

23390 (nM)

Fig. 4. Specific binding of [3H]-SCH 23390 to homogenates of E8C4 (embryonic day 8 retina and 4 days in vitro) mixed neuron–glia cultures. Cell membranes are incubated with various concentrations of [3H]-SCH 23390 (0.1–2.0 nM) in Tris–HCl buffer, pH 7.4 in a final volume of 0.2 mL. Data from saturation isotherms are transformed by the method of Scatchard and submitted to linear regression analysis. [3H]-SCH 23390 bound with high affinity to homogenates of retinal cells in culture, showing a Kd value of 0.18 nM. The total number of [3H]-SCH 23390 binding sites revealed by the Scatchard plot (inset) was 129 fmol/mg protein.

36

A.L.M. Ventura et al.

6. Discard the supernatant and add ice-cold binding buffer to the resulting membrane pellet. 7. Transfer material to Dounce homogenizer and homogenize the clumps until a homogeneous mixture is obtained 8. Store at 4°C (see Note 9). 3.5.2. Incubation of Membranes with [3 H]-SCH 23390

1. Add 50 μL of previously prepared diluted solutions of the antagonist [3H]-SCH 23390 (~70.3 Ci/mmol, Perkin-Elmer) to all incubation tubes (see Note 10). 2. Add 50 μL of buffer to the incubation tubes marked as “Total binding” and 50 μL of 20 μM (+)-Butaclamol to incubation tubes named “nonspecific binding” (see Note 11). 3. Begin incubation of 60 min, at room temperature, with the addition of 100 μL samples of membrane preparation (~0.1 mg protein). 4. While incubation of membranes is running, immerse GF/C glass fiber filters in a 0.3% solution of polyethyleneimine; this procedure decreases nonspecific adsorption of the ligand to the filter (see Note 12). 5. While incubation is running, precipitate a sample of membrane preparation with an equal volume of 10% Trichloroacetic acid to estimate protein content later; store at −20°C until use. 6. Terminate the incubation of tubes in a successive order; dilute the content of each tube with 3 mL of ice-cold washing buffer (50 mM Tris–HCl, pH 7.2). 7. Filter content of each tube under negative pressure through GF/C pretreated filters. 8. Wash tubes three times with 2 mL of washing buffer and filter. 9. Add 50 μL of [3H]-SCH 23390 diluted solutions directly to separate dry filters to estimate precisely the concentrations of ligand that were added to the incubation tubes. Estimate radioactivity bound to filters by scintillation spectroscopy (see Note 13). 10. Centrifuge precipitated membrane preparation obtained in step 5, add 0.1–0.2 N NaOH, and determine protein content. 11. Calculate the specific binding of the radioactive ligand by subtracting the nonspecific binding from the total binding. Transform values of radioactivity in cpm to fmol of ligand by using the specific activity of the radioactive ligand; divide values by protein content. This procedure will give an amount of receptors in fmol/mg protein.

3

Dopamine in Retina

37

12. Estimate radioligand parameters KD and Bmax using Graphpad Prism software (see Note 14). Representative results are detailed (11, 19). 3.6. cAMP Accumulation (See Fig. 5)

3.6.1. Stimulation of Cells

cAMP is the main signal generated by dopaminergic input in the retina and it can be estimated in this tissue or in retinal cultures according to a competitive binding assay described previously (3) (see Note 15). 1. Retina cells, mixed neuron–glial cells, or confluent Müller glial cells in culture are pre-incubated for 10 min at 37°C in DMEM medium buffered with 20 mM HEPES at pH 7.3, containing 0.5 mM isobutylmethylxantine and 100 μM ascorbic acid to inhibit cAMP-dependent phosphodiesterase and prevent oxidation of dopamine, respectively. 2. Add dopamine, D1-like agonists, or antagonist at the indicated final concentration and incubate further for 15 min; stop reaction by adding trichloroacetic acid to a final concentration of 5%. 3. Scrape cells and transfer all material to tubes; keep frozen until further use.

3.6.2. Ion Exchange Chromatography of Samples

1. Add trace amounts of [3H]-cAMP (50 nCi in 50 μL) to sample tubes and centrifuge for 30,000 × g for 30 min. Separate supernatants and dissolve precipitates with NaOH to measure protein content later. Add supernatants to ion-exchange resin columns (Dowex AG50W-X4, 200–400 mesh) previously equilibrated with 1 N HCl (see Note 16).

Fig. 5. cAMP accumulation in cells of cultures at E8C4 stimulated with dopamine (100 μM) is completely blocked when pre-incubated with the selective D1-like receptor antagonist, SCH-23390 (1 μM).

38

A.L.M. Ventura et al.

2. Wash columns with 6 mL of water each and discard; elute columns three times with 3 mL of water and collect. Determine radioactivity in 150 μL of the fractions by scintillation spectroscopy; also determine radioactivity in 50 μL of [3H]-cAMP tracer solution. Calculate the % recovery of [3H]-cAMP in tubes; Select the tubes with highest recovery to measure cAMP. 3.6.3. cAMP Determination in Samples

1. Incubate 50 or 100 μL samples with 25 μL solution of PKA / BSA and a fixed concentration of [3H]-cAMP (2 pmol/20 μL) in 50 mM acetate buffer, pH 4.0, at 4°C, for 90 min; add H2O to complete the volume of the reaction to 200 μL (see Note 17). 2. At the same time, construct a standard curve with known concentrations of nonradioactive cAMP (use 0, 1, 3, 5, 10, and 15 pmol). Incubate with PKA and [3H]-cAMP in acetate buffer as above. 3. Interrupt reaction in samples by adding 2 mL of 200 mM phosphate buffer, pH 6.0; filter samples through Millipore acetate filters (0.45 μm), washing tubes 3× with 2 mL phosphate buffer; dry filters and quantify the bound radioactivity by liquid scintillation. 4. Plot data from the standard curve by relating radioactivity versus concentrations of nonradioactive cAMP. Estimate nonradioactive cAMP in samples; correct values with the % recovery from the chromatography columns and divide by protein content of samples to obtain values in pmol/mg protein; detailed information are shown in refs.( 3, 10, 11, 14, 20.)

3.7. Dopamine Extraction and Quantification

Dopamine in cell extracts can be measured by HPLC analysis coupled with electrochemical detection (0.5 V) as described before (6, 7, 21). 1. Cell extracts are added in Tris–HCl solution (pH 8.8) in the presence of dihydroxybenzylamine (DHBA, the internal standard of extraction). 2. The amines are precipitated by addition of alumina, then wash three times with water and eluted with 100 mM perchloric acid. 3. After centrifugation, filter and inject the supernatant into the reverse phase column. Fast isocratic separation is obtained using an LC-18 column eluted with the mobile phase described in Subheading 2.5. 4. Express the results as nmol/mg cell protein.

3

Dopamine in Retina

39

In summary, dopamine is a key neurotransmitter in the retina as its effects guide the development of this tissue and mediates several distinct roles such as proliferation, signaling, differentiation and death. Several elements of the dopaminergic system (receptors, enzymes and transporters) emerge in a temporally defined way in this organ. It is also clear that dopamine regulates retinal morphogenesis by inhibiting the extension of neurites of retinal cells (22), reduces retinal apoptosis (23) and restricts retinal cell divisions (24). Dopamine is also known to inhibit the function of the N-Methyl D-Aspartate receptor in the avian or rodent retina as assayed through neurotransmitter release (25) or whole cell currents (26). This chapter describes key approaches to study dopamine in the retina, which might be useful to evaluate dopaminergic roles in the normal retina as well as in deficits such as myopia and albinism and disorders in the brain such as Parkinson’s disease.

4. Notes 1. It is important to keep serum at a low percentage. At low density, neurons are the vast majority of cells in the first 72 h. Glial appearance could be further avoided with cytosine arabinoside (10 μM) treatment. These cultures are ideal for neurotoxicity assays as dopamine have been shown as a potential neuroprotective factor in the retina (23). 2. In order to improve DDC labeling, radial sections are dipped in 10 mM citrate buffer, pH 6.0, and heated in microwave oven for 1 min. 3. Use new tubes and tips for RNA extraction and treatment; solutions always should be made in 0.1% DEPC-treated water. 4. Working in a laminar flow reduces contamination with RNases from bacteria and other contaminants. 5. Reading at 260 nm allows calculation of the concentration of nucleic acid in the sample. An OD of 1 corresponds to approximately 40 μg/mL for single strand DNA or RNA. The ration between readings at 260 and 280 nm (OD260/OD280) provides the purity of nucleic acids. Pure preparations of RNA should have values of 2.0. Contamination with protein or phenol will give values significantly less than 2.0; If samples are not pure, extract RNA with buffered phenol as described in step 6 of the DNase I treatment procedure.

40

A.L.M. Ventura et al.

6. For PCR amplification of cDNA, it is useful to prepare first a master mix containing water, 1× enzyme buffer, dNTPs and MgCl2. 7. To compare the expression of mRNA for different receptor subtypes in the tissue it is necessary to work in the linear portion of the PCR amplification curve, where the amount of amplified PCR products are directly proportional to the initial amount of cDNA used in the PCR amplification. To determine this, make a PCR amplification curve for each pair of specific oligonucleotides by varying the number of PCR amplification cycles. 8. Membrane preparations of cultured retinal cells can be obtained by adding lysis buffer to cultures and transferring material to centrifugation tubes with a pipette. 9. Other homogenizers such as Polytron or Potter can also be used; prepare incubation tubes while homogenate is under centrifugation. 10. A stock solution of 400 nM of [3H]-SCH 23390 should be prepared with binding buffer. For each experiment, successive dilutions from this stock should be prepared; concentrations of 0.4, 0.8, 1.6, 2.4, and 6.4 nM will result in 0.1, 0.2, 0.4, 0.8, and 1.6 nM final concentrations. As SCH 23390 is a high affinity antagonist, final concentrations should be in the range of 0.1–4 nM. 11. Total binding tubes will contain only membranes and the radioactive ligand and will provide the specific binding of the ligand to receptors + nonspecific binding of the ligand. Nonspecific binding tubes will contain membranes + radioligand + excess of a nonradioactive ligand to block the specific binding of the radioactive ligand to receptors, thus providing an estimate of the nonspecific binding of the ligand. Values obtained in these tubes should be subtracted from the values obtained in the total binding tubes in order to estimate the specific binding of the radioactive ligand to receptors; three total binding + three nonspecific binding tubes should be made for each ligand concentration. 12. GF/B glass fiber filters can also be used; however, they have a higher nonspecific adsorption of the ligand. 13. The reaction mixtures should be filtered under a medium negative pressure. If the pressure is too high, membranes will pass through filters; if pressure is too low, the ligand will dissociate from receptors by dilution. The ideal pressure is the one at which the buffer flows constantly through filters, thus washing them.

3

Dopamine in Retina

41

14. The amount of D1 receptors can also be estimated by performing displacement experiments; in this case, membranes are incubated with a constant concentration of [3H]-SCH 23390 (around the KD value) and increasing concentrations of nonradioactive compounds. 15. Although cAMP is the major signal stimulated by dopamine, other second messengers have been reported to be stimulated by D1 receptors such as inositol triphosphate, in the striatum (27). Retinal cells should be incubated with 1 μCi myo-[2-3H] inositol (Perkin-Elmer) for 3 h. Cultures or retina pieces are treated with selective agonists for 15 min in the presence of 10 mM LiCl (induce accumulation of inositol phosphates). Stop the reaction by the addition of trichloroacetic acid followed by ether extraction; the supernatant containing inositol polyphosphates are separated by ion exchange chromatography (Dowex AG1-X8 resin, formate form; Bio-Rad). Inositol triphosphate (ip3) is quantified in a liquid scintillation analyzer (28). 16. Use ~8 cm (height) by 1 cm (diameter) Dowex-AG50W-X4 columns. Equilibrate columns with 10 mL 1 N HCl + 10 mL H2O immediately before use. 17. A stock solution of the regulatory sub-unit of PKA should be prepared before cAMP assays. Prepare a 1 mg/mL PKA solution; keep aliquots frozen until use; prepare a 20 mg/mL solution of BSA; keep frozen. Dilute the necessary amount of PKA for the entire assay (five to tenfold) in a working 4 mg/ mL solution of BSA.

Acknowledgements This work was supported by grants from FAPERJ, CNPq, PROPPiUFF and INCT-CNPq (INNT). References 1. Witkovsky P, Dearry A (1992) Functional roles of dopamine in the vertebrate retina. Prog Retinal Res 11:247–292 2. Reis RAM, Ventura ALV, Kubrusly RC, de Mello MC, de Mello FG (2007) Dopaminergic signaling in the developing retina. Brain Res Rev 54:181–188 3. de Mello FG (1978) The ontogeny of dopamine-dependent increase of adenosine 3¢,5¢-cyclic monophosphate in the chick retina. J Neurochem 31:1049–1053 4. Gardino PF, dos Santos RM, Hokoc JN (1993) Histogenesis and topographical distribution of

tyrosine hydroxylase immunoreactive amacrine cells in the developing chick retina. Brain Res Dev Brain Res 72:226–236 5. Ventura ALM, Klein WL, de Mello FG (1984) Differential ontogenesis of D1 and D2 dopaminergic receptors in the chick embryo retina. Brain Res 314:217–223 6. Kubrusly RCC, Guimarães MPZ, Vieira APB, Hokoç JN, Casarini DE, de Mello MC, de Mello FG (2003) L-DOPA supply to the neuro retina activates dopaminergic communication at the early stages of embryonic development. J Neurochem 86:45–54

42

A.L.M. Ventura et al.

7. Kubrusly RC, Panizzutti R, Gardino PF, Stutz B, Reis RA, Ventura AL, de Mello MC, de Mello FG (2008) Expression of functional dopaminergic phenotype in purified cultured Müller cells from vertebrate retina. Neurochem Int 53:63–70 8. de Melo Reis RA, Ventura ALV, Schitine CS, de Mello MC, de Mello FG (2008) Müller glia as an active compartment modulating nervous activity in the vertebrate retina: neurotransmitters and trophic factors. Neurochem Res 33:1466–1474 9. Soares HC, Reis RA, De Mello FG, Ventura AL, Kurtenbach E (2000) Differential expression of D(1A) and D(1B) dopamine receptor mRNAs in the developing avian retina. J Neurochem 75:1071–1075 10. Kubrusly RC, da Cunha MC, Reis RA, Soares H, Ventura AL, Kurtenbach E, de Mello MC, de Mello FG (2005) Expression of functional receptors and transmitter enzymes in cultured Müller cells. Brain Res 1038:141–149 11. Ventura ALM, de Mello FG (1990) D1 dopamine receptors in neurite regions of embryonic and differentiated retina are highly coupled to adenylyl cyclase in the embryonic but not in the mature tissue. Brain Res 530:301–308 12. Sambrook J, Fritsch E, Maniatis T (1989) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory, New York 13. Reis RA, Cabral da Silva MC, Loureiro dos Santos NE, Bampton E, Taylor JS, de Mello FG, Linden R (2002) Sympathetic neuronal survival induced by retinal trophic factors. J Neurobiol 50:13–23 14. Kubrusly RC, Ventura AL, Reis RA et al (2007) Norepinephrine acts as D1-dopaminergic agonist in the embryonic avian retina: late expression of β1-adrenergic receptor shifts norepinephrine specificity in the adult tissue. Neurochem Int 50:211–218 15. Guimarães MZP, Hokoç JN, Duvoisin R et al (2001) Dopaminergic retinal cell differentiation in culture: modulation by forskolin and dopamine. Eur J Neurosci 13:1931–1937 16. Borba JC, Henze IP, Silveira MS et al (2005) Pituitary adenylate cyclase-activating polypeptide (PACAP) can act as determinant of the tyrosine hydroxylase phenotype of dopaminergic cells during retina development. Brain Res Dev Brain Res 156:193–201 17. Dos Santos RM, Gardino PF (1998) Differential distribution of a second type of tyrosine hydroxylase immunoreactive amacrine cell in the chick retina. J Neurocytol 27:33–43

18. Taveira da Silva R, Hokoç JN, de Mello FG et al (2009) Differential immunodetection of L-DOPA decarboxylase and tyrosine hydroxylase in the vertebrate retina. Int J Dev Neurosci 27:469–476 19. De Mello MC, Pinheiro MC, de Mello FG (1996) Transient expression of an atypical D1-like dopamine receptor system during avian retina differentiation. Braz J Med Biol Res 29:1035–1044 20. de Mello MC, Ventura ALM, Paes de Carvalho R, Klein WL, de Mello FG (1982) Regulation of dopamine and adenosine-dependent adenylate cyclase systems of chick embryo retina cells in culture. Proc Natl Acad Sci USA 79:5708–5712 21. Arita DY, Di Marco GS, Schor N, Casarini DE (2002) Purification and characterization of the active form of tyrosine hydroxylase from mesangial cells in culture. J Cell Biochem 87:58–64 22. Lankford KL, De Mello FG, Klein WL (1988) D1-type dopamine receptors inhibit growth cone motility in cultured retina neurons: evidence that neurotransmitters act as morphogenic growth regulators in the developing central nervous system. Proc Natl Acad Sci USA 85:2839–2843 23. Varella MH, de Mello FG, Linden R (1999) Evidence for an antiapoptotic role of dopamine in developing retinal tissue. J Neurochem 73:485–492 24. Tibber MS, Whitmore AV, Jeffery G (2006) Cell division and cleavage orientation in the developing retina are regulated by L-DOPA. J Comp Neurol 496:369–381 25. Do Nascimento JLM, Kubrusly RCC, Reis RAM, De Mello MC, De Mello FG (1998) Atypical effect of dopamine in modulating the functional inhibition of NMDA receptors of cultured retina cells. Eur J Pharmacol 343:103–110 26. Castro NG, de Mello MC, de Mello FG, Aracava Y (1999) Direct inhibition of the N-methyl-D-aspartate receptor channel by dopamine and (+)-SKF38393. Br J Pharmacol 126:1847–1855 27. Wang HY, Undie AS, Friedman E (1995) Evidence for the coupling of Gq protein to D1-like dopamine sites in rat striatum: possible role in dopamine-mediated inositol phosphate formation. Mol Pharmacol 48:988–994 28. Reis RA, Kubrusly RC, de Mello MC, de Mello FG (1995) Transient coupling of NMDA receptor with ip3 production in cultured cells of the avian retina. Neurochem Int 26:375–380

Chapter 4 Capture of D2 Dopamine Receptor Signaling Complexes in Striatal Cells for Mass Spectrometry Proteomic Analysis Nadine Kabbani and Jacob C. Nordman Abstract In recent years advancements in proteomic techniques have contributed to the understanding of protein interaction networks (Interactomes) in various cell types. Today, high throughput proteomics promises to define virtually all of the components of a signaling and a regulatory network within cells for various molecules including membrane-spanning receptors. The D2 dopamine receptor (D2R) is a primary mediator of dopamine transmission in the brain. Signaling through D2Rs has been linked to dopamine-mediated effects on motivation, reward, locomotion and addiction to drugs of abuse. In the striatum, the D2R is a key mediatory of dopamine transmission. Actions on this receptor are an important pharmacological property of various drugs including typical antipsychotics and drugs of abuse. Here we provide an approach for the identification protein interaction networks of the D2R within striatal cells. We discuss key assays and techniques, such as cellular membrane protein fractionation, western blot analysis, magnetic bead coimmunoprecipitation, and liquid chromatography electrospray ionization (LC-ESI) mass spectrometry, that can be used for the isolation and characterization of D2R protein interaction networks. This approach presents a reliable method for the identification and characterization of D2R signaling within cells. Key words: Proteome, Dopamine signaling, Mass spectrometry, Antibody, Membrane spanning protein, G protein coupled receptors, Interactome

1. Introduction Dopamine receptors are members of a large family of membrane spanning proteins that have seven transmembrane domains and can functionally couple to heterotrimeric GTP binding proteins (G proteins) (1, 2). The endogenous ligand for these receptors, dopamine, is one of the main catecholamine transmitters in mammals (2, 3). Dopamine has been found to play a key role in regulating a range of neural functions including locomotor behavior, motivational state, and reward processes (4). In addition, it has been demonstrated that an increase in dopamine transmission Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_4, © Springer Science+Business Media, LLC 2013

43

44

N. Kabbani and J.C. Nordman

caused by drugs of abuse can lead to addiction in many species (5). Given their central role in dopamine transmission, the dopamine receptor (DR) family represents a key class of molecular targets for pharmacological therapy (6). Within mammals, DRs are divided into five types (D1-D5). Based on their pharmacologic profile and genetic sequence similarity, DRs are subdivided into D1-like and D2-like categories (1, 2). D1-like (D1 and D5) receptors interact with Gs (stimulant) proteins to increase the production of cellular cAMP by stimulating adenyl cyclase at the membrane, while D2-like (D2, D3, D4) receptors interact with Gi (inhibitory) proteins and decrease the production of cellular cAMP by inhibiting adenyl cyclase activity also at the membrane (7–9). In addition to their noted effects on cellular cAMP production, DRs are known to exert effects on cellular excitability by regulating various ligand and voltage-gated ion channels, ion transporters, and the sodium-potassium pump (10). The activation of D2Rs in various cell types has been found to impact signaling of cellular molecules such as cAMP regulated Protein Kinase A (PKA), Protein Kinase C (PKC), and calmodulin regulated protein such as calcineurin (PP2B) and Ca2+/Calmodulin Dependent Protein Kinase II (CaMKII) (8). The recent discovery of diverse dopamine receptor interacting proteins (DRIPs) in cells (9, 11) suggests that the signaling and regulatory properties of DRs are mediated via their direct protein interactions in cells. Indeed, certain DRIPs appear to regulate the major signaling and regulatory features of the D2R including its transport and localization within neurons, stability at the plasma membrane, and downstream signaling (7, 8). A growing demand in understanding molecular functions has spurred efforts in the design of tools for enhanced detection of intracellular signaling. In addition to advancements in cellular imaging, which have provided a mean to examine the spatial aspects of protein expression and trafficking, mass spectrometry techniques such as matrix assisted laser desorption/ionization-time of flight (MALDI-TOF) or liquid chromatography electrospray ionization (LC-ESI) now allow for the identification of signaling and interaction networks within various compartments of living cells (12). This enables a rapid (high throughput) strategy for the capture of information on the dynamics and composition of a signaling protein network within the cell (13, 14). Mass spectrometry analysis of affinity captured protein complexes has been effectively used by our group and others to define high and low abundance interactions as well as small and large size protein binding partners of neurotransmitter receptors in the brain (15–17). The aim of this chapter is to provide a step-by-step guide for the isolation and characterization of D2R complexes (D2Rs + DRIPs) from cells.

4

Proteomics of D2 Receptors

45

Since the striatum is an abundant site of D2R expression, we describe procedures for detecting D2Rs endogenous to striatal cells. Information gained from understanding receptor–protein interactions may aid in the treatment of brain disease.

2. Materials 2.1. Protein Preparation from Cultured Cells

1. Cell Culture Medium: Neurobasal (NB) with 2% B-27 supplement (Invitrogen, Life Technologies, Grand Island, NY), 5% Horse Serum (Gibco, Life Technologies, Grand Island, NY), 1% Pneumococcal Streptomycin (Gibco). 2. Poly-L-Lysine (Invitrogen). 3. Laminin (Invitrogen). 4. Non-denaturing protein extraction solution: 20 mM Tris–HCl, pH 7.4, 1% Triton X-100, 2 mM EDTA, 137 mM NaCl, 10% glycerol, and 1× protease inhibitor cocktail (Roche, Indianapolis, IN). 5. Sterile, optically clear polystyrene petri dishes. 6. Sterile cell scrapers.

2.2. Protein Preparation from Brain Tissue

1. Cold dissection buffer solution: 4 mM Hepes, 1 mM EDTA, 0.32 M sucrose, pH 7.4. 2. Cold cutting buffer: 10 mM Tris, 320 mM sucrose, 1 mM Phenylmethanesulfonylfluoride (PMSF) and 1× protease inhibitor cocktail, pH 7.4 (Roche). 3. Bradford dye reagent kit (Bio-Rad, Hercules, CA). 4. Non-denaturing protein extraction solution (20 mM Tris–HCl (pH 7.4)), 1% Triton X-100. (VWR), 2 mM EDTA, 137 mM NaCl, 10% glycerol, 1× protease inhibitor cocktail (Roche). 5. Glass dounce homogenizer.

2.3. Protein Quantification

1. Bovine Serum Albumin (BSA). 2. Quickstart Bradford Dye Agent (1×) (Bio-Rad). 3. Spectrophotometer.

2.4. Immunoprecipitation of Dopamine Receptors

1. Protein G Dynabeads® (Invitrogen). 2. Magnetic Tube Rack (Invitrogen). 3. Phosphate Buffer Saline (PBS). 4. Tween-20. 5. Phosphate Buffer Saline with 0.1% Tween 20 (PBST).

46

N. Kabbani and J.C. Nordman

6. Monoclonal Anti-D2R (EMD, Millipore, Billerca, MA). 7. NuPAGE LDS Sample Buffer (4×) (Invitrogen). 8. NuPAGE Sample Reducing Agent (10×) (Invitrogen). 9. Mass Spectrometry elution buffer: 49% acetonitrile, 2% trifluoroacetic acid in deionized water. 2.5. Mass Spectrometry

1. 10 mM DTT (Sigma-Aldrich). 2. 20 ng Β-casein (Sigma-Aldrich). 3. 8 M Urea (Sigma-Aldrich). 4. Sequencing grade trypsin (Invitrogen). 5. Glacial acetic acid (Sigma-Aldrich). 6. Angiotensin I & II (Sigma-Aldrich). 7. C-18 Zip-Tip Desalting Columns (Millipore, Billerca, MA). 8. Zip-Tip washing buffer: 0.1% trifluoroacetic acid in deionized water. 9. Zip-Tip elution buffer: 50% acetonitrile, 0.1% trifluoroacetic acid in deionized water. 10. 0.1% Formic acid (Thermo Fisher Scientific, Rockford, IL). 11. SpeedVac.

2.6. Protein Cross-linking

1. 2 mM DSP (dithiobis[succinimidylpropionate]) (Pierce, Rockford, IL). 2. 2.5 mM BS3 (Pierce). 3. Glass dounce homogenizer (5–7 mL volume capacity). 4. DMSO (solvent for use with DSP) (Sigma-Aldrich). 5. 20 mM Tris–HCl, pH 7.5.

2.7. ReverseCross-linking and In-Gel Digestion

1. Coomassie stain: 50% methanol, 10% acetic acid, 0.25% Coomassie Blue R-250 (Sigma-Aldrich) in deionized water. 2. Destaining Solution: 16.5% ethanol, 5% acetic acid in deionized water. 3. 50 mM ammonium bicarbonate (NH4HCO3). 4. In-Gel Reducing Solution: 25 mM DTT, 50 mM NH4HCO3 in deionized water. 5. In-Gel Alkylating Solution: 20 mM iodoacetamide (dissolve in 500 mM NH4HCO3), 50 mM NH4HCO3 in deionized water. 6. Drying Solution: 80% acetonitrile, 50 mM NH4HCO3 in deionized water. 7. In-Gel Digestion Solution: 10 ng/μL in ice-cold 50 mM NH4HCO3.

4

Proteomics of D2 Receptors

47

8. In-Gel Extraction Buffer: 50% acetonitrile, 2% acetic acid in deionized water. 2.8. Centrifugation

1. Low to medium speed centrifugation was achieved using the Eppendorf Centrifuge 5810R series (Eppendorf AG, Hamburg, Germany). 2. High speed ultracentrifugation was achieved using the Sorvall WX Ultra Series Centrifuge (Thermo Scientific). 3. A variety of rotors can be used for the described centrifugation procedures. In our experience, both swinging bucket and fixed angle rotors work well for the isolation of membrane protein fractions. In general, swinging bucket rotors are considered better suited for particle separation and advantageous when working with smaller volume.

3. Methods The D2R complex is made up of numerous direct and indirect interacting proteins. Many of these interactions are dynamic, being sensitive to disruptions in the state of the D2R during various forms of experimentation. In particular, molecular and biochemical methods for the dissociation of tissue, lysis of cells, and the efficient solubilization of membrane bound proteins are all known barriers in the study of receptor–protein interactions. For this reason we have emphasized experimental conditions that maximize on the preservation of endogenous protein interactions in the proteomic analysis of D2R protein complexes within striatal cells. This procedure can be used for D2R analysis from cultured striatal neurons as well as native striatal tissue. In addition, we present a cross-linking strategy that can preserve protein–protein interactions in order to define the D2R complex within cells. The advantages of this procedure include preservation of the D2R complex in the live cell, a direct visualization of several multi-protein complexes on an SDS-PAGE gel using a western blot or a Coomassie stain, and an identification of the proteins which make up those complexes using mass spectrometry peptide detection. 3.1. Protein Preparation from Primary Striatal Neurons

1. Primary cultures of striatal neurons can be derived from late embryonic day 18 embryos or, less preferably, neonate day 0 or 1 rat or mouse. Techniques for tissue dissection, cellular disassociation and plating of striatal cells have been well described in the literature. 2. For proteomic analysis, striatal cells are best plated at medium density concentration of 100 cells/mm2 in NB/B27 medium

48

N. Kabbani and J.C. Nordman

(see Note 1). Cells adhere well onto dishes precoated with poly-L-lysine (12.5 μg/mL) and laminin (5 μg/mL). 3. Perform medium changes every 2–3 days or as needed and maintain cells in culture for up to 2 weeks for proteomic analysis (see Note 2). 4. For protein identification, gently remove the culturing medium and replace with 3 mL of room temperature, sterile, PBS. This volume is per 100 mm size petri dish; however, when using different size dishes it possible to scale down/up the volume of PBS according to size. 5. Gently remove cells using a cell scraper (see Note 3). For less adherent cells, it is possible to remove cells via a gentle suction. 6. Pool samples into a 15 mL or a 50 mL conical tube as appropriate (Falcon). 7. Spin down the cells at 129 × g for 5 min at 4°C. 8. Carefully remove the supernatant and discard. 9. Wash the cells one time using an ice-cold PBS solution. The volume of PBS is not very critical; however, we suggest a volume consistent with the pooled final volume in step 6. 10. Gently wash the cells in PBS by inverting the capped tube once or twice. 11. Spin down the cells at 129 × g for 5 min at 4°C. 12. Carefully remove the supernatant and discard. 13. Resuspend cells into 5 mL of ice-cold PBS and transfer into a dounce homogenizer. 14. Manually homogenize the cells using gentle, slow, full strokes. 15. Do not cause bubbles during homogenization. 16. Homogenize until the solution turns consistent and fairly smooth. On average five to six strokes are sufficient for the homogenization of cultured cells. 17. Transfer the homogenate into prechilled centrifugation tubes. 18. Spin down the supernatant at 39,000 × g for 1 h for the isolation of membrane proteins. 19. A visible off white colored pellet should appear at the end of the ultracentrifugation step. This is the membrane fraction. 20. Discard the supernatant by slowly decanting or pipetting. The pellet should be tightly bound to the centrifugation tube. 21. Resuspend the pellet in 1 mL of the non-denaturing protein extraction buffer to solubilize membrane proteins (see Note 4).

4

Proteomics of D2 Receptors

49

22. Triturate the pellet gently, several times, by pipetting up and down in the extraction buffer. 23. Do not try to break up or dissolve the entire pellet in the buffer. 24. Incubate the pellet in the protein extraction buffer for 1 h (or overnight) at 4ºC with gentle mixing. 25. Centrifuge at 12,851 × g for 20 min at 4ºC. 26. Remove and keep the supernatant. This is the triton soluble membrane protein fraction containing D2Rs and their interacting proteins. 27. Optional: retain the pellet and store at −20ºC for up to 3 months. This pellet corresponds to the triton insoluble fraction, which maybe enriched in subcellular domains such as lipid rafts and postsynaptic densities. 28. Determine the final protein yield and concentration using the Bradford kit (Bio-Rad) in accordance with the manufacturer’s protocols and the established method of Bradford (18). 3.2. Protein Preparation from Striatal Tissue

1. Fresh striatal tissue can be obtained from either rat or mouse brains of various ages depending on the experimental paradigm. Elaborate protocols on how to dissect the striatum of rodents can be found elsewhere. 2. For high fidelity protein analysis, rapidly place the striatal tissue into a freshly made cold dissection buffer solution kept on ice. 3. We recommend a 1:1 ratio of rat striatum to dissection buffer (i.e., pool 4 whole striata of adult rats into a 4 mL solution of cold dissection buffer). 4. Transfer the tissue (in the dissection buffer) into a prechilled dounce homogenizer. 5. Keep the homogenizer on ice throughout the procedure. 6. Manually homogenize the tissue using gentle, slow, full strokes. 7. Do not over stroke or cause bubbles during homogenization. 8. Homogenize until the solution turns consistent and most of the large tissue pieces have broken. 9. On average eight to ten strokes are sufficient for the homogenization of brain tissue. 10. Transfer the homogenate into a prechilled centrifugation tube. 11. Spin down the homogenate at 700 × g for 10 min at 4°C. 12. Collect the supernatant fraction (S1) and store on ice during steps 13–18.

50

N. Kabbani and J.C. Nordman

13. Resuspend the pellet in the same volume of cold dissection as before (step 4). 14. Rehomogenize the pellet as before (steps 5–9). 15. Transfer the homogenate into a prechilled centrifugation tube. 16. Spin down the homogenate at 700 × g for 10 min at 4°C. 17. Collect the supernatant fraction (S2) and keep on ice. 18. Combine the supernatant fractions (S1 + S2) in prechilled ultracentrifugation tubes. This is the total striatal homogenate. 19. Ultracentrifuge the S1 + S2 sample at 40,000 × g for 1 h at 4°C. 20. A visible pellet should appear at the end of the ultracentrifugation step. This is the cellular membrane fraction from striatal tissue. 21. Discard the supernatant by slowly decanting or pipetting. The pellet should be tightly bound to the centrifugation tube. 22. Resuspend the pellet in non-denaturing protein extraction buffer to solubilize membrane proteins. We recommend a volume of 2 mL extraction buffer for four whole striata of adult rats for high protein yield content. 23. Triturate the pellet gently, several times, by pipetting up and down in the extraction buffer and proceed through the solubilization procedure of the membrane fraction as previously indicated in Subheading 3.1. 3.3. Coimmunoprecipitation of Dopamine Receptor Complexes

1. Place a 1.5 mL microcentrifuge tube on a magnetic rack and add 50 μL of Protein G Dynabeads to the tube. 2. Add 500 μL PBST to the bead matrix and wash the matrix a total of three times in this volume of PBST. Remove the tube from the rack during each wash and gently invert the tube several times to wash thoroughly. 3. Use the magnetic tube rack to sediment the beads and carefully remove the PBST making sure not to remove any of the bead resin (see Note 5). 4. Add 5 μg of the monoclonal anti-D2R (in 500 μL PBST) onto the bead matrix. 5. Incubate on a rocking platform for 20 min at room temperature (see Note 6). This will allow the antibodies to bind to the beads (Fig. 1a). 6. Place the tube on the magnetic rack for 1 min and carefully remove the supernatant making sure not to disturb the bead matrix.

4

Proteomics of D2 Receptors

51

Fig. 1. Major steps in the immunoprecipitation of the D2R complex using the batch method. (a) Immobilization of the antibody bait on a bead matrix: Prewashed agarose Protein A/G Dynabeads are incubated with purified monoclonal antibodies. Protein A/G selectively binds to the IgG region of the antibody. (b) The antibody–bead matrix is incubated with a solution of solubilized membrane proteins expressing D2R complexes. D2R complexes (D2R + DRIPs) are then selectively immunoprecipitated from the solution.

7. Wash the beads with 500 μL PBST a total of three times as described above in steps 2–3. 8. To immunoprecipiate the D2R protein complex, add 100 μg of solubilized membrane proteins from primary striatal cultures and 50–100 μg of solubilized membrane proteins from striatal tissue to the bead matrix (proteins obtained from Subheadings 3.1 and 3.2). 9. For immunoprecipitation controls, we highly recommend performing the same procedures in mice lacking the D2R in order to gain confidence in the specificity of the anti-D2R antibody and the results of the immunoprecipitation procedure. 10. Adjust the total volume of each sample tube to 500 μL using PBST.

52

N. Kabbani and J.C. Nordman

11. Place the tube on a rocking platform for 20–30 min at room temperature to allow the bead complex to bind D2Rs and their associated proteins (Fig. 1b). Alternatively, it is possible to incubate the immunoprecipitation overnight at 4°C on the same rocking platform. 12. Use the magnetic tube rack to carefully remove unbound supernatant and isolate receptor–antibody–bead complex. 13. Thoroughly wash the bead complex with 500 μL PBST a total of five times (with 1 min incubation on ice in between washes) (see Note 7). 14. Make sure to gently invert the tube to assure complete washing at every step. 15. To elute the protein complex from the Protein G Dynabeads, add 30 μL of a premixed elution buffer containing 25% LDS, 10% reducing agent (Invitrogen) and 65% room temperature PBS to the bead matrix. 16. Alternatively if mass spectrometry is necessary use an elution buffer consisting of 49% acetonitrile + 2% trifluoroacetic acid in deionized water, and proceed as below. 17. Gently pipette the elution solution with the bead matrix several times to make sure they mix well. 18. Incubate the beads in the elution buffer for 10 min at 70ºC. This should dissociate the antibody–receptor complex from the Dynabeads. 19. Place the solution back onto the magnetic rack and aim to recover the entire eluted solution sparing the Dynabeads from the final eluant. 20. At this point you can analyze your eluant sample using mass spectrometry or a western blot method (Fig. 2). 3.4. Mass Spectrometry Identification of Dopamine Receptor Interacting Proteins

Immunoprecipitated samples must be processed for mass spectrometry analysis. Depending on the ion source, different mass spectrometry methods require the sample to be processed into various concentrations, volume, and composition. In addition, for mass spectrometry the protein must be cleaved into peptides before analysis using either in-gel digestion or proteolysis in solution procedures. Described here is the method we have used for the preparation of the immunoprecipitated D2R complex for LC-ESI mass spectrometry using an LTQ Orbitrap (Thermo Fisher). This instrument was coupled to a computer that ran the SEQUEST software (BioWorks software from Thermo Fisher, 3.3.1) in order to identify the proteins in the NCBI protein database. This procedure is summarized in Fig. 3.

4

Striatal cell culture

Membrane protein preparation & solubilization

Proteomics of D2 Receptors

53

Striatal tissue

Protein crosslinking in situ

Membrane protein preparation & solubilization

D2R Protein Complex Immunoprecipitation

LC-ESI/ MALDI-TOF Mass spectrometry

SDS-PAGE electrophoresis Gel Extraction

Reverse crosslinking

Western Blot Confirmation of Interactions

Identification of Interacting Proteins

D2R Protein Complexes Fig. 2. A flow chart showing proteomic strategies for the identification of the D2R complexes from cultured cells and native brain tissue.

1. Reduce the eluted D2R complex by incubating the entire sample in 10 mM DTT, 20 ng of β-casein, and 8 M urea for 1 h at room temperature. 2. Since protein identification is enhanced by protein abundance, we strongly recommend using most, if not the entire, immunoprecipitated sample from an experiment for mass spectrometry analysis.

54

N. Kabbani and J.C. Nordman

Fig. 3. Identification of proteins that coimmunoprecipitate with the D2R using mass spectrometry. (a) Enzymatic digestion and processing of immunoprecipitated D2R complexes. (b) LC separation of peptide fragments using a C18 chromatography column. The sample is then emitted towards the sensor for ESI mass spectrometry. Ions are fragmented by collision induced dissociation (CID). (c) Tandem mass spectrometry ESI yields a mass to charge (m/z) ion spectra. (d) The fragment ion spectra are assigned peptide sequences based on database comparison using SEQUEST software. An example of D2R interacting proteins identified using this approach. (e) Mass spectrometry proteomics can be used to generate data for determining D2R signaling and interactions within cells.

3. Alklyate the sample with 50 mM iodoacetamide for 20 min in the dark at room temperature. 4. Enzymatically digest sample with 0.5 μg sequencing grade trypsin in 50 mM NH4HCO3 (pH 8) at 37°C overnight. 5. Quench the trypsin digestion using 2 μL of 98–100% grade glacial acetic acid. 6. For detection of phosphorylated proteins, add 500 fmol Angiotensin II to the sample. If phosphorylated proteins are not of concern, skip this step. 7. Desalt samples using C-18 elution Zip-Tips (Millipore). 8. Load 20μL of the solution into one Zip-Tip at a time. 9. Activating Zip-Tips with Zip-Tip elution buffer 2× (see Note 8). 10. Wash Zip-Tip in Zip-Tip washing buffer.

4

Proteomics of D2 Receptors

55

11. Repeat the washing three times. 12. Elute with Zip-Tip elution buffer, leaving a final eluted volume of 20–30 μL (see Note 9). 13. Dry sample in a SpeedVac concentrator until final volume is approximately 5 μL (see Note 10). 14. Reconstitute dried samples in 1–5 μL of 0.1% Formic Acid. 15. Add 100 fmol Angiotensin I (see Note 11). 3.5. Cross-linking of Protein Complexes from Cells of Culture or Tissue Origin

It is possible to cross-link protein complexes thereby capturing a “snapshot” of the receptor interaction network in space and time during an experiment. A number of cross-linking tools have been developed that complement the range of information that can be gained by proteomic analysis. Here we describe three main approaches that can significantly aid in the study of D2R complexes within various cell types.

3.5.1. DSP Cross-linking of D2R Complexes in Live Cells

This method allows for DSP cross-linking of proteins in primary cultures or brain tissue (Fig. 4). Because DSP is membrane-permeable, it has been used to cross-link proteins within living cells under various experimental procedures (19, 20). Below we describe a method for DSP cross-linking within cultures of primary striatal neurons. This method can be used for the detection of D2R protein complexes in living cells under various experimental conditions. Once cross-linked, the proteins can be directly visualized on an SDS-PAGE gel using a Coomassie stain. In addition, crosslinked proteins can be identified using a western blot or a mass spectrometry method. 1. Prepare a 10 mM DSP stock solution by dissolving DSP into DMSO. 2. Dilute the DSP stock solution into PBS (1:10 dilution) to create a working solution of 1 mM DSP. 3. Gently remove the culture medium from cultured striatal neurons and wash the cells once with 5 mL of room temperature PBS. 4. Gently apply 3 mL of the 1 mM DSP solution into each dish of cells. We recommend using confluent 100 mm petri dishes for protein analysis since protein yield is rate limiting for accurate detection. This volume of DSP is just enough to cover the surface area of the dish; however, higher or lower volumes can be used. 5. Incubate the cells in DSP solution for 2 h at 4°C. We recommend occasional gentle mixing to ensure that the cells are covered in solution throughout the incubation. 6. A white precipitate will form. This is normal.

56

N. Kabbani and J.C. Nordman

Fig. 4. Intracellular protein cross-linking of the D2Rs and their interacting proteins. DSP cross-linking provides a method that facilitates in the identification of weak and/or transient receptor–protein interactions in cells. (a) The chemical structure of DSP promotes its association with free amine (NH2) groups on various proteins. The proteins need to be in close proximity for chemical cross-linking to occur. (b) A model of a cross-linked D2R protein complex at the plasma membrane. Since DSP is membrane-permeable, the D2R complex can be cross-linked within living cells. The DSP cross-linker can be reversed with the addition of DTT thereby providing a tool for studies on the dynamics of the D2R complex under various conditions.

7. End the DSP reaction by diluting the 2× quenching buffer (50 mM Tris–HCl, pH 7.5) to a final concentration of 25 mM Tris–HCl, pH 7.5 for 30 min at 4°C. If you used 3 mL DSP solution per dish, then add the same volume of quenching buffer (3 mL) to the dish. 8. Remove cells from the dish using a cell scraper. 9. Collect cells in a prechilled 15 mL conical Falcon tube and keep cells on ice throughout. 10. Spin cells down at 120 × g for 10 min at 4°C. 11. Discard the supernatant and keep the pelleted cells on ice. 12. Cultured cells can now be manually homogenized and processed for membrane preparation as described in Subheading 3.1. 13. Cross-linked D2R complexes can then be isolated from membrane preparations using the immunoprecipitation protocol

4

Proteomics of D2 Receptors

57

(see Subheading 3.3), and then analyzed by western blot or mass spectrometry (see Subheading 3.4). 14. If using gel electrophoresis, it is important to know that the addition of DTT or β-mercaptoethanol can reverse the DSP cross link. 3.5.2. BS3 Cross-linking of Membrane Proteins

Solubilized membrane proteins derived from either cultured striatal cells or tissue can be cross-linked in vitro prior to immunoprecipitation with an anti-D2R antibody. In this case, cross-linking of proteins, prior to immunoprecipitation, provides a way for preserving weak or transient protein–protein interactions that can be lost during the immunoprecipitation procedure. This method may also be useful in eliminating nonspecific interactions during an experiment. 1. Prepare a 5× BS3 stock solution by dissolving BS3 into ice-cold deionized water to a concentration of 12.5 mM. 2. Prior to the immunoprecipitation experiment (see Subheading 3.3), add BS3 to the membrane protein fraction at a final concentration of 2.5 mM. 3. Incubate the BS3 with the membrane proteins for 2 h at 4°C with gentle mixing. 4. Quench the BS3 reaction by adding 250 mM Tris–HCl, pH 7.5 to a final concentration of 25 mM Tris–HCl, pH 7.5. 5. Mix the solution for 30 min at 4°C with gentle mixing. 6. Cross-linked samples can now be immunoprecipitated as described above (see Subheading 3.3) followed by analysis using western blot or mass spectrometry (see Subheading 3.4).

3.5.3. ReverseCross-linking of Protein Complexes from Samples Immobilized in SDS-PAGE Gel

The use of the membrane-permeable, reversible cross-linker DSP provides a strong tool in the proteomic analysis of signaling pathways in cells (19, 20). Protein complexes can be directly visualized on an SDS-PAGE gel using a Coomasie stain. The bands can then be manually excised and interacting proteins identified using a reverse cross-linking approach. Here we summarize the steps involved in reversing the cross-linking of proteins within an SDSPAGE gel. This technique has proven useful in detecting specific changes within a protein–protein interaction network under various experimental conditions. 1. A Coomassie-stained SDS-PAGE gel loaded can be processed for reverse cross-linking using DTT. 2. To do this, the Coomassie-stained gel should be placed in a plastic container and incubated with a gel in a Destaining Solution for 1–2 h at room temperature, on an orbital shaker at a low speed setting.

58

N. Kabbani and J.C. Nordman

3. Once protein bands become visible, manually excise the bands of interest from the Coomassie-stained SDS-PAGE gel. The bands of interest will be established by the experiment. In our experiments, we have compared immunoprecipitated D2R complexes between cross-linked and non-cross-linked cellular samples. This method has allowed us to identify a series of higher molecular weight bands, within the cross-linked sample, that have been identified as direct binding proteins. 4. Cut the desired gel band into 1 mm × 1 mm pieces, placing individual bands into separate 1.5 mL microcentrifuge tubes. 5. Destain the gel pieces further by adding 500 μL of a Destain Solution to each tube. 6. Incubate the tubes at 37°C for 15 min with gentle mixing on a nutator. 7. If after 15 min the gel piece is still blue, dispose of the Destain solution and replenish with a fresh batch of Destain solution at the same volume as before. 8. Once the gel piece is destained (a light blue color will remain), apply 500 μL of the In-Gel Reducing Solution to each tube. 9. Incubate each tube at 37°C for 30 min with gentle mixing. This is the reverse-cross-linking step. 10. Discard the In-Gel Reducing Solution but slow decanting and then apply 500 μL of In-Gel Alkylating Solution per tube. 11. Incubate the tubes at room temperature for 20 min in the dark with gentle mixing. 12. Discard the In-Gel Alkylating Solution by slow decanting and then add 500 μL NH4HCO3 to each tube. 13. Incubate the tubes at room temperature for 5 min with gentle mixing. 14. Discard the NH4HCO3 then add 500 μL of the Drying Solution. 15. Incubate the tube(s) for 15 min at room temperature over gentle mixing. 16. Remove the Drying Solution by slow decanting and then dry the gel pieces within the microcentrifuge tubes for 20 min using a SpeedVac (at low setting). 17. After drying, place the tubes on ice then add 50 μL of In-Gel Digestion Solution. 18. Incubate the samples for 30 min at 4°C. 19. Transfer the tubes into a 37°C water bath and incubate the samples overnight. This should free most of the proteins from the gel matrix.

4

Proteomics of D2 Receptors

59

20. The following day pellet the gel pieces using a quick spin at room temperature. 21. Transfer the supernatant (S1) to a separate 1.5 mL microcentrifuge tube. 22. Add 30 μL of the In-Gel Extraction Buffer to the remaining gel pieces and incubate for 15 min at room temperature. 23. Pellet the gel pieces using a quick spin. 24. Obtain the supernatant (S2) and combine with S1. 25. SpeedVac the pooled fractions (S1 + S2) at a slow speed to a volume of 20 μL (see Note 12). 26. Proceed with mass spectrometric analysis as previously described (see Subheading 3.4) to identify interacting proteins within the cross-linked complex.

4. Notes 1. Primary striatal cultures are naturally composed of neuronal and non-neuronal cells. Since D2Rs are expressed in glia, as well as neurons, we have added 10 μM fluoro-5¢-deoxyuridine (Sigma) during the first 4 days of culturing in order to minimize glial proliferation. 2. DMEM medium must be room temperature when being introduced to cells. DMEM viability is colorometric: fresh DMEM is red; overtly basic (high pH) DMEM is purple; overtly acidic (low pH) DMEM is yellow. 3. Cells should detach from surface with light pressure. Make sure not to apply too much force as this could damage cell integrity. 4. For optimal results, ratio of cells to non-denaturing protein extraction buffer should be 1:2. 5. Taking care to avoid disturbing beads or bead–antibody– protein complex to ensure that the immunoprecipitated sample is consistent between different experimental groups. 6. For the immunoprecipitation incubation steps, a nutating mixer works well. 7. During the last wash and before the elution of protein complexes, a quick centrifugation step is recommended. 8. Avoid creating bubbles when desalting as this compromises the integrity of the filter. 9. This can be accomplished by simply resuspending the Zip-Tip sample in the same 20 μL volume of Zip-Tip elution buffer.

60

N. Kabbani and J.C. Nordman

10. SpeedVac time using a 20 μL volume is about 3–5 min. 11. Samples can be frozen at this point for up to 1 week before processing; however, immediate proteomic analysis is most preferable. 12. SpeedVac time should be about 5 min. References 1. Missale C, Russel N, Robinson SW, Jaber M, Caron MG (1998) Dopamine receptors: from structure to function. Physiol Rev 78:189–225 2. Sidhu A, Niznik HB (2000) Coupling of dopamine receptor subtypes to multiple and diverse G proteins. Int J Dev Neurosci 18:669–677 3. Binda AV, Kabbani N, Levenson R (2005) Regulation of dense core vesicle release from PC12 cells by interaction between the D2 dopamine receptor and calcium-dependent activator protein for secretion (CAPS). Biochem Pharmacol 69:1451–1461 4. Goldman-Rakic PS (1998) The cortical dopamine system: role in memory and cognition. Adv Pharmacol 42:707–711 5. Schultz W (2002) Getting formal with dopamine and reward. Neuron 36:241–263 6. Strange PG (2001) Antipsychotic drugs: importance of dopamine receptors for mechanisms of therapeutic actions and side effects. Pharmacol Rev 53:119–133 7. Obidiah J, Avidor-Reiss T, Fishburn CS, Carmon S, Bayewitch M, Vogel Z, Fuchs S, Levavi-Sivan B (1999) Adenylyl cyclase interaction with the D2 dopamine receptor family; differential coupling to Gi, Gz, and Gs. Cell Mol Neurobiol 19:653–664 8. Neve KA, Seamans JK, Trantham-Davidson H (2004) Dopamine receptor signaling. J Recept Signal Transduct Res 24:165–205 9. Kabbani N, Levenson R (2007) A proteomic approach to receptor signaling: molecular mechanisms and therapeutic implications derived from discovery of the dopamine D2 receptor signalplex. Eur J Pharmacol 572:83–93 10. Bertorello AM, Hopfield JF, Aperia A, Greengard P (1990) Inhibition by dopamine of (Na+ + K+) ATPase activity in neostriatal neurons through D1 and D2 dopamine receptor synergism. Nature 347:386–388 11. Bergson C, Levenson R, Goldman-Rakic PS, Lidow MS (2003) Dopamine receptor-interacting proteins: the Ca(2+) connection in

12.

13.

14.

15.

16.

17.

18.

19.

20.

dopamine signaling. Trends Pharmacol Sci 24:486–492 Downard KM (2006) Ions of the interactome: the role of MS in the study of protein interactions in proteomics and structural biology. Proteomics 6:5374–5384 Baerenfaller K, Grossman J, Grobei MA, Hull R, Hirsch-Hoffmann M, Yalovsky S, Zimmermann P, Grossninklaus U, Gruissem W, Baginsky S (2008) Genome-scale proteomics reveals Arabidopsis thaliana gene models and proteome dynamics. Science 320: 938–941 Hinkson IV, Joshua EE (2011) The dynamic state of protein turnover: it’s about time. Trends Cell Biol 21(5):293–303 Santos SD, Manadas B, Duarte CB, Carvalho AL (2010) Proteomic analysis of an interactome for long-form AMPA receptor subunits. J Proteome Res 9:1670–1682 Xiao K, McClatchy DB, Shukla AK, Zhao Y, Chen M, Shenoy SK, Yates JR 3rd, Lefkowitz RJ (2007) Functional specialization of betaarrestin interactions revealed by proteomic analysis. Proc Natl Acad Sci U S A 104: 12011–12016 Free RB, Hazelwook LA, Cabrera DM, Spalding HN, Namkung Y, Rankin ML, Sibley DR (2007) D1 and D2 dopamine receptor expression is regulated by direct interaction with the chaperone protein calnexin. J Biol Chem 282:21285–21300 Bradford MM (1976) Rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein dye binding. Anal Biochem 72:248–254 Ollom CM, Denny JB (2008) A crosslinking analysis of GAP-43 interactions with other proteins in differentiated N1E-115 cells. Int J Mol Sci 9:1753–1771 Thomas DDH, Taft WB, Kaspar KM, Groblewski GE (2001) CRHSP-28 regulates Ca2+ stimulated secretion in permeabilized acinar cells. J Biol Chem 276:28866–28872

Chapter 5 Modeling Spatial Aspects of Intracellular Dopamine Signaling Kim T. Blackwell, Lane J. Wallace, BoHung Kim, Rodrigo F. Oliveira, and Wonryull Koh Abstract Dopamine binding to various dopamine receptors activates multiple intracellular signaling molecules, some of which interact with calcium activated signaling pathways. Many experiments measure agoniststimulated elevations in signaling molecules using prolonged, diffuse application, whereas the response of neurons to transient and spatially localized stimuli is more important. Computational modeling is an approach for investigating the spatial extent, time course, and interaction of postsynaptic signaling molecules activated by dopamine and other transmembrane receptors. NeuroRD is a simulation algorithm which can simulate large numbers of pathways and molecules in multiple spines attached to a dendrite. We explain how to gather the information needed to develop computational models, to implement such models in NeuroRD, to perform simulations, and to analyze the simulated data from these models. Key words: Computer model, Signaling pathways, Dopamine signaling, Reactions, Diffusion

1. Introduction Dopamine binding to various dopamine receptors activates multiple intracellular signaling molecules. The importance of dopamine activated signaling pathways is evident from Parkinson’s disease, a degenerative disease in which dopamine neurons of the substantia nigra pars compacta degenerate. The lack of dopamine input from the substantia nigra pars compacta to the basal ganglia produces abnormalities in voluntary motor activity (1). In vivo recordings have shown that the lack of dopamine alters the firing dynamics of these neurons and the striatum as a whole (2). In vitro experiments have shown that

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_5, © Springer Science+Business Media, LLC 2013

61

62

K.T. Blackwell et al.

dopamine modulates several voltage dependent and synaptic channels of striatal spiny projection neurons, and that dopamine activated signaling pathways interact with glutamate and acetylcholine activated pathways to produce neuromodulation (3). Most in vitro experiments demonstrating the effect of dopamine on signaling pathway molecules or ionic channels use prolonged, diffuse application of agonists. Because of adaptations such as receptor desensitization, inferences made from prolonged, diffuse application might not accurately represent the effect of transient “in vivo like” inputs. Computational modeling is an approach for investigating the spatial extent, time course and interaction between dopamine activated and other signaling pathways (4). Computational modeling allows integration of information obtained from biochemical, pharmacological, and molecular biology experiments with information obtained from electrophysiology and calcium imaging experiments. Simulation experiments using the resulting integrative model provides insight into mechanisms underlying neuromodulatory effects in response to realistic synaptic inputs, and can generate testable hypotheses.

2. Materials Several software packages are available for modeling signaling pathways (5–8). They differ in terms of user interface and simulation algorithm. The computational efficiency of the various algorithms depends both on the level of biophysical detail and the numerical approximations made. The computational efficiency then constrains the spatial scale and complexity of the signaling network that can be practically investigated. The example provided here uses NeuroRD (5), which has a simple, text-based user interface, and is freely available (http:// krasnow1.gmu.edu/CENlab/software.html). The simulation algorithm can simulate large numbers of pathways and molecules in multiple spines attached to a dendrite. The model is defined in a set of files, each specifying a different attribute of the model: morphology, reaction, initial conditions, stimulation protocol, and output, and one additional file which specifies the name of the other files. The following procedure explains how to gather the information needed to create the model files, and how to run and analyze the resulting simulation output.

5

Modeling Dopamine Signaling

63

3. Methods 3.1. Signaling Pathways 3.1.1. Identify Bimolecular and Enzymatic Reactions that Form the Signaling Pathways of Interest 3.1.2. Find Measures of Rate Constants from Various Biochemical Experiments

The set of reaction pathways is typically determined from multiple experiments, some of which demonstrate that blocking specific molecules prevents a downstream effect, and some of which demonstrate that two substrate molecules interact to create a product. Perusal of published literature might be aided by consultation of online databases. 1. Assays of bimolecular reactions typically provide KD = KB/KF. If separate estimates of KB or KF are not provided, it is necessary to estimate the rates of forward and reverse reactions. 2. Enzyme reactions consist of a pair of reactions in which the enzyme and substrate form a complex in the first reaction and free enzyme is regenerated in the second reaction. Under certain conditions (constant enzyme concentration, irreversible), which usually apply in enzyme assays, product formation is given by Eq. 1, K cat ·S dP = dt S + K M

(1)

where KM = (KB + Kcat)/KF. Assays typically measure enzyme activity at various substrate concentrations and provide KM and Kcat values (9). When a separate estimate of KB is not provided, it is assumed that KB = 4⋅Kcat (10). 3. One method of estimating rate constants is to create simulations of experimental assays and adjust the rate constants in the assay simulations until the output is a reasonable match to experimental assay data. Experimental assays that show dose response and/or time course relationships are the best. After choosing the experiment assays to simulate, a separate model is created to match the conditions of the assay. Enzyme assays are usually run for many minutes to provide time to accumulate sufficient substrate to measure. If conditions are such that product accumulation is a linear function of time, a few seconds of simulation will yield the same enzymatic rate as many minutes of simulation. In using results from simulations that run for a few seconds, one must be certain that the system is at steady state for all species except reaction substrate and product. Typically, there are many sets of rate constant values that will produce a simulation output that matches experimental data for a single assay. Although the same simulation output is obtained from these various sets of values, the time for various species in the system to reach steady state will be different.

64

K.T. Blackwell et al.

If one can find several different kinds of time course experiments to simulate, these can be used to constrain the final choice of rate constants. 4. Constants describing interactions of drugs with various components of the system can be estimated from functional data. When the concentration of agonist in an assay is low, the IC50 concentration for an antagonist is often a reasonable estimate of the dissociation constant for antagonist binding. If the concentration of agonist is higher and the agonist’s dissociation constant is known, then the Cheng Prusoff equation (11, 12) is used to calculate the dissociation constant: KI = IC50/ (1 + (substrate/KS)), where KS is the dissociation constant for the substrate. For agonists, the situation is more complex: because of signal amplification following interaction with receptor, EC50 values are often substantially lower than dissociation constants. Thus, EC50 values represent a lower boundary on the possible range of dissociation constants for an agonist. Dissociation constants also can be obtained from radioligand binding data. The vast majority of the radioligand binding studies reported involve equilibrium binding methodology. These provide excellent data relative to affinity of interactions but do not provide information on rates of association and dissociation. Although association and dissociation studies can be done using radioligand binding methods, they are not commonly used. 5. Several databases tabulate rate constants and literature sources of these rate constants (see Note 1). The modeler is always encouraged to look into the original references and protocols when trying to understand the source of possible discrepancies. 3.1.3. Find Measures of Diffusion Constants

In some cases the diffusion constant of a molecule in the cytosol is available. In most cases, it is necessary to estimate diffusion constant from molecular weight and viscosity of cytosol using Eq. 2 (5) D = 8.34e −8 ·T η·M 1/3

(2)

where the diffusion coefficient D is in cm2/s, T is temperature in K, the solution viscosity, η, is in cP (estimated at 1.2–1.4 for cytosol), and molecular weight M is in g/mol. 3.1.4. Create the ReactionScheme File that Lists Molecules, Diffusion Constants, and Reactions

The reaction file has two parts. The first part lists all molecular species. Each molecular species is defined by four attributes: name, id, diffusion constant, and units for the diffusion constant. The second part of the reaction file lists all reactions (Fig. 1). Each reaction has six attributes: reaction name, id, reactant, product,

5

Modeling Dopamine Signaling

65

Fig. 1. DopamineReac.xml. ReactionScheme file listing molecules, diffusion constants, and reactions.

forward reaction rate, and reverse reaction rate. Either one or two reactants and products can be specified. Enzyme reactions are specified as two bimolecular reactions, with the enzyme regenerated in the second step. Additional details are found in the README file available on the Web site. For example, when the D1 type of dopamine receptor binds to dopamine, the receptor becomes an active enzyme that catalyzes the activation of the Gαolf variety of G protein. The GαolfGTP activated subunit binds to and

66

K.T. Blackwell et al.

activates adenylyl cyclase, which catalyzes the production of cAMP from ATP. The following reactions describe the first steps of this pathway: Da + DIR  DaDIR DaDIR + G  G − DaDIR → DaDIR + Ga Olf GTP Ga Olf GTP → Ga Olf GDP Reactions are represented by the ReactionScheme file (Fig. 1). Each enzymatic reaction requires specification of two reactions; thus forward Rate in the third reaction is turnover or catalytic rate for the enzymatic reaction. 3.2. Morphology 3.2.1. Describe the Morphology of the Neuron to be Modeled

At present, due to computational burden, only a part of the neuron can be simulated in a reasonable time, e.g., a dendrite with several spines, or several dendrites. The morphology of neurons is available from many publications and various databases. Neuromorpho. org provides morphology files of many different neuron types from many species and brain regions in a standard format in which a traced neuron is given as a set of points (x, y, z coordinates) with associated radii. Alternatively, the morphology of whole neurons or parts of neurons is often available from immunocytochemistry figures in publications. Lengths and radii of discrete segments can be extracted from the figures. For simulations of experimental assays, it is advisable to use a single large compartment.

3.2.2. Create the Morphology File

The NeuroRD morphology file (Fig. 2) specifies the shape of the neuron by specifying the shape of all segments and how they are connected. Each segment has four attributes: id, region, start and end. The id attribute is required and must be unique whereas the optional region attribute does not have to be unique. Regions are used to group segments with the same initial conditions. The start and end attributes are specified using x, y, z, (r)adius, and label. The label is optional and can be used as a site where molecules are injected into the system. In general, segments are specified with the starting x, y, z coordinates and radius, and the ending x, y, z coordinates and radius. To connect the second segment to the first, it must start from the endpoint of the first compartment using the attribute start on (Fig. 2) (see Note 2).

3.2.3. Include Spine Specifications in the Morphology File

SpineType specifies a spine prototype and SpineAllocation (Fig. 2) applies the spine prototype to the surface of a structure. This allows for random placement of spines according to a specified density in a constrained region or segment of the defined morphology. Multiple spine prototypes can be defined, e.g., to randomly distribute long, thin spines among short, stubby spines. SpineType has an id attribute and is defined using multiple section elements,

5

Modeling Dopamine Signaling

67

Fig. 2. DopamineMorph.xml. Specifications in the morphology file. An example of two dendrites attached to the soma, with spines on both dendritic branches.

which have four attributes: width, at, regionClass, and label. The width attribute specifies the radius of that section and the at attribute indicates the distance from the dendrite at which that radius begins to apply. The regionClass and label attributes are optional. SpineAllocation has four attributes: id, spineType, region, and lengthDensity. The region attribute indicates the region to which spines of type spineType will be added. The lengthDensity attribute is the average number of spines per micron. Alternatively, areaDensity, which is the number per unit area, can be used instead of lengthDensity (Fig. 2). 3.3. Molecule Quantities and Initial Conditions

1. Find measures or estimates of molecule quantities and, for nondiffusible molecules, their subcellular distribution. For example, in spiny neurons which have glutamatergic synapses on the spines, metabotropic glutamate receptors are located in the spine head. Plasma membrane pumps are located in the plasma membrane and do not diffuse through the cytosol. Once control conditions are selected, it is possible to determine changes in molecule quantities that will be used to run simulation experiments. For example, to simulate pharmacological block of a particular molecule, its value could be set to zero.

68

K.T. Blackwell et al.

Then, results of the control simulation and the blocked simulation can be compared to evaluate the role of that blocked molecule. 2. For quantities that are not available, use experiments that measure time course of a molecule in the signaling pathway as a constraint. For example, if the amount of substance produced by a stimulation is known and this substance is produced by an enzyme that has been purified and tested so that its Kcat is known but its concentration in the tissue of interest is not known, one can calculate the minimum concentration of enzyme that must be present to produce the response. Also, sufficient substrate must be present to generate the products of enzymatic reactions. The location of various molecules often is determined from immunogold labeling (13) or subcellular fractionation techniques (14). 3. These values are used to create the InitialConditions file, which specifies the starting concentration or density for each molecule. The file must contain one general ConcentrationSet, which applies to everything unless overridden. Each NanoMolarity element names the species and provides a value for its concentration, entered in nanoMoles per liter. In addition, non-diffusing molecules can be assigned to a specific region using additional ConcentrationSets together with a region attribute. This should correspond to a specified region from the morphology file and indicates the parts of the structure to which the initial conditions apply. For membrane localized molecules, it is possible to specify initial conditions as a density (picomoles per square meter) using the SurfaceDensitySet which places these molecules only in the submembrane areas of the morphology. If the region attribute is specified, then that submembrane initial condition applies only to that region. If the region attribute is not specified, then the initial condition applies to all submembrane areas. In Fig. 3, the dopamine receptor (D1R) density is zero in general, but non-zero in the submembrane part of the dendrite region. 3.4. Stimulation

1. Determine the pattern of stimulation for a particular experiment. For example, a brief electrical stimulation leads to release of dopamine from terminals (15), which then binds to postsynaptic dopamine receptors. Sometimes, a molecule downstream from the experimental stimulation can be used for the model stimulation, such as with NMDA type of glutamate receptors. In this case each glutamate release event and binding to receptors may be approximated as influx of calcium, and stimulation at high frequency would produce accumulation of individual calcium transients. Note that neither presynaptic terminal nor diffusion of dopamine in the extracellular space can be modeled presently.

5

Modeling Dopamine Signaling

69

Fig. 3. DopamineIC.xml. Receptor quantities and initial conditions.

Fig. 4. DopamineStim.xml. StimulationSet file specifying time and location for injection of molecules during a simulation.

2. The StimulationSet file specifies the time and location for (optional) injection of molecules during a simulation (Fig. 4). Each InjectionStim specifies the molecule injected and its site of injection. Additional required attributes include onset (in ms), duration (in ms) and rate (particles/ms). Optional attributes, period and end, can be used to specify a train of input. Thus, 100 Hz stimulation for 1 s would be specified with a period of 10 ms, and end equal to onset plus 1,000 ms. Multiple trains are possible with two more optional attributes: interTrainInterval and numTrains. Variations in stimulation protocols, e.g., brief, large amplitude (1,000 particles during 1 ms) versus long, low amplitude (10 particles during 100 ms) dopamine release, are used to run simulation experiments. Stimulation can be delivered to a single spine or to a range of spines as explained in more detail at the Web site.

70

K.T. Blackwell et al.

3.5. Output

1. To evaluate the result of the simulation it is necessary to analyze the concentration or quantity of one or more molecules from one or more parts of the neuron morphology. For example, if studying synaptic plasticity, then quantity of activated kinase, calcium concentration, or phosphorylation state of particular target molecules may be of interest. To understand mechanisms underlying the simulation results, it may be necessary to evaluate the time course of several molecular species in the model. One advantage of simulations is the possibility to examine every molecular species. 2. The OutputScheme file specifies the file to which output is written and which molecules from which compartments are written to the file at which frequency (time interval). Multiple output files are allowed: for example, information about some molecules, or information from some regions can be written more frequently than others. Every OutputSet specifies a single output file using two required attributes, filename and dt (for data output interval), and an optional attribute, region. If the region attribute is not specified, then the concentrations (or number of particles) for all subvolumes in the system are saved. Each OutputSpecie in the OutputSet indicates a molecule to be included in that output file (Fig. 5).

3.6. Additional Details on the Simulation

One additional file, called the Model file (Fig. 6), serves to identify all these other files and provides additional information such as discretization options, simulation seed(s) and various control parameters. This model file is input to the software to run the simulation, as explained in Subheading 3.7. 1. Discretization options For numerical calculations within the model, each of the morphology segments is subdivided into smaller compartments called subvolumes. The discretization element indicates the size of these subvolumes. Smaller sizes produce larger number of subvolumes which require more calculations and result in a longer run time. The defaultMaxElementSide specifies the

Fig. 5. DopamineIO.xml.

5

Modeling Dopamine Signaling

71

Fig. 6. DopamineModel.xml.

largest size (in microns) for each side of the subvolume in each segment. This is the default, and can be overridden using MaxElementSide with the region attribute: the value supplied will control the size of subvolumes for that region. Similarly, spineDeltaX specifies the size of subvolumes in spines. Spines have a one-dimensional discretization along the spine axis. The geometry element is used to specify how the morphology is interpreted. 2D implies that there are multiple voxels in x and y dimensions, but only a single layer of subvolumes in the z dimension. Thus, the morphology is three-dimensional, but diffusion occurs in two dimensions only. For 2D, the user also specifies the depth of the subvolumes.

72

K.T. Blackwell et al.

2. Additional parameters. Runtime is used to specify run time in milliseconds. fixedStepDt specifies the time step, in milliseconds, which must be smaller than the fastest reactions or diffusion processes. The required simulationSeed specifies the seed for the random number generator. In morphologies with spines, a separate seed, spineSeed, is used to randomly place spines within the specified region. outputQuantity specifies whether quantity of molecules in the output is in number of molecules or concentration. 3.7. Run Simulations and Evaluate Output 3.7.1. Experimental Design

3.7.2. Running Simulations

The key to meaningful experiments is asking good questions that can be answered using data from the simulations. Simulations are excellent tools for addressing questions such as the effect of morphology and spatial constraints, or the conditions which would be required to produce an experimentally observed response. Similar to experiments, to address particular effects, it is necessary to run two or more simulations which differ only in the desired parameter. For example, the effect of morphology on output may be evaluated by performing two simulations which differ only in their morphologies. To evaluate the effect of an antagonist requires a control simulation (the default simulation with no antagonist) and a simulation either with antagonist added (simulating competitive or allosteric binding) or, alternatively, with the antagonist target concentration close to zero. 1. To run a simulation from the command line in Unix, the following command should be issued: java -jar NeuroRD.jar DopamineModel.xml Dopamine.out >>Dopamine.log 2. To run a simulation in Windows, type “cmd” in the run command from the start menu to obtain a command line window. Then, type the following: java -jar NeuroRD.jar DopamineModel.xml Dopamine.out 3. The NeuroRD.jar file is the simulation software downloaded from the Web site (http://krasnow1.gmu.edu/CENlab/ software.html), DopamineModel.xml is the model file (“master” file that specifies the other files), and Dopamine.out is the main output file. 4. The name of additional output files are composed of the main output file name plus the filename specified in OutputScheme file.

3.7.3. Evaluating Simulation Output

The first set of simulations typically is used to validate the model; thus simulation output is compared to experimental data. Due to stochastic variability, multiple simulation runs must be performed to generate several observations, analogous to experiments which require multiple trials or samples. Then, conventional statistical tests are used to compare levels of relevant molecules from simulations to

5

Modeling Dopamine Signaling

73

experimental data. For more quantitative comparison to imaging data, simulations can include fluorescent indicator molecules and calculate the change in fluorescence. Two additional types of simulations are typically performed. One set of simulations implements the experimental design, e.g., evaluating the role of a particular molecule, or the effect of stimulation pattern. Again, multiple simulation runs coupled with statistical analysis are required for hypothesis testing. The other set of simulations addresses the crucial question of how robust the conclusions are. It is important to verify that biological conclusions do not change with small changes in parameter values or different stimulation protocols. The ideal parameters for robustness simulations are those whose experimental measurements span a wide range (i.e., molecule quantities and diffusion constants) and those to which the results are more sensitive. The ability to evaluate dynamics of many different molecules simultaneously, and to perform manipulations that are not possible experimentally such as changing diffusion coefficients, allows evaluating the biochemical mechanisms underlying experimental observations. Evaluating simulation output is challenging due to the amount of numerical data that is generated. In principle, the quantity of every molecular species in the system in every subvolume is available in the numerical experiments. This abundance necessitates additional processing of the data for efficient presentation of these vast numerical results generated from computational modeling. For example, numerical results can be converted to multidimensional visualization (movies) to facilitate understanding and scientific intuitions, e.g., using VNRD which is available on the Web site (http://krasnow1.gmu.edu/CENlab/software.html). Figure 7 shows average concentrations in the dendrite and soma

Fig. 7. Concentration of Dopamine bound D1 receptor and GolfGTP in the dendrite, where it is produced, and in the soma, to which it diffuses. Fluctuations are due to stochastic nature of simulation.

74

K.T. Blackwell et al.

for the model presented in Figs. 1–6, and illustrates that the concentration of GolfGTP (diffusible in the cytosol for the sake of illustration) increases first in the dendrite (the location of the dopamine receptors), and subsequently in the soma. The average concentrations were generated by VNRD.

4. Notes 1. These databases can be found at: http://doqcs.ncbs.res.in http://www.brenda-enzymes.info http://www.cyclic-nucleotides.org 2. Subsequent segments, which do not start on an existing segment, will not be connected. 3. Branching segments are made by creating two segments beginning at the same site but terminating at different points. 4. The two branches need not have the same radius at the point of connection. In this case, a radius value must be specified.

Acknowledgements This work was supported by the NIH-NSF CRCNS program on Collaborative Research in Computational Neuroscience through NIH grants R01 AA18060 and R01 AA16022. References 1. Wichmann T, DeLong MR (1998) Models of basal ganglia function and pathophysiology of movement disorders. Neurosurg Clin N Am 9:223–236 2. Goldberg JA, Rokni U, Boraud T, Vaadia E, Bergman H (2004) Spike synchronization in the cortex/basal-ganglia networks of Parkinsonian primates reflects global dynamics of the local field potentials. J Neurosci 24:6003–6010 3. Kitai ST, Surmeier DJ (1993) Cholinergic and dopaminergic modulation of potassium conductances in neostriatal neurons. Adv Neurol 60(40–52):40–52 4. Kotaleski JH, Blackwell KT (2010) Modelling the molecular mechanisms of synaptic plasticity using systems biology approaches. Nat Rev Neurosci 11:239–251

5. Oliveira RO, Terrin A, Di Benedetto G, Cannon RC, Koh W, Zaccolo M, Blackwell KT (2010) The role of type 4 phosphodiesterases in generating microdomains of cAMP: large scale stochastic simulations. PLoS One 5:e11725 6. Andrews SS, Addy NJ, Brent R, Arkin AP (2010) Detailed simulations of cell biology with Smoldyn 2.1. PLoS Comput Biol 6:e1000705 7. Byrne MJ, Waxham MN, Kubota Y (2010) Cellular dynamic simulator: an event driven molecular simulation environment for cellular physiology. Neuroinformatics 8:63–82 8. Kerr RA, Bartol TM, Kaminsky B, Dittrich M, Chang JC, Baden SB, Sejnowski TJ, Stiles JR (2008) Fast Monte Carlo simulation methods for biological reaction-diffusion systems in

5

9. 10.

11.

12.

solution and on surfaces. SIAM J Sci Comput 30:3126 Stenesh J (1993) Core topics in biochemistry. Cogno Press, Michigan, p 283 Bower JM, Beeman D (1998) The book of genesis: exploring realistic neural models with the GEneral NEural SImulation System, 2nd edn. Springer, New York Cheng Y, Prusoff WH (1973) Relationship between the inhibition constant (K1) and the concentration of inhibitor which causes 50 per cent inhibition (I50) of an enzymatic reaction. Biochem Pharmacol 22:3099–3108 Cer RZ, Mudunuri U, Stephens R, Lebeda FJ (2009) IC50-to-Ki: a web-based tool for converting IC50 to Ki values for inhibitors of

Modeling Dopamine Signaling

75

enzyme activity and ligand binding. Nucleic Acids Res 37:W441–W445 13. Garzon M, Vaughan RA, Uhl GR, Kuhar MJ, Pickel VM (1999) Cholinergic axon terminals in the ventral tegmental area target a subpopulation of neurons expressing low levels of the dopamine transporter. J Comp Neurol 410:197–210 14. Xie Z, Adamowicz WO, Eldred WD, Jakowski AB, Kleiman RJ, Morton DG, Stephenson DT, Strick CA, Williams RD, Menniti FS (2006) Cellular and subcellular localization of PDE10A, a striatum-enriched phosphodiesterase. Neuroscience 139:597–607 15. Rice ME, Cragg SJ (2004) Nicotine amplifies reward-related dopamine signals in striatum. Nat Neurosci 7:583–584

Part II Cellular Imaging

Chapter 6 A Biophysical Approach for the Study of Dopamine Receptor Oligomerization Sylwia Lukasiewicz, Agata Faron-Górecka, and Marta Dziedzicka-Wasylewska Abstract The ability of certain neurotransmitter receptors to form oligomers provides an additional level of fine-tuning of intracellular signaling. Among the techniques allowing study of receptor oligomerization as well as influence of specific ligands on these processes, a biophysical approach with the use of fluorescently tagged receptors is the most sensitive. Measurement of the fluorescence resonance energy transfer (FRET) phenomenon between two fluorescently tagged receptors is considered a very useful and measurable tool to study the physical interactions between receptors either in a single cell or in a population of living cells. Here we describe the use of FRET measurement specifically to monitor protein oligomer formation between dopamine D1R and D2R, but the same methodology can be used to study other receptor proteins as well as their mutants. Key words: FRET, Neurotransmitters, Oligomer formation, D1R, D2R, ECFP, EYFP

1. Introduction The idea of various neurotransmitter receptors forming dimers or higher order oligomers has been well documented (1, 2). The formation of hetero-complexes by certain receptors can take place only if they are co-expressed in the same cell. It has been widely accepted that dopamine D1 and D2 receptors are expressed rather in two separate populations of medium spiny neurons in the striatum, with about 20% of all striatal neurons co-expressing the two receptors (3). However, it has also been disputed that dopamine D1 and D2 receptors are significantly co-localized in the nucleus acumbens septi (4). Such co-localization provides a cellular basis for the interaction of these receptors and may be

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_6, © Springer Science+Business Media, LLC 2013

79

80

S. Lukasiewicz et al.

important for the elucidation of the mechanism of action of certain drugs (e.g., clozapine). Among the techniques allowing studies of receptor oligomerization as well as the influence of specific ligands on these processes, a biophysical approach with the use of fluorescently tagged receptors is the most sensitive. Fluorescence resonance energy transfer (FRET) measurement between two fluorescently tagged receptors is considered a very useful and measurable tool (5–7). FRET measurements allow for the study of physical interactions between receptors either in single cell or in a population of living cells. The FRET phenomenon is observed between a fluorescent donor and an acceptor chromophore when they are located within 100 Å of each other and are arranged properly in terms of their transition dipole moments (8). Here, we describe the use of FRET to monitor protein oligomer formation between dopamine D1R and D2R, but the same methodology can be used to study other receptor proteins as well as their mutants. We labeled dopamine receptors with enhanced cyan fluorescent protein (ECFP, the fluorescence donor) and enhanced yellow fluorescent protein (EYFP, the fluorescence acceptor) and expressed the receptors in human embryonic kidney (HEK) 293 cells. Generally, receptor oligomerization using the biophysical approach is difficult to analyze in native cells. The HEK cell line has been widely used in resonance energy transfer studies of membrane receptors because these cells provide an accepted model in which fluorescently tagged receptor proteins can be efficiently expressed.

2. Materials 2.1. Construction of Fusion Proteins and Genetic Variants of the Dopamine Receptors

1. All molecular biology reagents are obtained from Fermentas (Vilnius, Lithuania). 2. Oligonucleotides (IBB PAN, Warsaw, Poland). 3. pECFP-N1 and pEYFP-N1 vectors (BD Biosciences, Clontech, Palo Alto, CA). 4. pcDNA3.1(+) plasmids encoding human dopamine receptor proteins (UMR cDNA Resource Center, University of Missouri-Rolla, MO). 5. Escherichia coli DH5α (Dam+) (Novagen, EMD Chemicals, Merck, Germany). 6. LB: 10 g/L peptone, 10 g/L NaCl, 5 g/L yeast extract, pH 7.0.

2.2. Cell Culture and Transfection

1. HEK 293 cells (American Type Culture Collection, Manassas, VA).

6

Study of Dopamine Receptor Oligomerization

81

2. HEK 293 cells are grown in Dulbecco’s modified essential medium (DMEM) supplemented with 1% l-glutamine and 10% heat-inactivated fetal bovine serum (FBS). The cells were cultured at 37°C in an atmosphere of 5% CO2. All cell culture materials are purchased from GIBCO (Life Technologies, Grand Rapids, NY) and Sigma-Aldrich (St. Louis, MO). 3. Solution of trypsin: 137 mM NaCl, 5 mM KCl, 10 mM NaHCO3, 0.5 mM EDTA, 20 mM Hepes, 5.5 mM glucose, 0.25% trypsin, pH 7.6. 4. Phosphate-buffered saline (PBS): 140 mM NaCl, 10 mM NaH2PO4, 1.8 KH2PO4, 2.7 mM KCl, pH 7.4. 5. Buffer HBS (transfection): (2×) 280 mM NaCl, 50 mM HEPES, 1.5 mM Na2HPO4, pH 7.05. 6. Solution of calcium chloride (transfection): 2.5 M CaCl2. 2.3. Radioligand Binding Assay

1. Phosphate-buffered saline (PBS): 140 mM NaCl, 10 mM NaH2PO4, 1.8 KH2PO4, 2.7 mM KCl, pH 7.4. 2. Binding buffer: 50 mM Tris–HCl, containing 120 mM NaCl, 5 mM KCl, 4 mM MgCl2, pH 7.4. 3. Washing buffer: 50 mM Tris–HCl, pH 7.4. 4. Dopamine D1R specific radioligand: [3H]SCH23390, specific activity: 86 Ci/mmol (NEN, Boston, MA) (see Note 1). 5. Dopamine D2R specific radioligand: [3H]spiperone, specific activity: 15.7 Ci/mmol (NEN, Boston, MA) (see Note 1). 6. Specific dopamine receptor ligands (Lundbeck) and (+) butaclamol (Sigma).

cis(Z)-flupentixol

7. Membrane protein was determined by Lowry’s modified method: Bicinchoninic Acid Protein Assay Kit and Protein standard, 2 mg BSA/mL (Sigma). 8. Estimation of the radioligand parameters are calculated using the GraphPad Prism 2.0 curve—fitting program (GraphPad Software, La Jolla, CA). 9. Glass test tube (15 mL) (see Note 2). 10. Brandel Cell Harvester. 11. Whatman GF/C filter paper. 12. Scintillation fluid (Aqascynt. BioCare, Guangzhou, China). 13. Liquid scintillation counting (Beckman LS 650). 2.4. FRET and Confocal Measurements

1. All fluorescence spectra are collected using a spectrofluorimeter and 10 mm quartz cuvette (Hellma, Mullheim Germany). 2. Isotonic buffer 1 used as an incubating medium during measurements: 137.5 mM NaCl, 1.25 mM MgCl2, 1.25 CaCl2, 6 mM KCl, 5.6 mM glucose, 10 mM HEPES, 0.4 mM NaH2PO4, pH 7.4.

82

S. Lukasiewicz et al.

3. Specific dopamine receptor ligands (Hartmann Analytic, Sigma). 4. TCSPC methodology and lifetime measurements are the most sensitive for estimation of highly quantitative data of FRET efficiency (see Note 3). 5. Co-localization analysis is done using Image ProPlus 4.5 software.

3. Methods 3.1. Construction of Fusion Proteins

1. The human dopamine receptor genes cloned into the pcDNA3.1(+) plasmid are used as the starting point to construct the fusion proteins. Appropriate genes are tagged with cDNA encoding enhanced cyan or yellow fluorescent proteins (ECFP or EYFP). 2. The full-length cDNAs encoding the dopamine receptor is PCR-amplified. The forward primer is universal for pcDNA3.1 (+), and the reverse primers removed the STOP codon and introduced a unique restriction site, XhoI. The resulting fragment is inserted, in-frame, into the NheI/XhoI sites of the pECFP-N1 and pEYFP-N1 vectors. The obtained fusion proteins constructs are used after expression as the fluorescence donor (receptor-ECFP) or acceptor (receptor-EYFP) (see Note 4).

3.2. Construction of Genetic Variants of the Dopamine Receptors

1. The genetic variants of the dopamine receptors are generated according to the manufacturer’s protocol Quik-Change II Site-Directed Mutagenesis Kit (Stratagene). 2. Dopamine receptor genes inserted into pECFP-N1 and pEYFP-N1 vectors, respectively, are used as the mold for the PCR-Quik reaction. Incorporating the oligonucleotide primers, each complementary to the opposite strand of the vector and containing the desired mutations, generates a mutated plasmid. 3. The resulting product is treated with endonuclease DpnI, specific for methylated and hemimethylated DNA, in order to select synthesized DNA containing the introduced mutations. E. coli DH5α cells are then transformed with the mutated plasmid. All mutated sequences are verified by DNA sequencing.

3.3. Cell Culture and Transfection

1. HEK 293 cells are cultured at 37°C in an atmosphere of 5% CO2. When the confluence is about 90%, the cells are passaged. They are washed with phosphate-buffered saline (PBS) and then treated with trypsin.

6

Study of Dopamine Receptor Oligomerization

83

2. Transient transfections of HEK 293 cells are performed using the calcium phosphate precipitation method, as described by Sambrook et al. (9). HEK 293 cells are transfected with plasmids encoding either ECFP-tagged or EYFP-tagged dopamine receptor proteins separately or co-transfected with different combinations of both ECFP- and EYFP-tagged plasmids. 3. One day before transfection, the cells are seeded into 100 mm dishes at a density of 3 × 106 cells/dish for fluorescence spectra measurements and binding assays. For fluorescence lifetime measurements and confocal imaging cells are plated on glass cover slips in 10-mm dishes at a density of 2 × 105 cells/dish. 4. One hour before transfection cells are incubated with the fresh medium containing 10% FBS in 37°C. 5. The cells are transfected with 15 μg of DNA per 100 mm dish or 2 μg of DNA per 10-mm dish. The ratio of DNA encoding the receptor-ECFP fusion protein (fluorescence donor) to DNA encoding the receptor-EYFP fusion protein (fluorescence acceptor) is 1:1 (see Note 5). 3.4. Binding Assay

Binding of specific radioligands to receptors is an important step since it allows us to determine whether fluorescently tagged receptors display the same binding parameters (Bmax and Kd) as the native receptor. To obtain the information, the saturation analysis is employed. On the other hand, competition analysis is used to determine the affinity (Ki) of the studied receptors for certain ligands, e.g., clozapine. For all binding assays (saturation and competition for D1R and D2R) the steps 1–4 are performed in the same way. 1. The transfected HEK 293 cells are washed with phosphatebuffered saline (PBS), scraped from the dish in PBS and centrifuged at 160 × g for 5 min. The pellet is then frozen at −30°C until used. 2. Frozen pellets are resuspended in the binding buffer (50 mM Tris–HCl, pH 7.4, containing 120 mM NaCl, 5 mM KCl, 4 mM MgCl2, and 1 mM EDTA), using an Ultra Turrax homogenizer (30 s). The homogenates are centrifuged at 30,000 × g for 10 min. The step should be repeated twice (see Note 6). 3. Binding studies are performed on a fresh membrane preparation (final protein concentration usually 20 μg/tube or 40 μg/ tube, for D1 and D2 dopamine receptor, respectively). Membrane protein is determined by the modified Lowry’s method. 4. 10 μM cis(Z)-flupentixol, (Lundbeck, Deerfield, IL) and 50 μM (+) butaclamol (Sigma) are added to determine nonspecific binding for D1R and D2R, respectively.

84

S. Lukasiewicz et al.

Fig. 1. Representative saturation data for (a) [3H] SCH23390 binding to dopamine D1R and fusion protein D1R-EYFP; (b) [3H] spiperone binding to dopamine D2R and fusion protein D2R-ECFP.

3.4.1. Saturation Binding Assay

A saturation binding assay for D1R is performed in 200 μL Tris–HCl buffer (pH 7.4) with 20 μg of membrane homogenate and increasing 12 concentrations (0.06–6 nM) of [3H]SCH23390 (200 μL). For D2R—200 μL Tris–HCl buffer (pH 7.4) with 40 μg of membrane homogenate and increasing 12 concentrations (0.01–4 nM) of [3H]spiperone (50 μL) are used (see Note 7) (Fig. 1). 1. Incubation time for D1R is 90 min. at 25°C, and for D2R is 30 min at 37°C. 2. At the end of the incubation, bound ligands are isolated by rapid filtration through glass fiber filters (GF/C, Whatman). The filters are washed four times with 5 mL of ice-cold washing buffer (Tris–HCl, pH 7.4) (see Note 8). 3. Bound radioactivity is determined by liquid scintillation counting. 4. Estimation of the radioligand binding parameters, Kd (the equilibrium dissociation constant) and Bmax (maximal binding capacity) is calculated using the GraphPad Prism version 2.0 (see Note 9).

3.4.2.Competition Binding Assay

Competition binding studies are carried out under similar conditions to saturation experiments. Competition analysis can be useful

6

Study of Dopamine Receptor Oligomerization

85

to see whether the oligomerization of the studied receptors changes their affinity for the given compound (10). Since the HEK 293 cell line allows us to express various proteins it is also a useful model to study the genetic variants of receptors of interest, either to search for specific regions responsible for receptors oligomerization or to study the impact of certain polymorphisms linked to the diseases. The exchange of Ser-311 residue for Cys in the dopamine D2 receptor sequence is often linked to the susceptibility to schizophrenia. Below the affinity for clozapine is presented of dopamine D1 and D2 receptors as well as of genetic variant, D2Ser311Cys. The results indicate that the value of Ki depends on whether these receptors are expressed alone or simultaneously in the same cell. 1. Competition assays should be done on a fresh membrane preparation (final protein concentration was 20 μg/tube or 40 μg/ tube for D1 and D2 dopamine receptor, respectively) using a fixed concentration of [3H]SCH23390 or [3H]spiperone (see Note 8). 2. One concentration of radiolabeled ligand is used: [3H]spiperone (0.3 nM) (see Fig. 2) and [3H]SCH23390 (1 nM) (see Fig. 3) (see Note 10). The range of nonradioactive ligand (clozapine, Sigma) is from 10−3 to 10−12 M. 3. Tubes are incubated for 90 min at room temperature ([3H] SCH 23390) or for 30 min at 37°C ([3H] spiperone).

Fig. 2. Competition of [3H]SCH23390 with clozapine for binding to membrane preparation from HEK 293 cells transiently transfected with dopamine receptors—representative plots. Two-site binding model was fitted to the data. HEK 293 cells transiently transfected with dopamine D1R (black line ); HEK 293 cells transiently co-transfected with dopamine D1 and D2 receptors (blue line ); HEK 293 cells transiently co-transfected with dopamine D1 and D2S311C receptors (green line ). The estimated Ki values depended on the presence of the second dopamine receptor.

86

S. Lukasiewicz et al.

Fig. 3. Competition of [3H]spiperone with clozapine for binding to membrane preparation from HEK 293 cells transiently transfected with dopamine receptors—representative plots. One- and two-site binding model was fitted to the data. HEK 293 cells transiently transfected with dopamine D2R (black line); HEK 293 cells transiently co-transfected with dopamine D1 and D2 receptors (blue line); HEK 293 cells transiently transfected with dopamine D2S311C receptors (red line); HEK 293 cells transiently co-transfected with dopamine D1 and D2S311C receptors (green line).

4. As in the saturation analysis, binding is terminated with rapid filtration through glass fiber filters (GF/C, Whatman). The filters are washed four times with 5 mL of ice-cold washing buffer (50 mM Tris–HCl, pH 7.4) and bound radioactivity is determined by liquid scintillation counting (Beckman LS 650) (see Note 7). 5. Estimation of the radioligand binding parameter, Ki, is calculated using the GraphPad Prism 2.0 curve-fitting program (GraphPad Software) (see Note 11). 3.5. FRET Measurements

Qualitative as well as quantitative methods are used to monitor whether FRET occurs between the ECFP-tagged protein (donor) and the EYFP-tagged protein (acceptor of fluorescence).

3.5.1. Fluorescence Spectroscopy Measurements

Although steady state fluorescence spectroscopy measurements in cell suspension enable only the qualitative estimation of the FRET phenomenon, this approach is very demonstrative and allows for a quick answer as to whether there is any energy transfer in the examined sample. 1. Spectrofluorimetric measurements of the cell suspensions are recorded at 37°C, 48 h after transfection. Cells (expressing desired combination of receptor fusion proteins) cultured on a single 100 mm dish are washed and detached from the plate

6

Study of Dopamine Receptor Oligomerization

87

Fig. 4. Fluorescence emission spectra of HEK 293 cells expressing the ECFP- and EYFPtagged proteins coupled to D1R and D2R. FRET control, spectra from a 1:1 mixture of cells individually expressing the D1-ECFP fusion protein (the cyan line excited at 434 nm) and the D2-EYFP fusion protein (the yellow line excited at 475 nm). The blue line represents FRET spectrum of HEK 293 transfected with ECFP-EYFP construct. The black line is the spectrum of HEK 293 co-transfected with D1-ECFP and D2-EYFP.

using warm (37°C) PBS buffer. Afterwards, the suspensions are centrifuged at 1,000 rpm for 5 min and resuspended in 1 mL of warm (37°C) isotonic buffer 1. 2. ECFP is excited at 434 nm, and EYFP—at 475 nm. Fluorescence is detected at 450–550 nm through a double monochromator (Fig. 4). The spectral contributions arising from light scattering and nonspecific fluorescence of cells and incubated medium are eliminated by subtracting the emission spectra of mocktransfected cells from the fluorescence spectra of cells expressing the receptor-ECFP and -EYFP constructs. The analysis is done according to Stanasilla et al. (11). 3.5.2. FRET Measurement by Fluorescence Lifetime Microscopy

1. Cells dedicated to TCSPC experiments are grown on cover slips. The fluorescence decay is measured from single living cells transfected with fusion protein constructs. All measurements are performed at 37°C (see Note 12), 48 h after transfection. Cells are incubated in the same isotonic buffer 1 as the one used for fluorescence spectra measurements. For each receptor combination, at least four independent experiments should be performed and during each experiment, fluorescence decay from at least 15 cells on the given cover slip should be measured (Fig. 5). 2. Each fluorescence decay measurement is analyzed with the multiexponential model, given by the equation: I (t ) = ∑ i =1 αi e −t / τi n

88

S. Lukasiewicz et al.

Fig. 5. Time-dependent fluorescence intensity decays of ECFP attached to the D1 receptor with and without EYFP attached to the D2 receptor. The blue dotted curve shows the intensity decay of the donor alone (D), and the green dotted curve shows the intensity decay of the donor in the presence of acceptor (DA). The green and blue solid lines and weighted residuals (lower panels) are for the best double exponential fits. The black dotted curve represents the excitation pulse diode laser profile, set up at 434 nm.

where I(t) stands for fluorescence intensity in time t, αi are pre-exponential factors representing amplitudes of the components at t = 0, τi are the decay times, and n is the number of decay times. Best fit parameters are obtained by minimizing the reduced χ 2 value and residual distribution. The average fluorescence lifetime ‹τ› is calculated from the equation: τ =

∑ α ·τ ∑ α ·τ i

i

2 i

i

i

i

The average efficiency of energy transfer ‹E› is calculated from the average fluorescence lifetime of donor in the absence ‹τD› or presence ‹τDA› of an acceptor from the equation: E =1−

τ DA αD

6

Study of Dopamine Receptor Oligomerization

89

3. In order to investigate the influence of specific ligands on oligomerization processes, the cells are incubated in the presence of agonists and antagonists for 15 min at 37°C before the measurement (see Note 13). 3.5.3. Control Experiments

1. The often-discussed problem concerning the experiments performed in heterologous expression systems is the issue concerning “overcrowding” of the protein of interest. Dimerization might simply be promoted at relatively high expression levels and may partially be an artifact of over-expression. Moreover FRET might be observed because of microdomain clustering. Therefore additional control experiments should be performed in order to prove that the estimated FRET efficiency reflects specific receptor–receptor interaction and does not result from random molecular interaction within the membrane. A good control is a measurement of FRET efficiency between membrane-targeted and noninteracting fusions of EYFP and ECFP, for example membrane receptor-EYFP and cytosolic protein-ECFP. 2. Measurement of the fluorescence lifetime of ECFP expressed in the cells alone or together with EYFP—when cells are loaded with both fluorescent proteins, not linked to any receptors also should be done as additional control experiment. During the experiment no FRET should be observed, despite high expression level of both fluorescent proteins. 3. In order to estimate the highest value of FRET efficiency, which is possible to be measured in the proposed model, the fusion construct of two fluorescent proteins used (ECFP_ EYFP) should be prepared and all kinds of the experiments described above should be done with cells expressing that construct (Fig. 7). 4. To avoid over-interpretation of FRET efficiency data, the measurements of homo-FRET (occurring between ECFP_ECFP) should be also conducted. 5. The interpretation of FRET efficiency alterations as a result of ligands presence should take into account the possible conformational changes upon ligand binding within the receptor molecule tagged with the fluorescent protein (Fig. 6). In order to avoid misinterpretation of the data, control experiments should be conducted. The donor-acceptor distance is calculated using the following equation: r = R0 ⎡⎣(E −1 − 1)1/6 ⎤⎦ The anticipated possible alteration in energy transfer induced by the conformational change is estimated according to the following equation:

90

S. Lukasiewicz et al.

Fig. 6. FRET efficiency measured in HEK 293 cells co-expressing dopamine D1-EYFP and D2-ECFP receptors in presence of ligand (clozapine) dependent on both clozapine concentration and on the time of ligand presence in the incubation medium.

E=

6r 5 ⎡1 + (r / R0 )6 ⎤ R06 ⎣ ⎦ 2

r

6. Moreover, it is very important to show that the used ligands do not change the fluorescent properties of ECFP labeling the receptor. Therefore, the lifetime measurements of the receptor-ECFP upon treatment with specific ligands should be done. In such a case, no changes in the fluorescence lifetime should be observed (Fig. 7). 3.6. Confocal Microscopy

Confocal microscopy is used to analyze the localization of the fluorescently tagged dopamine receptors in HEK 293 cells. It is especially important in the studies of mutant protein, often performed in order to identify the role of certain amino acid residues/ regions/domains within the receptor molecule in the oligomerization process. Such genetic manipulations within the receptor sequence might sometime change the cellular localization of the studied receptor. Cells grown on cover slips are transiently transfected with the cDNA encoding the fluorescently labeled dopamine receptors. 1. ECFP and EYFP fluorescence is excited by 457 nm and 514 nm wavelength lights, respectively. 2. For co-localization analysis Image ProPlus 4.5 software is used. Co-localization describes the existence of two or more fluorescently labeled molecule types in the same spatial positions. Pearson’s correlation coefficient is used to measure the overlap of the pixels and reflects the degree of relationship

6

Study of Dopamine Receptor Oligomerization

91

Fig. 7. Data obtained in our studies concerning the interactions between the GPCRs (D1R, D2R) and the appropriate alpha subunit of G protein confirm the specificity of the methodology. Using the same expression system and the same amount of DNA for transient transfections, FRET did not occur when two noninteracting fusion proteins (bearing ECFP and EYFP, respectively) were co-expressed in the same cell. Representative fluorescence emission spectra of HEK 293 cells cotransfected with either D1-EYFP or D2-EYFP and Gα-ECFP or GαI-ECFP fusion proteins. (a) Co-transfection of HEK 293 cells with D1-EYFP and GαS-ECFP (green line) or D1-EYFP and Gα1-ECFP (blue line); (b) Co-transfection of HEK 293 cells with D2-EYFP and GαI-ECFP (green line) or D2-EYFP and GαS-ECFP (blue line).

between two variables. It is one of the standard measures in pattern recognition: R=

∑ (Ri − Rav)·(Gi − Gav) ∑ (Ri − Rav) ·∑ (Gi − Gav) i

2

i

2

i

where Ri and Gi are the red and green intensities of voxel I, respectively, and Rav and Gav the average values of Ri and Gi, respectively. It is used for describing the correlation of the intensity distributions between red and green component of each dualchannel image. Pearson’s correlation coefficients should be calculated from randomly selected parts of the image (membrane signal) from individual cells co-transfected with different construct combinations. The average intensity of the fluorescence signal is measured for every image in a determined individual area of interest free of cells, and subtracted as a background. For analysis these regions are used, of which fluorescence intensities are correlated. For each combination of proteins, a minimum of 20 individual regions from different, independently transfected cells should be counted. 3. Interpretation of Pearson’s correlation coefficients, especially relative to each other is difficult, as their relative magnitudes are not proportional. By that reason coefficients of determination

92

S. Lukasiewicz et al.

(which are squared value of correlation coefficients) are estimated. The resulting coefficient of determination allows estimating the proportion of overlapping variance between two sets of pixels, thus making the interpreting correlation coefficients much easier.

4. Notes 1. Radioligand should be stored at −20°C and diluted in washing buffer (Tris–HCl, pH 7.4) just before use. 2. The plastic tubes do not promote robust results, probably owing to adhesion of membrane preparation or drugs to the test tube wall. 3. The fluorescence lifetime measurements are independent of any change in fluorophore concentration or excitation intensity; therefore that kind of measurement provides quantitative information about the interaction between labeled proteins of interest. In contrast, the steady state fluorescence spectroscopy measurements in cell suspension enable only the qualitative estimation of the FRET phenomenon. 4. Construction of fusion protein step-by-step: (a) PCR—mold pcDNA 3.1 plasmid with desired dopamine receptor cDNA primers: Forward—universal for pcDNA 3.1 Reverse—removed the STOP codon and introduced a unique restriction site (b) Agarose gel electrophoresis—identification and purification of PCR product (c) Enzymatic cleavage of obtained PCR product and plasmids pECFP and pEYFP (d) Agarose gel electrophoresis—identification and purification of desired DNA (e) Ligation—introduction of desired DNA fragment encoding appropriate dopamine receptor into vectors encoding fluorescence proteins (f) Transformation of E. coli DH5α cells (g) Bacterial cells cultured, plasmid DNA isolation and identification of colony containing vector encoding appropriate receptor fusion protein. 5. The amount of cDNA used for transfection does not always correspond with protein expression levels, therefore sometimes

6

Study of Dopamine Receptor Oligomerization

93

it is necessary to use a different donor/acceptor ratio in order to obtain the comparable levels of the donor and acceptor molecules. However, the total amount of cDNA used for transfection cannot exceed 15 μg for 3 × 106 cells. 6. To reduce the degradation of receptors, low temperature (4°C) must be used throughout these steps. All buffers should be prepared fresh on the day of analysis. 7. The final volume of tubes was 0.5 mL. Each sample should be prepared in triplicate. The concentrations of radioligands are measured using liquid scintillation counting (Beckman LS 650). Calculation of radioligands concentration are analyzed according to the equation: CL =

DPM 2, 200 × V P × AS

where: CL [nM]—concentration of radioligand DPM—disintegrations per minute Vp [mL]—final volume of tube As [Ci/mmol]—specific activity of radioligand. 8. Washing buffer should be prepared on the day of analyses and cooled to 4°C before use. Prior to counting, washed filters should be incubated with 5 mL of scintillation liquid Aquascynt (BioCare) for 12 h in room temp. 9. Deduction of the nonspecific disintegrations per minute (dpm) from the total at each concentration of radioligand gives a specific binding in dpm. The dpm are then converted to picomole per milligram of protein. 10. The radioligand concentration should be close to the dissociation constant, Kd, obtained from saturation binding analysis. 11. The dpm remaining for each displacement point can then be expressed as a percentage of total specific binding. 12. It is very important to control the temperature during the TCSPC experiments because fluorescence lifetime strongly depends on temperature. 13. Appropriate stock concentration of ligands required for the treatment of cells should be made right before the experiment.

Acknowledgments The authors would like to dedicate this work to the memory of the late professor Zygmunt Wasylewski, who encouraged us to employ fluorescence spectroscopy in our studies of dopamine receptors.

94

S. Lukasiewicz et al.

References 1. Hansen JL, Sheikh SP (2004) Functional consequences of 7TM receptor dimerization. Eur J Pharm Sci 23:301–317 2. Prinster SC, Hague C, Hall RA (2005) Heterodimerization of G protein-coupled receptors: specificity and functional significance. Pharmacol Rev 57:289–298 3. Aizman O, Brismar H, Uhlén P, Zettergren E, Levey AI, Forssberg H, Greengard P, Aperia A (2000) Anatomical and physiological evidence for D1 and D2 dopamine receptor colocalization in neostriatal neurons. Nat Neurosci 3:226–230 4. Hasbi A, Fan T, Alijaniaram M, Nguyen T, Perreault ML, O’Dowd BF, George SR (2009) Calcium signaling cascade links dopamine D1-D2 receptor heteromer to striatal BDNF production and neuronal growth. Proc Natl Acad Sci U S A 106:21377–21382 5. Dziedzicka-Wasylewska M, Faron-Górecka A, Andrecka J, Polit A, Kuśmider M, Wasylewski Z (2006) Fluorescence studies reveal heterodimerization of dopamine D1 and D2 receptors in the plasma membrane. Biochemistry 45: 8751–8759

6. Janetopoulos C, Devreotes P (2002) Monitoring receptor-mediated activation of heterotrimeric G-proteins by fluorescence resonance energy transfer. Methods 27:366–373 7. Elangovan M, Day RN, Periasamy A (2002) Nanosecond fluorescence resonance energy transfer-fluorescence lifetime imaging microscopy to localize the protein interactions in a single living cell. J Microsc 205:3–14 8. Lakowicz JR (1999) Principles of fluorescence spectroscopy. Kluwer Academic/Plenum Publishers, New York 9. Sambrook J, Fritsch EF, Maniatis T (1996) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, New York 10. Faron-Górecka A, Górecki A, Kuśmider M, Wasylewski Z, Dziedzicka-Wasylewska M (2008) The role of D1-D2 receptor hetero-dimerization in the mechanism of action of clozapine. Eur Neuropsychopharmacol 18:682–691 11. Stanasila L, Perez JB, Vogel H, Coteechia S (2003) Oligomerization of the alpha 1a- and alpha 1b-adrenergic receptor subtypes. Potential implications in receptor internalization. J Biol Chem 278:40239–40251

Chapter 7 Detection of Receptor Heteromers Involving Dopamine Receptors by the Sequential BRET-FRET Technology Gemma Navarro, Peter J. McCormick, Josefa Mallol, Carme Lluís, Rafael Franco, Antoni Cortés, Vicent Casadó, Enric I. Canela, and Sergi Ferré Abstract Until very recently, dopamine receptors, like other G-protein-coupled receptors, were believed to function as individual units on the cell surface. Now it has been described by several groups including ours that dopamine receptors not only function as homomers but also form heteromers with other receptors at the membrane level. Bioluminescence and fluorescence resonance energy transfer (BRET and FRET) based techniques have been very useful to determine the interaction between two receptors, but to demonstrate the existence of higher-order complexes involving more than two molecules requires more sophisticated techniques. Combining BRET and FRET in the Sequential BRET-FRET (SRET) technique permits heteromers formed by three different proteins to be identified. In SRET experiments, the oxidation of a Renilla Luciferase substrate triggers acceptor excitation by BRET and subsequent energy transfer to a FRET acceptor. Using this methodology here we describe the heteromerization between adenosine A2A, dopamine D2, and cannabinoids CB1 receptors in living cells. Key words: Dopamine receptors, Dopamine receptors interacting proteins, BRET, FRET, Sequential resonance energy transfer, GPCR, Receptor oligomerization, Heteromer, Protein–protein interaction

1. Introduction Dopamine exerts many of its physiological functions by interacting with dopamine receptors. Dopamine receptors are classified in D1like, with the D1 and D5 receptor subtypes (D1R and D5R), which usually couple to Gs/olf proteins, and D2-like, with the D2, D3, and D4 receptor subtypes (D2R, D3R, and D4R), which couple to Gi/o proteins (1). Like many other G-protein coupled receptors (GPCRs), dopamine receptors function as oligomers forming homomers (2–6) and heteromers with other dopamine receptors (7–10) or other GPCRs (11–13). A receptor heteromer has been Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_7, © Springer Science+Business Media, LLC 2013

95

96

G. Navarro et al.

recently defined as a macromolecular complex composed of at least two functional receptor units with biochemical or functional properties that are demonstrably different from those of its individual components (14). For this reason, receptor heteromers provide many implications for pharmacology, since they constitute new targets for drug development (15–17). The use of biophysical techniques, such as bioluminescence resonance energy transfer (BRET) and fluorescence resonance energy transfer (FRET) techniques has been fundamental in taking the issue of GPCR oligomerization to the front of GPCR research, providing evidence for an increasing number of receptor heteromers in living cells (18–20). Nevertheless, to detect higher order receptor oligomers a significant development in biophysical energy transfer techniques has been needed. To this end, two techniques have been developed in our laboratory to study oligomers formed by three different proteins and applied to determine heterotrimers involving dopamine receptors. One is BRET with bimolecular fluorescence complementation (BiFC), that we used to demonstrate heteromultimerization between adenosine A2A (A2AR), D2R, and cannabinoid CB1 (CB1R) receptors and between A2AR, D2R, and glutamate mGlu5 receptors (21, 22). The other technique is Sequential-BRET-FRET (SRET) (23). In SRET, the oxidation of an RLuc substrate by an RLuc-fusion protein triggers the excitation of the BRET acceptor (i.e., protein fused to GFP2) and subsequent energy transfer to the FRET acceptor (i.e., protein fused to YFP). SRET will only occur with these fusion proteins if the two acceptor–donor pairs, Rluc/GFP2 and GFP2/YFP, are at a distance of less than 10 nm. Here the technique is described to detect heterotrimers formed by A2AR, D2R, and CB1R in living cells. In general we conclude that SRET is an invaluable technique to identify oligomeric complexes of more than two proteins localized at the plasma membrane, including more than two GPCRs, which will enable us to better understand how signals are integrated at the plasma membrane level.

2. Materials 2.1. Fusion Proteins and Expression Vectors

1. The cDNA for functionally validated fusion proteins in suitable mammalian expression vectors are used. The human cDNAs for A2AR, D2R, CB1R, and the negative control human dopamine D4.4 receptor, cloned in pcDNA3.1. 2. pRluc-N1 vector (Rluc expressing vector, PerkinElmer, Wellesley, MA). 3. pGFP2-N3(h) vector (humanized PerkinElmer (Waltham, MA)).

pGFP2-N3(h)

from

7

SRET Technology to Detect Receptor Oligomers

97

4. pEYFP-N1 vector (enhanced yellow variant of GFP; Clontech, Heidelberg, Germany). 5. DH5α competent bacterial cells (Invitrogen, Carlsbad CA). 6. DH5α growing medium (LB): 10 g/L NaCl, 10 g/L Tryptone, and 5 g/L Yeast extract in mQ water. 7. Xtra Maxi kit (Nucleobond®, Düren, Germany). 2.2. Cell Culture

1. Human Embrionic Kidney (HEK) 293T cells are grown in 6-well cell culture plates (Techno Plastic Products, Lausanne, Switzerland). 2. As suitable growth medium, Complete Medium (Dulbecco’s modified Eagle’s medium (DMEM; Gibco (Carlsbad, CA)) supplemented with 2 mM L-glutamine, 100 U/ml penicillin– streptomycin, 5% (v/v) heat inactivated Fetal Bovine Serum (FBS), and 5% (v/v) nonessential amino acids (all supplements are from Invitrogen, Paisley, Scotland, UK) are used. 3. Protein quantification reagent; Bradford solution (Bio-Rad, Hercules, CA) diluted 1/5 (v/v) in milliQ (mQ water).

2.3. Transfection

1. Branched PEI (PolyEthylenImine, Sigma, Steinheim, Germany). Prepare a 40 μM solution in mQ water. 2. NaCl solution: NaCl 150 mM prepared in mQ water. 3. 0.05% trypsin (Gibco). 4. HBSS buffer: 0.185 g of CaCl2·12H2O, 0.370 g of KCl, 0.060 g of KH2PO4, 0.100 g of MgCl2·2H2O, 0.100 g of MgSO4·7H2O, 8.000 g of NaCl, 0.121 g of Na2HPO4·12H2O, and 2.385 g of HEPES in 1 L of mQ water. Use 1 M NaOH to adjust the pH to 7.4.

2.4. SRET

1. Assay buffer: HBSS buffer containing 1 g/L D-glucose. Add glucose to the buffer 10 min before using. 2. 500 μM DeepBlueC (Perkin Elmer) in anhydrous ethanol as luciferase substrate stock solution. Store at −20°C protected from light. 3. 500 μM coelenterazine h (Perkin Elmer) in anhydrous ethanol as luciferase substrate stock solution (Panreac, Barcelona, Spain). Store at −20°C protected from light.

2.5. Equipment

1. Multiskan Ascent Photometer (Thermo Labsystems, San Diego CA). 2. Fluostar Optima Fluorimeter equipped with a high-energy xenon flash lamp and appropriate filters (excitation filter at 485 nm and 410 nm and emission filter corresponding to 530 nm and 510 nm) (BMG Labtechnologies, Offenburg, Germany).

98

G. Navarro et al.

3. Mithras LB 940 equipped with detection filters for shortwavelength (400 nm) and long-wavelength (530 nm) (Berthold Technologies, DLReady, Germany). 4. 96-well white microplates for BRET (Porvair, Norfolk, UK). 5. 96-well black microplates with transparent bottom for fluorescence detection (Porvair).

3. Methods 3.1. Preparation of Fusion Proteins

Generate fusion constructs in Rluc, GFP2, or YFP expression vectors consisting of the cDNA for the protein of interest, inserted in-frame with the cDNA for the bioluminescent or fluorescent donor or acceptor molecule. 1. Select the donor and acceptor combination to perform SRET (i.e., A2AR-Rluc, D2R-GFP2, CB1R-YFP, and a negative control D4.4R-Rluc). Since in SRET experiments, the oxidation of an Rluc substrate triggers acceptor excitation by BRET and subsequent energy transfer to a FRET acceptor (Fig. 1) it is important to select the optimal combination (see Note 4).

Fig. 1. Sequential BRET-FRET (SRET). SRET combines BRET and FRET involving two energy donors and two acceptors. BRET and FRET techniques are combined to detect heterotrimers at the membrane level. Signal is initiated by oxidation of DeepBlueC by the Rluc-fused protein (A2AR-Rluc) that generates light emission at the indicated wavelength (blue ). The acceptor in BRET is a GFP2-fused protein (D2R-GFP2) that, after excitation, results in emission at the indicated wavelength (green ) that excites a YFP-fused protein (CB1R-YFP) by a FRET process with concomitant light emission peaking at the indicated wavelength (yellow ). Emission of YFP after addition of the Rluc substrate is only possible if the three fusion proteins are in close proximity (<10 nm) allowing bioluminescent and fluorescent sequential resonance energy transfer (SRET) to occur. A representation of excitation (top) and emission (bottom) spectra of fused proteins is shown in the right .

7

SRET Technology to Detect Receptor Oligomers

99

2. The human cDNAs for A2AR, D2R, CB1R, or D4.4R cloned in pcDNA3.1 are amplified, without their stop codons, using sense and antisense primers harboring unique EcoRI and BamHI sites to clone A2AR in the Rluc corresponding vector, EcoRI and KpnI to clone D2R in the GFP2 corresponding vector and BamHI and EcoRI to clone CB1R in EYFP corresponding vector or XhoI and BamHI sites to clone D4.4R in pRluc-N1 vector. To avoid transcription mutations, it is recommendable to use a high-fidelity DNA polymerase that offers extreme performance for all PCR applications (i.e., iProof™ from Bio-Rad). It is important to select the annealing temperature for each primer as indicated by the supplier and use 15–30 s/kbase for extension times, since longer times could induce a loss of bases. 3. The amplified fragments are subcloned to be in-frame into restriction sites of a multiple cloning region within pRluc-N1, pGFP2N3(h), or pEYFP-N1 vectors respectively yielding the plasmids corresponding to A2AR-Rluc, D4.4R-Rluc, D2R-GFP2, and CB1RYFP. To do this, the amplified fragments (dephosphorylated or not, depending on the activity of the enzymes) and vectors are cut with the specific enzymes (see step 3). Then 100 ng of vector and 60 ng/Kb of insert are mixed with 1 unit of T4 cDNA ligase (Promega, Fitchburg MA) for 3 h at room temperature. The ligation product is transformed in DH5α competent bacterial cells and the subcloned vectors are selected using the specific selection antibiotics of each vector. The positive colonies are grown in LB medium in the presence of the selection antibiotic concentration indicated by the vector supplier. 4. Sequence cDNA to test the correct sequence of the fusion protein. Obtain enough cDNA to do several transfections using a Xtra-Maxi kit (usually we obtain 500 μL of cDNA from 1–5 μg/μL concentration). 5. Check that the luminescence or fluorescence is detectable after fusion protein expression (represent the amount of cDNA transfected in HEK 293T cells versus bioluminescence or fluorescence detected). If possible, use confocal microscopy to visualize (by its own fluorescence or using antibodies) correct cellular localization of fusion proteins. 6. Validate the fusion proteins of interest, including suitable control proteins, by comparing fusion and wild-type proteins in functional assays. ERK 1/2 phosphorylation and cAMP production can be used for this purpose as described previously (13, 24). 7. Generate positive controls for SRET experiments. Rlucexpressing vector (pRlu-N1) is amplified without its stop codon using sense and antisense primers harboring unique HindIII and KpnI sites to clone Rluc in-frame into restrictions sites of a multiple cloning site of GFP2-YFP vector (pcDNA3.1GFP2-YFP; Biosignal Packard) to obtain the fusion protein Rluc-GFP2-YFP.

100

G. Navarro et al.

8. Reconstitute and store the luciferase substrate stock solution containing DeepBlueC with anhydrous ethanol. Protect the solution from light. 3.2. Cell Transient Transfection

1. HEK 293 T cells are passaged when approaching confluence with trypsin/EDTA to provide new maintenance cultures in 150 cm2 flasks. One 150 cm2 flask is required for transfection in order to obtain enough transfected cells to perform a SRET saturation curve (see Subheading 3.4, step 4). Aliquot cells in 6-well cell culture plate in growth medium. They should be 60–80% confluent after 24 h. Maintain at 37°C, 5% CO2 and 90% of humidity. 2. Transfect the expression vectors corresponding to the desired fusion proteins at the suitable ratios (see legends of Figs. 2 and 3) using the PolyEthylenImine (PEI) method. Other methods of transfection may be used as well. To do this, two Falcon tubes

Fig. 2. SRET for A2AR, D2R, and CB1R in living cells. SRET assays are performed 48 h posttransfection in cells expressing A2AR-Rluc (2 μg of cDNA; approximately 100,000 luminescence units), D2R-GFP2 (3 μg of cDNA; approximately 6,000 fluorescence units), and CB1R-YFP (9 μg of cDNA; approximately 18,000 fluorescence units) or the equivalent amounts of the fluorescence or luminescence proteins or transfected with the positive SRET construct (1 μg of cDNA of Rluc-GFP2-YFP construct). Net SRET was obtained by monitoring the YFP fluorescence emission after DeepBlueC addition, with subtraction of the value obtained with cells expressing the same amount of A2AR-Rluc and the corresponding BRET acceptor. Significant net SRET was detected for A2AR-Rluc/D2R-GFP2/ CB1R-YFP coupling or for the positive SRET control, while negligible net SRET was obtained in cells expressing equivalent amounts of A2AR-Rluc, GFP2, and CB1R-YFP, or A2AR-Rluc, D2R-GFP2, and YFP. Data are expressed as the mean net SRET ± S.E.M. of four independent experiments performed in duplicate. One-way ANOVA followed by Newman–Keuls test showed significant differences with respect to negative controls (***: P < 0.001).

7

SRET Technology to Detect Receptor Oligomers

101

Fig. 3. SRET saturation curve for A2AR-D2R-CB1R heteromers in living cells. SRET saturation curves were obtained using HEK-293T cells transfected with 2 μg of the cDNA for A2AR-Rluc (approximately 100,000 luminescence units) and 3 μg of the cDNA for D2RGFP2 (approximately 6,000 fluorescence units) and increasing amounts of the cDNA for CB1R-YFP (8,000 to 18,000 fluorescence units). Values, expressed as net SRET, represent the mean ± S.E.M. of two independent experiments performed in triplicate. Negative control is constituted by cells expressing the equivalent amounts of D4R-Rluc/A2AR-GFP2/ CB1R-YFP giving linear (nonspecific) SRET with similar amounts of fluorescence and luminescence as those giving saturable SRET.

are needed. In the Falcon A each μg of cDNA is mixed with 25 μL of NaCl solution. In Falcon B the same amount of NaCl solution is mixed with 1.25 μL/μg cDNA of 40 μM branched PEI solution. Both Falcon tubes are vortexed a few seconds. After this, Falcon B solution is added to Falcon A. The final solution is strongly vortexed for 10 s. Ten minutes later, 1 mL of serum starved medium is added. Cells are incubated for 4 h with the cDNA-PEI solution. Then cells are placed in a fresh complete culture medium. 3. Around 48 h after transient transfection, detach the cells and resuspend them in HBSS buffer containing 1 g/L D-glucose. Wash cells twice with the same buffer for 5 min and resuspend them in the same buffer. 4. Using 10 μL of cell suspension, quantify the amount of protein using a Bradford assay and dilute cells to 200 μg/mL. It is important to maintain the same amount of protein in each sample. Aliquot cells into 96-well white and black isoplates (100 μL per black well and 90 μL per white well). 3.3. SRET Detection

Using aliquots of transfected cells (20 μg of protein), four different determinations are performed in parallel: 1. Quantification of protein-YFP expression by determination of the fluorescence due to protein-YFP. Cells distributed into 96-well microplates (black plates with a transparent bottom), are read in a Fluostar Optima Fluorimeter using an excitation

102

G. Navarro et al.

filter at 485 nm, and a 10 nm bandwidth emission filter corresponding to 530 nm (527–536 nm). 2. It is also important to quantify the protein-GFP2 expression to control for BRET energy transmission and to develop the linear un-mixing arrangement. Cells distributed into 96-well microplates (black plates with a transparent bottom), are read in a Fluostar Optima Fluorimeter using an excitation filter at 410 nm and a 10 nm bandwidth emission filter corresponding to 510 nm (506–515 nm). To determine the exact fluorescence amount of proteinGFP2 and protein-YFP it is necessary to calculate the linear un-mixing. First, analyze the contribution of GFP2 and YFP proteins alone using the two detection channels (see Note 6). Fluorescent determinations are measured in parallel in experiments with cells expressing only one of these proteins and normalized to the sum of the signal obtained in the two detection channels. Values obtained in 1 and 2 are corrected considering this linear un-mixing. The sample fluorescence is the emission at 530 nm corrected as described minus the fluorescence of cells expressing only protein-Rluc and protein-GFP2. The protein-GFP2 fluorescence is the emission at 510 nm corrected as described minus the fluorescence of cells expressing only protein-Rluc. 3. Quantification of protein-Rluc expression by determination of the luminescence due to protein-Rluc. Cells are distributed in 96-well microplates (white plates) and luminescence is determined 10 min after the addition of 5 μM coelenterazine H in a Mithras LB 940 multimode reader. 4. SRET measurements. Cells are distributed in 96-well microplates (white plates) and 5 μM DeepBlueC is added. SRET signal is collected using a Mithras LB 940 reader 30 s after the substrate addition with detection filters for shortwavelength (400 nm (370–450 nm)) and long-wavelength (530 nm (510–590 nm)). Net SRET is defined as [(longwavelength emission)/(short-wavelength emission)] − Cf, where Cf corresponds to [(long-wavelength emission)/ (short-wavelength emission)] for cells expressing proteinRluc, protein-GFP2 and the other protein partner not fused to a fluorescence protein (similar values are obtained measuring Cf in cells expressing protein-Rluc only and proteinGFP2). Linear un-mixing is done for quantification, taking into account the spectral signature to separate the two fluorescence emission spectra (see Note 6). SRET determined in cells expressing A2AR-Rluc, D2R-GFP2 and CB1RYFP or the corresponding positive or negative controls is shown in Fig. 2.

7

3.4. SRET Saturation Curves

SRET Technology to Detect Receptor Oligomers

103

SRET saturation curve is further proof for the specificity of the interaction observed. 1. Transiently transfect HEK 293T cells with a constant amount of the constructs corresponding to the protein-Rluc and proteinGFP2 and with increasing amounts of the construct corresponding to protein-YFP indicated in the legend of Fig. 3. 2. After 48 h of transient transfection determine SRET as indicated (see Subheading 3.3) for each transfection condition. 3. Both fluorescence and luminescence for each sample are measured to confirm similar donors expression (approximately 100,000 bioluminescence units and 6,000 GFP2 fluorescence units) while monitoring the increase in acceptor expression (1,000 to 20,000 fluorescence units) (see Note 7). 4. Represent the net SRET values as a function of the amount of the acceptor. In each saturation curve, the relative amount of acceptor is given as the ratio between the fluorescence of the acceptor (YFP) and the luminescence of the first donor (Rluc). Curves are fitted to a nonlinear regression equation, assuming a single phase (i.e., with Graph-Pad Prism software, San Diego, CA, USA). From these saturation curves, an apparent SRETmax and an apparent SRET50 can be determined (see Note 8). A SRET saturation curve obtained by increasing CB1R-YFP expression while maintaining the same A2AR-Rluc/D2-GFP2 ratio (Fig. 3) allows determination of the following parameters for the trimer A2AR-Rluc/D2R-GFP2/CB1R-YFP: apparent SRETmax of 0.18 ± 0.05 and apparent SRET50 of 0.013 ± 0.007.

4. Notes 1. If cells are dying after transfection refer to cytotoxicity of transfection reagent. 2. The amount of fusion proteins transfected must be near the physiological range. 3. If the donor protein expression is high, then the fluorescence background will be high and poor SRET signal will be detected. If there is low donor protein expression, the acceptor protein will not be excited. 4. The relative orientation of donor and acceptor can be unsuitable for SRET. 5. Add DeepBlueC immediately before SRET detection. 6. Given the spectral emission of the GFP2/YFP pair, the contribution of GFP2 or YFP proteins to the two detection channels

104

G. Navarro et al.

is significant. Thus, the spectral signature (25) of these fusion proteins must be determined and considered for SRET and YFP expression evaluation. 7. In SRET saturation curves, be sure that the amount of luminescence due to protein-Rluc and the amount of fluorescence due to protein-GFP2 are constant while increasing the fluorescence of the acceptor protein-YFP. 8. It should be noted that the SRETmax and SRET50 will differ when different ratios of donor–acceptor for BRET are used. This is reminiscent of what occurs in enzymology, when the Vmax and KM for an enzyme using two substrates are calculated by maintaining the concentration of one substrate constant but varying the concentration of the other. For such enzymes, the calculated values are known as “apparent Vmax” and “apparent KM”. Accordingly, we propose the denomination of “apparent SRETmax” and “apparent SRET50”.

Acknowledgments Study supported by grants from Spanish Ministerio de Ciencia y Tecnología (SAF2008-00146, SAF2008-03229-E, and SAF200907276), grant 060110 from Fundació La Marató de TV3 and by the Intramural Funds of the National Institute on Drug Abuse. References 1. Missale C, Nash SR, Robinson SW, Jaber M, Caron MG (1998) Dopamine receptors: from structure to function. Physiol Rev 78:189–225 2. Ng GY, Mouillac B, George SR, Caron M, Dennis M, Bouvier M, O’Dowd BF (1994) Desensitization, phosphorylation and palmitoylation of the human dopamine D1 receptor. Eur J Pharmacol 267:7–19 3. Kong MM, Fan T, Varghese G, O’Dowd BF, George SR (2006) Agonist-induced cell surface trafficking of an intracellularly sequestered D1 dopamine receptor homo-oligomer. Mol Pharmacol 70:78–89 4. George SR, Lee SP, Varghese G, Zeman PR, Seeman P, Ng GY, O’Dowd BF (1998) A transmembrane domain-derived peptide inhibits D1 dopamine receptor function without affecting receptor oligomerization. J Biol Chem 273:30244–30248 5. Guo W, Urizar E, Kralikova M, Mobarec JC, Shi L, Filizola M, Javitch JA (2008) Dopamine D2 receptors form higher order oligomers at physiological expression levels. EMBO J 27:2293–2304

6. Han Y, Moreira IS, Urizar E, Weinstein H, Javitch JA (2009) Allosteric communication between protomers of dopamine class A GPCR dimers modulates activation. Nat Chem Biol 9:688–695 7. Rashid AJ, So CH, Kong MM, Furtak T, El-Ghundi M, Cheng R, O’Dowd BF, George SR (2007) D1-D2 dopamine receptor heterooligomers with unique pharmacology are coupled to rapid activation of Gq/11 in the striatum. Proc Natl Acad Sci U S A 9:654–659 8. Marcellino D, Ferré S, Casadó V, Cortés A, Le Foll B, Mazzola C, Drago F, Saur O, Stark H, Soriano A, Barnes C, Goldberg SR, Lluis C, Fuxe K, Franco R (2008) Identification of dopamine D1-D3 receptor heteromers. Indications for a role of synergistic D1-D3 receptor interactions in the striatum. J Biol Chem 283:26016–26025 9. Fiorentini C, Busi C, Gorruso E, Gotti C, Spano P, Missale C (2008) Reciprocal regulation of dopamine D1 and D3 receptor function and trafficking by heterodimerization. Mol Pharmacol 74:59–69

7

SRET Technology to Detect Receptor Oligomers

10. So CH, Verma V, Alijaniaram M, Cheng R, Rashid AJ, O’Dowd BF, George SR (2009) Calcium signaling by dopamine D5 receptor and D5-D2 receptor hetero-oligomers occurs by a mechanism distinct from that for dopamine D1-D2 receptor hetero-oligomers. Mol Pharmacol 75:843–854 11. Ferrada C, Moreno E, Casadó V, Bongers G, Cortés A, Mallol J, Canela EI, Leurs R, Ferré S, Lluís C, Franco R (2009) Marked changes in signal transduction upon heteromerization of dopamine D1 and histamine H3 receptors. Br J Pharmacol 157:64–75 12. Juhasz JR, Hasbi A, Rashid AJ, So CH, George SR, O’Dowd BF (2008) Mu-opioid receptor heterooligomer formation with the dopamine D1 receptor as directly visualized in living cells. Eur J Pharmacol 581:235–243 13. Canals M, Marcellino D, Fanelli F, Ciruela F, de Benedetti P, Goldberg SR, Neve K, Fuxe K, Agnati LF, Woods AS, Ferré S, Lluis C, Bouvier M, Franco R (2003) Adenosine A2A-dopamine D2 receptor-receptor heteromerization. Qualitative and quantitative assessment by fluorescence and bioluminescence resonance energy transfer. J Biol Chem 278:46741–46749 14. Ferré S, Baler R, Bouvier M, Caron MG, Devi LA, Durroux T, Fuxe K, George SR, Javitch JA, Lohse MJ, Mackie K, Milligan G, Pfleger KD, Pin JP, Volkow ND, Waldhoer M, Woods AS, Franco R (2009) Building a new conceptual framework for receptor heteromers. Nat Chem Biol 5:131–134 15. Casadó V, Cortés A, Mallol J, Pérez-Capote K, Ferré S, Lluis C, Franco R, Canela EI (2009) GPCR homomers and heteromers: a better choice as targets for drug development than GPCR monomers? Pharmacol Ther 124: 248–257 16. Ferré S, Lluís C, Lanciego JL, Cortés A, Mallol J, Canela EI, Lluís C, Franco R (2010) G protein-coupled receptor heteromers as new targets for drug development. CNS Neurol Disord Drug Targets 9:596–600 17. Ferré S, Navarro G, Casadó V, Cortés A, Mallol J, Canela EI, Lluís C, Franco R (2010) G protein-coupled receptor heteromers as new targets for drug development. Prog Mol Biol Transl Sci 91:41–52

105

18. Milligan G (2004) Applications of bioluminescence- and fluorescence resonance energy transfer to drug discovery at G proteincoupled receptors. Eur J Pharm Sci 21: 397–405 19. Pfleger KD, Eidne KA (2005) Monitoring the formation of the dynamic G-protein-coupled receptor-protein complexes in living cells. Biochem J 385:625–637 20. Marullo S, Bouvier M (2007) Resonance energy transfer approaches in molecular pharmacology and beyond. Trends Pharmacol Sci 28:362–365 21. Cabello N, Gandía J, Bertarelli DC, Watanabe M, Lluís C, Franco R, Ferré S, Luján R, Ciruela F (2009) Metabotropic glutamate type 5, dopamine D2 and adenosine A2A receptors form higher-order oligomers in living cells. J Neurochem 109:1497–1507 22. Navarro G, Carriba P, Gandía J, Ciruela F, Casadó V, Cortés A, Mallol J, Canela EI, Lluis C, Franco R (2008) Detection of heteromers formed by cannabinoid CB1, dopamine D2, and adenosine A2A G-protein-coupled receptors by combining bimolecular fluorescence complementation and bioluminescence energy transfer. ScientificWorldJournal 8:1088–1097 23. Carriba P, Navarro G, Ciruela F, Ferré S, Casadó V, Agnati L, Cortés A, Mallol J, Fuxe K, Canela EI, Lluís C, Franco R (2008) Detection of heteromerization of more than two proteins by sequential BRET-FRET. Nat Methods 5: 727–733 24. Carriba P, Ortiz O, Patkar K, Justinova Z, Stroik J, Themann A, Müller C, Woods AS, Hope BT, Ciruela F, Casadó V, Canela EI, Lluis C, Goldberg SR, Moratalla R, Franco R, Ferré S (2007) Striatal adenosine A2A and cannabinoid CB1 receptors form functional heteromeric complexes that mediate the motor effects of cannabinoids. Neuropsychopharmacology 32:2249–2259 25. Zimmermann T, Rietdorf J, Girod A, Georget V, Pepperkok R (2002) Spectral imaging and linear un-mixing enables improved FRET efficiency with a novel GFP2-YFP FRET pair. FEBS Lett 531:245–249

Chapter 8 BRET Approaches to Characterize Dopamine and TAAR1 Receptor Pharmacology and Signaling Stefano Espinoza, Bernard Masri, Ali Salahpour, and Raul R. Gainetdinov Abstract It is evident that G protein-coupled receptors (GPCRs) such as D2 dopamine receptor and functionally related Trace Amine Associated Receptor 1 (TAAR1) can engage both in G protein-dependent (e.g., cAMP-mediated) and -independent β-arrestin-mediated signaling modalities. Both of these signaling events can be monitored in real-time and in live cells by using new biosensors based on a Bioluminescence Resonance Energy Transfer (BRET) approach. Here we discuss the practical applications of BRET to analyze dynamics of cAMP modulation via an EPAC biosensor as well as recruitment of β-arrestin2 to the D2 dopamine receptor. Combination of these approaches allows for a comparison of activity of pharmacological compounds on these signaling modalities as demonstrated for various antipsychotics as regard to D2 dopamine receptor. Furthermore, analysis of cAMP concentrations in cells expressing TAAR1 provides a simple high-throughput screening method to identify new ligands for this receptor. These BRET approaches could be applied for the characterization of pharmacology and signaling of variety of other GPCRs. Key words: D2R, β-Phenylethylamine

TAAR1,

cAMP,

β-arrestin2,

BRET,

EPAC,

3-Methoxytyramine,

1. Introduction Dopamine (DA) is a major monoaminergic neurotransmitter in the mammalian brain and controls many physiological functions, such as voluntary movement, emotion, reward, cognition and neuroendocrine secretion (1). DA is also involved in critical processes in the periphery, including gastrointestinal activity, regulation of kidney function and catecholamine secretion. A variety of human pathologies have been linked to a dysregulation of dopaminergic neurotransmission ranging from schizophrenia and addiction to Parkinson’s disease, Tourette’s syndrome, Attention Deficit Hyperactivity Disorder (ADHD), and hyperprolactinemia (1). DA exerts its physiological functions via action on five members of a Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_8, © Springer Science+Business Media, LLC 2013

107

108

S. Espinoza et al.

family of G protein-coupled receptors (GPCR) that are subdivided into D1-like (D1 and D5) and D2-like (D2, D3, D4) dopamine receptor subtypes. D1-like dopamine receptors are positively coupled to adenylate cyclase and their stimulation increase cAMP level inside the cells, while D2-like dopamine receptors negatively regulate the production of cAMP in the cell. In addition, recent studies have demonstrated a novel G protein-independent pathway for D2 dopamine receptors (D2Rs) that involves the multifunctional scaffolding protein β-arrestin2 (2). While best known for their role in GPCR desensitization, β-arrestins are emerging as key signaling proteins in a variety of physiological processes (3). Regarding D2R, β-arrestin2-orchestrated formation of a protein complex including D2R, and other proteins (PP2A and Akt) has been demonstrated to be important for the full manifestation of certain D2R-related behaviors (4). Intriguingly, a recent comparison of the ability of several clinically active antipsychotics to affect G protein-dependent (cAMP accumulation) versus G protein-independent processes (β-arrestin2 recruitment), has shown that antipsychotics are generally more effective in preventing β-arrestin2 recruitment to the D2R than regulating cAMP concentration within the cell (5). Another GPCR that is emerging as a functionally important modulator of dopaminergic activity in the brain is Trace Amine Associated Receptor 1 (TAAR1) (6–8). TAAR1 belongs to the family of Trace Amine Associated Receptors (TAARs) that is expressed in several mammalian brain areas including limbic regions and sites containing monoaminergic cell bodies such as the VTA (8). It has been shown that TAAR1 can modulate dopaminergic activity both by altering VTA neurons firing and D2 receptor signaling (9). TAAR1 is a Gs coupled receptor and it is activated by a class of compounds called trace amines (TAs) that include β-phenylethylamine (β-PEA), tyramine, octopamine, and tryptamine (6, 7, 10). Other interesting molecules that bind TAAR1 are amphetamines and the major extracellular dopamine metabolite 3-methoxytyramine (3-MT) (6, 10). This compound has been traditionally considered as an inactive o-methylated (by COMT) metabolite of extracellular DA but recently its role in neuromodulation and DA-independent movement control that is in part mediated by TAAR1 has been described (11). Classical assays that evaluate cAMP concentration consist in detection of the accumulation of the second messenger inside the cell, or in the evaluation of its signaling as measured by a reporter gene. The main drawback of these techniques is their low versatility in terms of temporal monitoring of the sample. In the last few years, new biosensors based on Fluorescence Resonance Energy Transfer (FRET) and Bioluminescence Resonance Energy Transfer (BRET) have been described to measure in real-time physiological events within living cells (12–14). To monitor cAMP variations, a genetically encoded BRET biosensor has been developed (10). The exchange protein activated by cAMP (EPAC) biosensor binds

8

Dopamine/TAAR1 Signaling

109

cAMP in the cell resulting in a conformational change and in an energy transfer between Rluc and YFP in a concentration dependent manner. With a similar BRET approach, it is also possible to monitor, in real time, β-arrestin2 recruitment to the D2 receptor (5). These two systems are increasingly useful in the study of two important processes in vitro that involve GPCRs signaling: cAMP levels modulation and β-arrestin recruitment to the receptor. The major advantages of these approaches are (1) the possibility to monitor the kinetics of signaling events (second by second) within live cells, (2) a direct comparison of activity for a given pharmacological compound on G protein-dependent and independent signaling events within the same cell.

2. Materials 2.1. Cell Culture and Compounds

1. Dulbecco’s Modified Eagle’s Medium (DMEM) (Invitrogen, Carlsbad, CA) supplemented with 10% (vol/vol) of fetal bovine serum (FBS, Sigma, St. Louis, MO), 10 mM of HEPES (store at 4°C) (Invitrogen), and 0.01 mg/mL of gentamicin (store at room temperature (RT)) (Invitrogen) is used for normal cell culture purpose and it is stored at 4°C. FBS is aliquoted in 50 mL tube and stored at −20°C. 2. Phenol red free Minimum Essential Medium (MEM) containing 2% of FBS, 10 mM HEPES, and 2 mM L-glutamine (Invitrogen) is used for plating the cells before the experiments and it is stored at 4°C. L-glutamine is a 100× solution stocked at −20°C in 5 mL aliquots. 3. Poly-D-lysine (Sigma) is a powder dissolved in tissue-culture water at 0.1 mg/mL and stored at 4°C. Do not discard after the use since it could be reused several times. 4. Phosphate Buffer Saline (PBS) (Invitrogen) is used to wash the cells. PBS containing calcium and magnesium (Sigma) and 0.003% (wt/vol) of ascorbic acid (Sigma) is the medium for the BRET experiments and used to dissolve the compounds (store at 4°C). 5. Trypsin at 0.25% (wt/vol) with ethylenediamine tetraacetic acid (EDTA) (1 mM) (Invitrogen) is used to detach the cells and is kept at 4°C. 6. Burker chamber (Blaubrand, Germany) to count cells and trypan blue solution at 0.4% (Sigma) to stain live ones. 7. Human embryonic kidney 293 cells (HEK293T) (Invitrogen). 8. Coelenterazine h (Promega, Madison, WI) is dissolved in anhydrous ethanol to a final concentration of 1 mM and it must be

110

S. Espinoza et al.

stored at −20°C with some desiccant compound such as silica gel to keep the humidity at low levels. Coelenterazine h is light sensitive and the working solutions must be prepared immediately before the addition to the plate. 9. 3-Isobutyl-1-methylxanthine (IBMX), β-phenylethylamine (β-PEA), 3-methoxy-tyramine (3-MT), dopamine (DA), forskolin, haloperidol, quinpirole, and isoproterenol are from Sigma. IBMX is prepared at 200 mM in DMSO (1,000×) and stored at −20°C. Forskolin is dissolved at 10 mM in DMSO as stock solution and it is stable at RT at least for 6 months. All the other compounds are made ready at 10 mM in PBS (plus ascorbic acid—see Subheading 4) and then diluted to the desired concentration 1:10 each time in PBS except haloperidol that has to be dissolved initially in 10 mM in DMSO. 10. Cells are plated in a culture-treated 96-well plate (Corning, Corning, NY), white and clear-bottom (catalogue number 3610). Before the experiments a white backing tape (Perkin Elmer-catalogue number 6005199) has to be placed to cover the bottom of the plate. 2.2. Plasmids and Transfection

1. Plasmids containing the cDNA for the human Trace Amine Associated Receptor 1 (hTAAR1) were obtained from the cDNA Resource Center at the University of Missouri-Rolla and the American Type Culture Collection (Manassas, VA). A variant of the wild-type form is used to improve the expression at the plasma membrane (10). 2. HA-tagged D2 Long dopamine receptor is fused to Renilla Luciferase at the C-terminus (5). 3. Mouse β-arrestin2 is fused to YFP at the C-terminus (5). 4. The BRET biosensor EPAC is a modification from an existing FRET biosensor (10). 5. pRluc-N or pRluc-C vectors are from Perkin Elmer (Downers Grove, IL). 6. pEYFP-N or pEYFP-C vectors are from BD Biosciences (San Jose, CA). 7. pcDNA 3 is from Invitrogen. 8. Regarding calcium-phosphate transfection, calcium chloride (Sigma) is dissolved at 2.5 M in Milli-Q water. HEPES buffered saline (HBS) 2× is composed of sodium chloride (Sigma) (2.5 M), HEPES (0.5 M), and sodium phosphate dibasic (1 M) (Sigma). The pH of the solution is adjusted to 7.1. Both solutions should be stored at 4°C and kept sterile by opening only under the cell culture hood.

8

2.3. Experimental Machine and Software

Dopamine/TAAR1 Signaling

111

1. To read the plate use the Mithras LB940 (Berthold Technologies, Germany) or the Infinite F500 (Tecan, Switzerland) with a MicroWin 2000 or an i-control software respectively. 2. Data are analyzed using GraphPad Prism 5.

3. Methods EPAC cAMP biosensor, as well as β-arrestin2 recruitment to D2R, relies on the same principle of function that is BRET. BRET is a form of non-radiative energy transfer between a donor (Renilla Luciferase, Rluc) and an acceptor (YFP) (15). Different couples of donor and acceptor exists, please see ref. (16) for review. Upon oxidation of its substrate coelenterazine h, Rluc emits light with a peak at 475 nm. If the donor is in the proximity of the acceptor (around 0.1 nm), the energy emitted from Rluc can excite the YFP that in turn emits light with a peak at 530 nm. The change in the ratio between emissions by Rluc and the YFP reflects a change in the distance between the two molecules. In the EPAC biosensor, at basal or low cAMP levels, Rluc and YFP are in close proximity resulting in a constitutive elevated BRET signal. When cAMP binds EPAC, a conformational change occurs increasing the distance between the donor and the acceptor resulting in a decreased change of BRET ratio. In β-arrestin2-D2R experiments, when DA or another agonist stimulates the receptor (D2-Rluc), β-arrestin2YFP is recruited from the cytosol to the plasma membrane (to bind D2R) and this translocation results in an increase of BRET ratio. In this way, it is possible to study in real time these physiological events by integrating the light emission by the two tags with appropriate filters (17). Regarding TAAR1, EPAC biosensor has been shown to be a useful tool in studying trace amines pharmacology and in finding new agonists of the receptors (10, 18). It has demonstrated a good sensitivity and very similar results compared to other techniques (β-PEA and 3-MT potencies to TAAR1). Similarly, β-arrestin2 recruitment to the D2R as measured by BRET assay has been validated as an effective method to study this signaling pathway in antipsychotics pharmacology (5). 3.1. Cells Seeding and Transfection

1. The wild type HEK-293T cells are cultured in 150-mm dishes with DMEM for the maintenance of a certain number of cells according to the experiments planned. The medium should be warmed in a 37°C bath for at least 20 min before using it. Cells are passaged and seeded in 100-mm dishes the day before the transfection using trypsin-EDTA. All the media and solutions must be opened only under the hood (see Note 1).

112

S. Espinoza et al.

2. Wash with 10 mL of PBS the 150-mm dishes, add 2 mL of trypsin-EDTA to each dish and let the trypsin to act for 1–5 min at RT. 3. Add about 5 mL of DMEM in order to inactivate trypsin and to help cell harvesting. Gently pipette up and down to detach all the cells from the dish and to break cells lumps. 4. Pour the cells in a 15 mL Falcon tube and count them. Mix 20 μL of cells and 20 μL of trypan blue solution on a piece of parafilm and then pipette 20 μL in the bunker chamber and count the cells under the microscope. Make sure to make a homogeneous cell suspension in the medium with the pipette before collecting the 20 μL in order to have a representative concentration of the solution. 5. After counting the number of cells/mL seed about 3 million of cells for each 100-mm dish in 10 mL of DMEM (see Note 2). Each dish will be used for a single transfection, including control mock transfection with an empty vector (see Note 3). 6. The morning after the seeding, cells will be transfected. The solutions required are the CaCl2 (2.5 M), the HBS 2× and water. All the solutions must be sterilized by filtration with a 0.22 μm filter and then opened only under the hood. All the working aliquots of these solutions (10–50 mL) should be kept in a refrigerator and for more reproducible results equilibrated to room temperature (RT) for 30 min before using them. The remaining aliquots of HBS 2× should be stored at −20°C (see Note 4). 7. Take the DNA tubes out of the freezer (see Note 5) and let them thaw at RT. 8. Place 15 mL Falcon tubes under the hood in a rack, one for each transfection. 9. Put the DNA in each tube. It is important to have the same amount of DNA for each transfection, so equilibrate with empty vector (pcDNA 3). Usually, the amounts for a 1 mL of solution, that is enough for a 100-mm dish, are as follows: EPAC (3 μg), TAAR1 (5 μg), D2R (2 μg), and β-arrestin2 (2 μg). Regarding D1R, they have sufficient endogenous expression in HEK-293T cells to elicit a response detectable by BRET upon their stimulation. 10. Add 50 μL of CaCl2 2.5 M and enough water to have a total volume of 0.5 mL containing the DNA as well. 11. Pipette up and down for few times to mix the solution. 12. Add 0.5 mL of the HBS 2× drop by drop to each tube and mix by bubbling air inside the solution with the P1000 pipette for 10–15 times.

8

Dopamine/TAAR1 Signaling

113

13. Add the transfection solution drop wise to each dish, gently and trying to cover the entire plate. Rock the dishes back and forth for a few times before replacing them in the incubator. After few hours, a small dark precipitate should be observed in the cells under the microscope. 3.2. Plating

1. Cells are normally plated in the 96-well plate after 16–24 h, but for TAAR1 it is better to plate the same day, in the afternoon, so as to do the experiment 1 day after the transfection since TAAR1 is partially degraded in vitro (10). 2. After 4–6 h post transfection for TAAR1 and 16–24 h for dopamine receptors and β-arrestin2 (see Note 6) cells have to be plated. 3. The assay plate is a culture-treated, white, clear-bottom 96-well plate. The plate is sterile so it has to be properly handled under the hood. At this step the phenol red free Minimum Essential Medium (MEM) containing 2% of FBS, 10 mM HEPES, and 2 mM L-glutamine should be used to avoid color in the medium that could disturb the light emission during the experiment (“clear medium”). 4. Place the poly-D-lysine and the PBS under the hood. 5. While pre-warming the medium, pipette approximately 100 μL of poly-D-lysine in each well of the 96-well plate and incubate for 10–20 min (see Note 7). 6. After this period re-collect the poly-D-lysine with the pipette and wash once all the wells with PBS (about 150 μL for each well). Then remove the remaining PBS with a vacuum aspirator. 7. Wash twice the transfected cells in the 100-mm dishes with PBS and detach them by pouring 1 mL of trypsin-EDTA for each dish. 8. After 5 min of incubation (see Note 8), add 4 mL of clear medium in each dish. Gently pipette the cells up and down to detach them. Pour them in a labeled 15-mL Falcon tube, one for each transfection. 9. Spin the tubes in a centrifuge for 5 min at 200 × g to pellet the cells. 10. Remove the medium from the tube and resuspend the cells with 1 mL of clear medium and count the cells for each tube as described above. 11. Then dilute the cells solutions of each tube with the clear medium so as to have a concentration of 750,000 cells for each mL. 12. Put 100 μL of cells suspension for each well in the assay plate for all the different transfections, according to the experiment

114

S. Espinoza et al.

design. This step can be done either using a normal p100 pipette or a repeater pipette (see Note 9). 13. The 96-well plate is stored in the incubator for at least 12 h before the experiment. 3.3. BRET Experiment

24–48 h post-transfection the EPAC sensor and the other proteins should be sufficiently expressed. Since Rluc produces a signal independently from BRET, an internal control of the level of transfection is represented by the basal Rluc counts. For Mithras, 100,000 counts are the minimum for a good signal to noise ratio. 1. Pre-warm to RT or to 37°C (depending on the planned experiment) the PBS with calcium and magnesium and add 0.003%(wt/vol) of ascorbic acid. 30–50 mL of PBS solution is usually enough for one 96-well plate and preparation of the test compounds. 2. The compounds should be freshly prepared even if some compounds are stable for some time if stored at −20°C in working aliquots. For example IBMX is dissolved in dimethylsulfoxide (DMSO) at the concentration of 200 mM (1,000× stock) and it could be stored at −20°C in small aliquots (e.g., 200 μL). Prepare all compounds to a concentration of 10 mM in the same PBS with the ascorbic acid and then make the suitable dilutions. β-PEA (TAAR1 agonist) is soluble in water and PBS in the same way as 3-MT (TAAR1 agonist), isoproterenol (β2adrenergic receptor agonist), quinpirole hydrochloride (D2R agonist). Haloperidol (D2R antagonist) is soluble at 10 mM in DMSO but the subsequent dilutions (1:10, 1:100, 1:1,000, and so on) should be made in PBS (see Note 10). 3. In case of experiments at RT, all the media and solutions should be equilibrated at RT. In case of 37°C assay, switch on the plate reader heating system at least 20 min before the experiments and keep the PBS warm in a 37°C water bath (see Note 11). 4. Take the plate out of the incubator and check the cells under the microscope. Before proceeding with BRET, stick the white backing tape to the bottom of the plate to prevent light dispersion. 5. Remove the clear medium from all the wells and replace it with the PBS with calcium, magnesium and ascorbic acid. 6. Dilute the coelenterazine h stock solution 1:20 in PBS (10× solution) and add to each well to yield a final concentration of 5 μM (see Note 12). 7. For the evaluation of cAMP levels using the EPAC sensor with TAAR1, D1R or in general with a Gs-coupled receptor, after adding coelenterazine, if necessary, also add IBMX to a final concentration of 200 μM (see Note 13).

8

Dopamine/TAAR1 Signaling

115

8. To evaluate agonist activity, such as β-PEA or 3-MT on TAAR1, add the compounds 10 min after coelenterazine h (see Note 14) and immediately read the plate. If the experiment is conducted at 37°C, place the plate in the incubator during the lag of time before the agonist addition. β-PEA and 3-MT are active at low micromolar concentration (see Fig. 1a, b). 9. To evaluate antagonistic activity against a known agonist (e.g., putative antagonists of TAAR1), the different compounds are added 5 min before the agonist. 10. In case of unknown activity of the compounds or of the receptor, a positive control could be made with the endogenously expressed β2-adrenergic receptor and the agonist isoproterenol

BRET ratio

a

β-PEA

1.32

β-PEA(10–4M)

1.28

β-PEA(10–5M)

1.24

β-PEA(10–6M) β-PEA(10–7M)

1.20

β-PEA(10–8M) 1.16 1.12

0

500

b

1000 Time (sec.)

1500

2000

3-MT

3MT(10–4M)

1.32

3MT(10–5M)

1.28 BRET ratio

3MT(10–6M) 1.24

3MT(10–7M)

1.20

3MT(10–8M)

1.16 1.12

0

500

1000 Time (sec.)

1500

2000

Fig. 1. Evaluation of cAMP levels using cAMP BRET biosensor in HEK-293T cells expressing hTAAR1. (a) Time course evaluation of cAMP variations using different concentrations of β-PEA. BRET ratio is calculated as Rluc/YFP ratio and the reading started after β-PEA addition. The decrease in BRET ratio indicates an increase in cAMP levels. This experiment is performed at RT with the addition of 200 μM of IBMX. (b) Similar experiment using 3-MT as TAAR1 agonist.

116

S. Espinoza et al. 0.96

Forskolin + Quinpirole (10μM) Forskolin (25μM) Ctrl

0.94

Rluc/YFP

0.92 0.90 0.88 0.86 0.84 0.82 0.80 0

400

800

1200

Time (sec.)

Fig. 2. Evaluation of D2R activation using EPAC BRET biosensor at 37°C. Time course of cAMP variations in HEK-293T cells expressing D2R and EPAC. Forskolin is added at the beginning of the experiment at the time = 0. Quinpirole at 1 μM is added 5 min before forskolin and 5 min after coelenterazine h (see Subheading 3). Forskolin, by the activation of adenylate cyclase, increases cAMP levels and this effect is prevented by D2R activation with quinpirole.

that is active at nanomolar concentrations. Alternatively, the activator of adenylate cyclase forskolin may be used (2–20 μM). 11. For receptors such as D2R, that inhibit adenylate cyclase and the formation of cAMP, forskolin at 2–20 μM is used to increase basal cAMP level (see Note 15). 12. Add quinpirole at 1 μM 5 min after Rluc substrate, forskolin 5 min after quinpirole and then read the plate immediately (Fig. 2) (see Note 16). 13. In case of evaluation of antagonists on quinpirole effect, these compounds are added simultaneously with coelenterazine h. For example, for a haloperidol dose–response, add in each well different concentrations from 10−11 to 10−5 M (Fig. 3a). 14. For the evaluation of β-arrestin2 recruitment to D2R, the incubations are carried out at RT but the protocol remains the same, add antagonists 5 min and agonist 10 min after Rluc substrate and read the plate right after agonist addition. Haloperidol is active in preventing β-arrestin2 recruitment at low nanomolar concentrations (Fig. 3b). 15. Usually the plate is read for 20–30 min but, depending on the receptor, the time course could be longer or shorter (see Note 17). 3.4. Plate Reader Settings and Data Analysis

The plate reader used for this assay is the Mithras LB940 that allows the integration of the luminescent signal detected in the 465–505 nm and 515–555 nm windows by using filters with the appropriate band pass and the MicroWin software. The BRET

8

a

Dopamine/TAAR1 Signaling

117

140

cAMP accumulation (% of Quinpirole inhibition)

120 100 80 60 40 20 0 –20 –40 –12 –11 –10

–9

–8

–7

–6

–5

–4

–5

–4

Log [Haloperidol] (M)

b

120

β-arrestin 2 recruitment (% of Quinpirole effect)

100

80

60

40

20

0 –12 –11 –10

–9

–8

–7

–6

Log [Haloperidol] (M)

Fig. 3. Evaluation of antagonistic activity of haloperidol on Gi activation or on β-arrestin2 recruitment induced by quinpirole. (a) HEK-293 T cells are transfected with EPAC biosensor and D2R and are treated with different concentrations of haloperidol and with 1 μM of quinpirole in the presence of forskolin. (b) Dose–response of haloperidol to evaluate the inhibition of β-arrestin2 recruitment induced by 1 μM of quinpirole. Cells are transfected with D2R-Rluc and β-arrestin2-YFP. Data are plotted as SEM of 3–5 independent experiments.

signal is determined by calculating the ratio between the light emitted at 465–505 nm and the light emitted at 515–555 nm. The data are plotted as BRET ratio over the time in time course experiments. In dose–response graphs, curves are fitted using a nonlinear regression and log vs. response fit using GraphPad Prism 5 (Fig. 4). In the experiment regarding D2R antagonist effect on

118

S. Espinoza et al. 140

cAMP accumulation (% of maximum response)

120 100 80 60 40 20 0 –20 –40 –12 –11 –10

–9

–8

–7

–6

–5

–4

Log [β-PEA] (M)

Fig. 4. cAMP variations induced by a β-PEA dose–response in cells co-expressing TAAR1 and EPAC. Cells are treated with different concentrations of β-PEA and plotted as a dose– response experiment. β-PEA is used at concentrations ranging from 10−11 to 10−4 and the plate is read 15 min after agonist addition. A non-linear regression with one site-specific binding is used to draw the curve using GraphPad Prism5. Data are plotted as SEM of 3–5 independent experiments.

quinpirole activity, in the y-axis BRET ratio is transformed in the percent of quinpirole effect (Fig. 3a). In β-arrestin2 recruitment, the D2R antagonists should be plotted with the percentage of quinpirole effect over the log of the concentration of the antagonist (Fig. 3b) (see Note 18).

4. Notes 1. All the media, solutions and instruments used for cell culturing should be maintained sterile. It is important to open the medium only under the hood and clean every object by spraying some ethanol 70°C before bringing them under the hood. Bacteria or yeast contaminations of cell dishes are easily detected by looking at them under the microscope or by looking at the color and clearness of the medium. The water and the solutions for the transfection must be sterilized by filtration under the wood with a 0.22 μm filter. The water used for all procedures is a Milli-Q grade water, with a resistivity of 18.2 MΩ-cm and total organic content of less than 5 parts per billion. 2. Depending on the speed of growth of the clone of HEK-293T cells or the time of the day of seeding it is possible to seed less

8

Dopamine/TAAR1 Signaling

119

or more cells. It is important to have cells at 70–80% confluency the day of transfection. 3. Since each well (of the 96-well plate) should contain approximately 75,000 cells, usually one 100-mm plate is enough for single transfection to yield a full 96-well plate. In case of multiple transfections and when the experiment design calls for fewer cells, 35-mm dishes can be used to transfect cells. In this case the procedures remain the same by just scaling down all the reagents and solutions used proportionally. In contrast, when many cells with a single transfection are needed, more than one 100-mm dish could be used. In order to have a homogeneous sample, mix all the similar transfections together before seeding them in the assay plate. 4. The critical parameter for the success of the transfection is the pH of the HBS 2× solution. It is suggested to prepare a fairly good amount of solution since it could be tedious to find the right pH each time the solution has to be prepared. After all the reagents are dissolved, the pH should be adjusted to 7.1 ± 0.05. Since there are some variables such as incubator or pH meter dysfunctions or unpredictable ones, it is advisable to prepare different HBS 2× solutions in a range of pH from 7.0 to 7.2 and test them in different transfections in order to find the one with the highest transfection efficacy. Then sterilize by filtration and aliquot into 50 mL Falcon tube and store at −20°C. Working aliquots can be stored at 4°C. 5. Normally, keep the DNA at −20°C but for long-term storage put it at −80°C. Be sure that the solution is clear and there are no precipitate or strange materials that could indicate the presence of molds. In such a case, it is imperative to re-prepare the plasmid. 6. For TAAR1 it is better to do the experiments 24 h after transfection. D2R and β-arrestin2 also have a strong protein expression after 24 h of transfection, so in cases of co-transfecting TAAR1 and D2R, it is possible to do experiments at 24 h. 7. Sometimes the poly-d-lysine treatment is prolonged for hours, for low adherent cells, but usually for HEK cells 10–20 min should be enough. It is possible to reuse the poly-d-lysine up to three times, so re-collect it and keep it sterile in the refrigerator after two uses. 8. Some HEK clones are less or more “attached” to the dish. In some case, while passaging cells, PBS is enough to detach the HEK cells; in other cases trypsin is necessary. The latter is especially the case after a calcium-phosphate transfection. 9. With these dilutions, 75,000 cells/well is seeded and should be enough to have a good signal to noise ratio. The number may change from 25,000 to 100,000 per well depending on

120

S. Espinoza et al.

the BRET signal, cells availability (e.g., in case of BRET experiments in primary neurons) and other variables. 10. To minimize the final concentration of DMSO to a maximum of 0.1–0.5% in order to avoid DMSO toxicity, haloperidol and other water-insoluble compounds should be diluted as much as possible in PBS. Appropriate controls should be used to verify the inactivity of the dissolving medium. 11. For β-arrestin2 recruitment to D2R, RT experiment is more convenient since the recruitment is slower and more visible when analyzing the data. cAMP evaluation with EPAC sensor could be done at 37°C or even at RT. The desensitization of the cAMP response of the receptor of interest is more readily observable at 37°C. 12. For BRET experiments, it is convenient, in term of calculations, to have a final volume of 100 μl. In this case, add 10 μL of coelenterazine h and prepare all compounds as 10× solutions in Eppendorf tube in order to add 10 μL for each of them that have to be tested. In this case, make the calculations of how many microliters of PBS that have to be added at the beginning of the experiment (e.g., for testing 1 agonist (10 μL) and 1 antagonist (10 μL), plus coelenterazine h (10 μL) and IBMX (10 μL), put 60 μL of PBS in each well in order to attain a final volume of 100 μL, 10 + 10 + 10 + 10 + 60 = 100). 13. To increase cAMP levels, IBMX may be used. IBMX blocks the degradation of the cAMP by endogenous phosphodiesterases and may be useful to increase the signal induced by the receptor of interest. 14. The oxidation of coelenterazine h by Rluc takes several minutes to equilibrate, so to have a stable and good signal it is better to wait at least 5 min (ideally 10 min) from the addition of the substrate. If the transfection is efficient, each well of the plate consists of one data point and thus it is not necessary to perform the experiment in duplicates. Duplicate reads however are often recommended. 15. In vitro, without any stimuli, the inhibition of adenylate cyclase does not induce a significant decrease of cAMP that is detectable by our technique. So, to study Gi-coupled receptors such as D2R activity, adenylate cyclase has to be activated by forskolin. 16. Quinpirole has its maximum effect at 1 μM and the EC50 is 5.7 nM. 17. The effect of the agonist on cAMP production is quite fast even at RT, so it is necessary to work rapidly in order to measure the beginning of the effect. As an alternative, the injector in the machine could be used to dispense the agonist to different wells, but take into account the number of compounds

8

Dopamine/TAAR1 Signaling

121

that have to be dispensed and the availability as well as the cost of the substances. 18. Another machine tested for this application is the Infinite F500 (Tecan). It has two appropriate filters to select the light emitted by Rluc and the YFP and as the Mithras two injectors and the heating system. While the Mithras is fairly more sensitive, Infinite’s software is more easy-to-use and flexible.

Acknowledgments Supported in part by research grants from the Michael J. Fox Foundation for Parkinson’s Research, Fondazione Compagnia di San Paolo (Torino, Italy) and research grant from F. Hoffmann-La Roche Ltd. (Basel, Switzerland) to Raul R. Gainetdinov. Bernard Masri was a recipient of a European Marie-Curie Outgoing International Fellowship (FP6—2005-Mobility-6). Ali Salahpour is supported by grants from Canadian Institute of Health Research (CIHR # 210296) and Natural Sciences and Engineering Council of Canada (NSERC # 386422). References 1. Beaulieu JM, Gainetdinov RR (2011) The physiology, signaling and pharmacology of dopamine receptors. Pharmacol Rev 63: 182–217 2. Beaulieu JM, Gainetdinov RR, Caron MG (2007) The Akt-GSK-3 signaling cascade in the actions of dopamine. Trends Pharmacol Sci 28:166–172 3. Lefkowitz RJ, Shenoy SK (2005) Transduction of receptor signals by beta-arrestins. Science 308:512–517 4. Beaulieu JM, Gainetdinov RR, Caron MG (2009) Akt/GSK3 signaling in the action of psychotropic drugs. Annu Rev Pharmacol Toxicol 49:327–347 5. Masri B, Salahpour A, Didriksen M, Ghisi V, Beaulieu JM, Gainetdinov RR, Caron MG (2008) Antagonism of dopamine D2 receptor/ beta-arrestin 2 interaction is a common property of clinically effective antipsychotics. Proc Natl Acad Sci U S A 105:13656–13661 6. Bunzow JR, Sonders MS, Arttamangkul S, Harrison LM, Zhang G, Quigley DI, Darland T, Suchland KL, Pasumamula S, Kennedy JL, Olson SB, Magenis RE, Amara SG, Grandy DK (2001) Amphetamine, 3,4-methylenedioxymethamphetamine, lysergic acid diethylamide, and metabolites of the cate-

7.

8.

9.

10.

cholamine neurotransmitters are agonists of a rat trace amine receptor. Mol Pharmacol 60:1181–1188 Borowsky B, Adham N, Jones KA, Raddatz R, Artymyshyn R, Ogozalek KL, Durkin MM, Lakhlani PP, Bonini JA, Pathirana S, Boyle N, Pu X, Kouranova E, Lichtblau H, Ochoa FY, Branchek TA, Gerald C (2001) Trace amines: identification of a family of mammalian G protein-coupled receptors. Proc Natl Acad Sci U S A 98:8966–8971 Lindemann L, Meyer CA, Jeanneau K, Bradaia A, Ozmen L, Bluethmann H, Bettler B, Wettstein JG, Borroni E, Moreau JL, Hoener MC (2008) Trace amine-associated receptor 1 modulates dopaminergic activity. J Pharmacol Exp Ther 324:948–956 Bradaia A, Trube G, Stalder H, Norcross RD, Ozmen L, Wettstein JG, Pinard A, Buchy D, Gassmann M, Hoener MC, Bettler B (2009) The selective antagonist EPPTB reveals TAAR1mediated regulatory mechanisms in dopaminergic neurons of the mesolimbic system. Proc Natl Acad Sci U S A 106:20081–20086 Barak LS, Salahpour A, Zhang X, Masri B, Sotnikova TD, Ramsey AJ, Violin JD, Lefkowitz RJ, Caron MG, Gainetdinov RR (2008) Pharmacological characterization of

122

11.

12.

13.

14.

S. Espinoza et al. membrane-expressed human trace amineassociated receptor 1 (TAAR1) by a bioluminescence resonance energy transfer cAMP biosensor. Mol Pharmacol 74:585–594 Sotnikova TD, Beaulieu JM, Espinoza S, Masri B, Zhang X, Salahpour A, Barak LS, Caron MG, Gainetdinov RR (2010) The dopamine metabolite 3-methoxytyramine is a neuromodulator. PLoS One 5:e13452 Ayoub MA, Pfleger KD (2010) Recent advances in bioluminescence resonance energy transfer technologies to study GPCR heteromerization. Curr Opin Pharmacol 10:44–52 Marullo S, Bouvier M (2007) Resonance energy transfer approaches in molecular pharmacology and beyond. Trends Pharmacol Sci 28:362–365 Ni Q, Titov DV, Zhang J (2006) Analyzing protein kinase dynamics in living cells with FRET reporters. Methods 40:279–286

15. Pfleger KD, Seeber RM, Eidne KA (2006) Bioluminescence resonance energy transfer (BRET) for the real-time detection of protein-protein interactions. Nat Protoc 1:337–345 16. Pfleger KD, Eidne KA (2006) Illuminating insights into protein-protein interactions using bioluminescence resonance energy transfer (BRET). Nat Methods 3:165–174 17. Salahpour A, Espinoza S, Masri B, Lam V, Barak LS, Gainetdinov RR (2012) BRET biosensors to study GPCR biology, pharmacology, and signal transduction. Front Endocrinol (Lausanne) 3:105 18. Espinoza S, Salahpour A, Masri B, Sotnikova TD, Messa M, Barak LS, Caron MG, Gainetdinov RR (2011) Functional interaction between Trace Amine Associated Receptor 1 (TAAR1) and dopamine D2 receptor. Mol Pharmacol 80:416–425

Chapter 9 Dopaminergic Regulation of Dendritic Calcium: Fast Multisite Calcium Imaging Wen-Liang Zhou, Katerina D. Oikonomou, Shaina M. Short, and Srdjan D. Antic Abstract Optimal dopamine tone is required for the normal cortical function; however it is still unclear how corticaldopamine-release affects information processing in individual cortical neurons. Thousands of glutamatergic inputs impinge onto elaborate dendritic trees of neocortical pyramidal neurons. In the process of ensuing synaptic integration (information processing), a variety of calcium transients are generated in remote dendritic compartments. In order to understand the cellular mechanisms of dopaminergic modulation it is important to know whether and how dopaminergic signals affect dendritic calcium transients. In this chapter, we describe a relatively inexpensive method for monitoring dendritic calcium fluctuations at multiple loci across the pyramidal dendritic tree, at the same moment of time (simultaneously). The experiments have been designed to measure the amplitude, time course and spatial extent of action potential-associated dendritic calcium transients before and after application of dopaminergic drugs. In the examples provided here the dendritic calcium transients were evoked by triggering the somatic action potentials (backpropagation-evoked), and puffs of exogenous dopamine were applied locally onto selected dendritic branches. Key words: Action potential, Backpropagation, Voltage-gated calcium channels, Dopaminergic modulation, Dopamine receptors, Dendritic excitability, Phasic dopamine signal

1. Introduction The level of prefrontal cortex dopamine (DA) is critical for modulating normal cognitive/behavioral processes (1). One important hypothesis is that deviations from the critical levels can severely disrupt cognition and result in mental disorders (2). At present, it is not known how such levels of dopamine interact with prefrontal cortex (PFC) neural circuits. Electron microscope studies in monkey and human have found that dopamine synaptic contacts onto cortical neurons are located almost exclusively on the distal dendrites Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_9, © Springer Science+Business Media, LLC 2013

123

124

W.-L. Zhou et al.

and spines of distal dendrites (3). The same distal dendritic segments of pyramidal neurons primarily receive thousands of excitatory glutamatergic inputs (4). The first stage of synaptic integration takes place at the actual site of glutamatergic inputs, in thin (basal, oblique and apical tuft) dendrites of pyramidal neurons (5– 7). Dendritic function is closely linked to local fluctuations in cytosolic calcium. Calcium ions in the intracellular compartment trigger and regulate an impressive number of basic cellular functions (8–12). Based on recent published studies one can identify five main categories (Fig. 1a–e) of calcium transients that occur in dendrites of CNS neurons during biological processes: (A) Calcium sparks—spontaneous miniature releases (13). (B) Massive synaptically evoked calcium release from internal stores (14–16). (C) Action potential-mediated calcium influx (17–20). (D) Synaptically evoked dendritic calcium influx (21–25). (E) Local-dendritic-spike-associated calcium influx (26–28). In addition to dendritic calcium transients (Fig. 1) the modern experimental techniques have characterized action potential (AP)induced calcium transients in axons (29) and more impressively in axon terminals, both in vitro (30) and in vivo (31) settings. The experimental tools for monitoring calcium transients in small neuronal compartments have undergone a steady improvement in the

Fig. 1. Five categories of dendritic calcium signaling. Pyramidal neurons have basal, oblique and apical dendrites. The amplitudes and spatial distributions of neuronal calcium transients are indicated by the intensity and shape of black shadings. (a) Fast but small spontaneous releases of calcium have been detected along the apical dendrites of hippocampal pyramidal neurons (arrow), as well as in the apical oblique branches and the cell body (not shown but see ref. 13). These spontaneous events (sparks) represent miniature releases from internal stores (diameter = 2 μm, duration = 100 ms) and could be important in brain development (11). (b) Synaptically evoked internal release of calcium engulfs the proximal segment of the apical dendrite and slowly propagates into the cell body (15). (c) Action potential propagates backward from the axon initial segment into the dendritic tree (backpropagation) causing activation of dendritic voltage-gated calcium channels (18). AP-associated dendritic calcium signal regulates synaptic plasticity (61). (d) Activation of glutamatergic presynaptic axon terminals causes the opening of synaptic receptors and the influx of calcium into dendritic spines (23). (e) Synchronous activation of spatially segregated glutamatergic presynaptic terminals triggers a local regenerative dendritic potential, which is manifested by a spatially restricted peak of calcium influx (27).

9

Dopamine and Dendritic Calcium

125

last 40 years (22, 32–36). An optical measurement of dendritic calcium fluctuation has become a robust, reliable, and affordable technique, which is now regularly used in a large number of laboratories around the world. The time is right to ask whether and how stimulation of dendritic dopamine receptors modulate dendritic calcium signaling as shown in Fig. 1. In the current chapter we focus on only one of the five aforementioned categories: AP-associated dendritic calcium transients (Fig. 1c). The backpropagating action potentials invade distal dendritic segments, depolarize the dendritic membrane and open voltagegated calcium channels (18). The dendrites of PFC neurons express D1 receptors (37, 38) and N-type calcium channels (18, 39). A physical link between D1R and N-type calcium channels has been implicated as a key mechanism in the dopamine-induced suppression of dendritic calcium influx during action potential backpropagation (40). In their study the calcium measurements were performed from only one dendrite class (apical) and at only one location at the time before and after bath perfusion of dopamine (40). Although Kisilevsky et al. (40) provide experimental evidence for dopamine-induced suppression of dendritic calcium influx, this issue remains controversial because another group claimed that in the same brain region, same cell type, same dendrite (apical), same type of stimulation (dopamine in the bath) has no effect on AP-mediated calcium influx (41). In this chapter, we revisit dopaminergic modulation of action potential backpropagation, and we combine local dopamine applications with multisite dendritic imaging. We attempt to mimic the phasic DA signal (42) by locally applying DA from a glass pipette for a brief period of time (2 s). We address other classes of dendrites (basilar branches located in the deeper cortical layers where dopaminergic fibers cluster). We investigate how different regions along the same dendrite respond to DA receptor stimulation (a spatial aspect of modulation). Finally, we analyze the voltage waveform of a backpropagated AP in the same dendritic segments, where phasic DA stimuli caused suppression of the calcium influx.

2. Materials 2.1. Solutions and Dyes

1. Extracellular solution. Artificial cerebrospinal fluid (ACSF, in mM): 125 NaCl, 26 NaHCO3, 10 glucose, 2.3 KCl, 1.26 KH2PO4, 2 CaCl2, and 1 MgSO4 (pH 7.4, osmolality 300–310 mosmol/L). 2. Intracellular solution. We use a standard gluconate-based intracellular solution containing (in mM): 135 potassium gluconate, 2 MgCl2, 3Mg-ATP, 10 phosphocreatine, 0.3 GTP,

126

W.-L. Zhou et al.

and 10 Hepes (pH 7.3, adjusted with KOH; osmolality 290–300 mosmol/L). 3. Calcium sensitive dyes: Calcium Green-1 (CG-1), Oregon Green Bapta-1 (OGB-1), Fluo-5F, and bis-fura-2 were purchased from Invitrogen (Carlsbad, CA) and dissolved directly into the intracellular solution at 150–250 μM. Voltage-sensitive dyes JPW1114 and JPW3028 were kindly provided by J.P. Wuskell and Leslie Loew (University of Connecticut Health Center, Farmington, CT) and dissolved directly into the intracellular solution at 400 μM. 2.2. Electrophysiology

1. Amplifiers. We use Axon Instruments Multiclamp 700B, a digitizer, Digidata Series 1322A, and Clampex software for acquisition, and Clampfit software for the analysis of electrical recordings (Molecular Devices, Sunnyvale, CA). One personal computer (PC) is devoted to electrical recordings and for taking photographs for morphological analysis of fluorescently labeled neurons. 2. Second analog-to-digital board. The Neuroplex system (RedShirtImaging LLC, Decatur, GA) is running on a PC devoted to fast optical imaging only (e.g., calcium imaging). Neuroplex system is equipped with an A-D board for storing and synchronizing electrical data with the corresponding optical data. During a typical experiment the same electrical signal, which is recorded by Axon Instruments apparata (aforementioned) on the first PC, is also stored inside the optical file on a second PC. 3. Micromanipulators. Two motorized micromanipulators (P-285, Sutter Instrument, Novato, CA) are used for positioning the patching pipette onto the cell body, and the dopamine application pipette near the dendrite.

2.3. Optical Imaging Station

1. Microscope. Standard upright microscope for patching brain slices (Olympus BX51WI, Olympus Inc., Japan). 2. Lenses. 40× water immersion objective with high N/A. 5× objective for finding pipettes. 3. Microscope modules. Epi-illumination module, filter cubes, and two camera ports for fluorescence imaging (Olympus Inc.). 4. Camera port-1. One camera port is taken by a simple CCD camera for infrared DIC video microscopy (there are many models available—we use Dage IR-1000). 5. Camera port-2. The second camera port is taken by a fast lownoise low-resolution data acquisition CCD camera (RedShirtImaging LLC, Decatur, GA). We use 80 × 80 pixel model (NeuroCCD-SMQ), which is suitable for both calcium

9

Dopamine and Dendritic Calcium

127

and voltage imaging from dendrites of CNS neurons, thanks to superior sensitivity (low noise) and very fast frame rates (5, 27, 43, 44). The full frame rate is 2,000 frames per second. This camera can take 5,000 pictures per second after binning or 10,000 pictures per second after a partial reduction of the frame size (cropping) (45). 6. Software. Data acquisition software Neuroplex (RedShirtImaging) is endowed with several useful features including the examination of records immediately following the acquisition trial, an automatic spatial averaging of individually selected pixels, and spike-triggered averaging of consecutive sweeps during the actual data acquisition. 7. Light source. For epi-illumination we use 250 or 150 W xenon arc lamps (Opti-Quip, Highland Mills, NY). These are lowripple (low-noise) lamps that have excellent power for wholefield illumination, and they cover the entire relevant range for dye excitations; from 340 to 700 nm. This is especially useful for sequential voltage-calcium imaging using a cocktail of dyes (27) or for combining AlexaFluor594 with green calcium dyes (Calcium Green-1, Oregon Green Bapta-1, and Fluo-5F). For important practical details on the use of Opti-Quip arc lamps (see Notes 1 and 2). 8. Mechanical shutter. Electro-Programmable shutter system CS35-Uniblitz was purchased from Vincent Associates (Rochester, NY). The CS35 shutter is equipped with reflective blades. This option protects the shutter mechanism from the light source’s damaging effects by reflecting the energy away from the blade surface. 9. Optical filters. Optical filters were purchased from Chroma Technology (Rockingham, VT) and Omega Optical (Brattleboro, VT). The filter set (cube) for Fluo-5F consists of an Omega exciter 500AF25 (485–510 nm bandpass), dichroic 525DRLP and emitter 530ALP (530 nm longpass). Filters for AlexaFluor594 were a Chroma exciter HQ580/20× (570– 590 nm bandpass), dichroic Q595LP and emitter HQ630/60 m (600–660 nm bandpass). The filter cube for Ca-Green-1 contains an Omega exciter 500AF25, dichroic 525DRLP, and emitter 530ALP. The same filter set is used for voltage imaging with JPW1114 or JPW3028 voltage-sensitive dyes. This filter set consists of a Chroma exciter D510/60 (480–540 nm bandpass), dichroic 570dcxru and emitter E600lp (600 nm longpass). The filter cube for bis-fura-2 contains a Chroma exciter D380/30× (365–395 nm bandpass), dichroic 400dclp, and emitter E470lp (470 nm longpass). 10. Vibration isolation table. We use BM-1 platforms made by Minus K Technology, Inc. (Inglewood, CA).

128

W.-L. Zhou et al.

3. Methods 3.1. Acute Slice Preparation

Preparation of quality slices is a critical step in the experiment. Sprague-Dawley rats (postnatal day 21–35) were deeply anesthetized with isoflurane and decapitated according to an animal protocol approved by the UConn Health Center Animal Care and Use Committee. Coronal brain slices (300 μm thick) were harvested from the frontal lobe (anterior to genu of corpus callosum) in gassed (95% O2 and 5% CO2), ice-cold artificial cerebrospinal fluid (ACSF). Note that we cut slices in the same solution (ACSF) which is used for experimental recordings (see Note 3). From the ice-cold cutting chamber the slices were transferred to a warm and oxygenated holding chamber (35°C) and incubated for 30 min at 35°C. Following a 30 min incubation, the slices were removed from the warm water-bath and kept at room temperature (1–6 h) before being transferred to the recording chamber. Prefrontal cortical slices of good quality have an abundance of pyramidal cells in superficial (Layers 2–3) and deep layers (Layers 5–6). Healthy pyramidal cells have oval cell bodies, smooth membranes that appear to shine under infrared differential interference contrast (IR DIC) video microscopy, and very soft edges. Pyramidal cells that stand out from the brain slice background, show rugged edges and strong contrast are often bad and unhealthy cells. One side of the brain slice is always better than the other. It is important to quickly examine the slice (using a 40× objective) prior to securing it with a slice-anchor (see Note 4).

3.2. Patch Electrode Recordings and Dye Injections

Patch pipettes were pulled from borosilicate glass (G150F-3, Warner Instruments, Hamden, CT, USA) on a P-97 microelectrode puller (Sutter Instruments, Novato, CA). The ideal pipette resistance for loading pyramidal neurons with fluorescent dyes is 7 MΩ (see Note 5). All recordings were performed at 33–34°C. Whole-cell recordings from layer 5 pyramidal neurons were carried out using a Multiclamp 700B amplifier (Materials) and digitized with two input boards: (1) Digidata Series 1322A (Subheading 2.2, item 1) at a 10 kHz sampling rate and (2) Neuroplex (Subheading 2.2, item 2) at a 4 kHz sampling rate. Only cells having a membrane potential more hyperpolarized than −50 mV (not corrected for liquid junction potential) and AP amplitudes >70 mV (measured from the base line) were included in the study. Voltagesensitive dye (JPW1114 or JPW3028) and calcium-sensitive dyes (Ca-Green-1, Oregon Green Bapta-1, bis-fura-2, and Fluo-5F), as well as AlexaFluor594, were dissolved in intracellular solution and loaded into the patch pipette. Impurities and dust particles in the intracellular solution represent major obstacles for formation of the seal and later for dye-injection into the cytosol (see Note 6). To avoid extracellular deposition of the fluorescent dyes, glass

9

Dopamine and Dendritic Calcium

129

pipettes were filled from the tip with dye-free solution by applying negative pressure (front-loading), and were back-filled with dye solution (back-loading). This procedure is essential for loading voltage-sensitive dyes into neurons in brain slices (43), and it is considerably less important for calcium-sensitive dyes; though it may improve the viability of calcium-loaded neurons in long experiments. Intracellular staining was achieved by free diffusion of the dye from the pipette into the cell body. Duration of dye loading depends on the cell type, size and shape of the patch pipette, and most importantly on the water-solubility of the fluorescent dye. For example, voltage-sensitive dyes JPW1114 and JPW2030 are lipophilic, and it takes at least 2 h to fill the apical tuft branches. In the case of voltage-sensitive dyes the dye loading pipette must not stay in whole-cell configuration for longer than 60 min; because the overloading of the cell body compartment causes pharmacological and photodynamic damage (46). To prevent toxic effects of voltage-sensitive dyes, after 40–60 min of dye-injection an outsideout patch was formed and the patch electrode removed (46, 47). Dye-injected neurons were next incubated for 2–3 h at room temperature and repatched with a dye-free pipette, just prior to the optical recording session. Voltage-sensitive dye recordings from dendrites of CNS neurons are beyond the scope of this chapter. For detailed description of voltage-sensitive dye method see the most recent protocol (48). 3.3. Calcium Imaging

Calcium-sensitive dyes are soluble in water and it takes approximately 30–35 min to properly load the majority of basilar and oblique dendritic branches in layer 5 pyramidal neurons. For the most distal apical tuft branches it may take more than 100 min of dye loading. AlexaFluor594 is co-applied with calcium-sensitive dyes to allow a quicker and better examination of the dendritic tree, as well as to aid proper positioning of neurons inside the visual field of the NeuroCCD-SMQ (Fig. 2), without having to excite the calcium dye (see Note 7). While the calcium-sensitive dye is diffusing from the patch pipette into the soma, and from the soma into the target dendritic branch, the amplitude of AP-induced calcium transient will change. The unstable amplitude of a calcium transient during control measurements may compromise the results, therefore, it is important to evaluate the time-dependence of evoked dendritic calcium transients in the absence of any conditioning (e.g., before dopaminergic stimulation). We have established that approximately 45 min from the beginning of dye loading procedure (whole-cell breakthrough) the baseline measurements in basilar segments of the dendritic tree become stable and remain at one fixed level for another 30–40 min (Fig. 2).

130

W.-L. Zhou et al.

Fig. 2. Calcium imaging: Establish the baseline prior to testing working hypotheses. A PFC Layer 5 pyramidal neuron was loaded for 30 min with OGB-1 [200 μM] and Alexa Fluor [60 μM]. The basilar dendritic tree was projected onto the NeuroCCD using a ×40 objective lens. Action potential was evoked by the somatic current injection and the resulting dendritic signals were recorded every 2–4 min from the entire visual field. Each panel (a–d) is devoted to one region of interest marked by white box. The peak amplitude of calcium transient (obtained by averaging outputs of camera pixels inside the box) is expressed as dF/F (%) and plotted versus time in the graph on the right. Time zero marks the end of the dye-loading phase and it corresponds to the 30th minute after the whole-cell breakthrough. After 30 min of dye loading each dendritic segment (a–d) exhibits a relatively stable AP-induced calcium signal for the next 35 min, which is plenty of time to perform a biological experiment.

3.4. Calcium Imaging of Dopamine-Induced Changes

In order to mimic phasic dopaminergic signals (42) we loaded 5 mM dopamine into a glass micropipette. With the aid of a motorized micromanipulator the DA application micropipette was positioned in the vicinity of one basal branch. There is 20–30 μm from the tip of micropipette to the dendritic shaft (Fig. 3a). The positioning of the glass pipette onto a selected dendritic branch was done by alternating between IR DIC (not shown) and fluorescence video microscopy (Fig. 3a). Dopamine was pressureejected for 2 s (computer-driven picospritzer), just prior to a calcium-imaging sweep. Note that [5 mM] refers to the concentration of dopamine inside the application pipette. The concentration of dopamine that reaches the dendritic membrane at the end of a 2 s long puff is likely one or two orders of magnitude lower. The dopamine-induced suppression in dendritic calcium transient is only momentary, as the subsequent sweeps show rapid recovery of the signal amplitude (Fig. 3b, Wash).

3.5. Spatial Aspect of Dopamine-Induced Changes

In their current state, the confocal microscopy and two-photon imaging methods are not capable of monitoring the spatial distribution of dopamine-induced changes across the dendritic tree at a 200 Hz frequency (5 ms per full frame, Fig. 4). The present experimental setup has been designed to monitor AP-associated transients from multiple loci (regions of interest, ROIs) across several dendritic branches (Fig. 4), at the same time (simultaneously). While the “target” dendrite (Fig. 4, ROI 2) is receiving a phasic dopaminergic stimulus (DA), the neighboring branches belonging to the same nerve cell (Fig. 4, ROI 5–7) can be used as an ideal control (same cell, same instant of time). The spatial resolution of the system, when used with a 40× objective, is approximately

9

Dopamine and Dendritic Calcium

131

Fig. 3. Calcium imaging: phasic dopamine stimulation. (a) A PFC Layer 5 pyramidal neuron was loaded for 25 min with OGB-1 [200 μM] and Alexa Fluor [60 μM]. Time zero marks 25th minute after the whole-cell breakthrough. The basilar dendritic tree was projected onto the NeuroCCD using a ×40 objective lens and a red fluorescence cube for Alexa Fluor 594 (Materials). Action potential was evoked by a somatic current injection and the resulting dendritic signal was recorded from 8 camera pixels inside the white box, using the green fluorescence cube (calcium imaging). (b) AP-mediated dendritic calcium transients were measured in intervals of 1–4 min. Each trace is the product of 8-pixel spatial averaging. Dopamine was ejected for 2 s (total duration) just prior to the optical recording sweep. Recordings obtained before DA ejection are considered control recordings (Ctrl.). Recordings obtained after DA ejection are used to estimate the temporal dynamics of washout.

Fig. 4. Calcium imaging: spatially restricted effect of a phasic dopamine stimulus. A PFC Layer 5 pyramidal neuron was loaded for 30 min with CG-1 [200 μM] and Alexa Fluor [60 μM]. The basilar dendritic tree was projected onto the NeuroCCD using a ×40 objective lens. Scale bar, 50 μM. A single action potential was evoked by the somatic current injection and the resulting dendritic signals were recorded simultaneously from the entire visual field. Only seven regions of interest (ROIs) are selected for display (1–7). Optical traces were acquired before (Control) and after a local dopamine puff (DA). Control recordings are marked by a dashed grey line. Dopamine recordings are marked by thick black line. ROI 0 indicates a somatic whole-cell recording of evoked action potential. Asterisk marks dendritic segment experiencing the most severe amplitude reduction in response to local dopamine puff (duration, 2 s). Note that ROIs 5, 6 and 7 experience no change at the same moment of time (Antic lab, unpublished data).

5 μm × 5 μm per pixel. That is to say that each camera pixel covers an area 5 μm × 5 μm in the object field. With an array composed of 80 × 80 pixels (NeuroCCD-SMQ) we can simultaneously sample every branch in the visual field of roughly 400 μm × 400 μm. Such resolution provides ample means to study the spatial extent of the

132

W.-L. Zhou et al.

dopamine effect on dendritic calcium flux. In the example shown in Fig. 4 the greatest suppression of calcium influx was detected in dendritic segment “ROI 2,” which is closest to the dopamine application site (DA). Due to the orientation of the ejection pipette and the direction of dopamine jet (Fig. 4, arrow), the membranes in the dopamine path were all affected (ROIs 1, 3, and 4), while basal dendrites away from the dopamine stream showed no change in signal amplitude (Fig. 4, ROIs 5–7). 3.6. Voltage–Calcium Imaging of DopamineInduced Changes

The amplitude of a backpropagating action potential is not uniform along a weakly excitable dendrite (49, 50). Depending on a neuron type (e.g., Mitral cell versus Purkinje neuron), or dendrite type (e.g., basal dendrite versus apical tuft dendrite) and the physiological state of the dendrite (e.g., recent synaptic history), the amplitude of backpropagating AP may vary dramatically (51, 52). Some neurotransmitters/neuromodulators, including dopamine itself, have been reported to change AP amplitude in dendrites (53, 54). We have observed a dopamine-induced suppression of AP-associated dendritic calcium signal (Figs. 3 and 4). The first logical question that comes to mind is whether this suppression was caused by dopaminergic modulation of the action potential waveform? In order to address this issue we performed voltagesensitive dye recordings and calcium-sensitive dye recordings from the same neuron, same dendrite, same dendritic segment (Fig. 5a, white box) before and after local dopamine application (Fig. 5b). To achieve this dual-mode optical recording, the neurons were loaded with a cocktail consisting of one voltage-sensitive dye

Fig. 5. Voltage: Calcium imaging of DA-induced changes. A PFC Layer 5 pyramidal neuron was loaded for 45 min with a mixture containing bis-fura [200 μM] and JPW3028 [400 μM] and then the loading pipette was removed. Following a 90 min of post-loading incubation the neuron was repatched with solution containing bis-fura but not JPW3028. (a) The basilar dendritic tree was projected onto the NeuroCCD using a ×40 objective lens. Action potential was evoked by the somatic current injection and the resulting dendritic signals were recorded from 8 camera pixels inside the white box (region-of-interest, ROI). Scale bar, 50 μm. (b) The AP-evoked dendritic signal was first recorded using a filter cube for voltage-sensitive dye JPW3028 (Voltage) before (Control), upon dopamine ejection (DA) and 2 min after the dopamine ejection (Wash). Five minutes later, the AP-evoked signal was recorded using a filter cube for bis-fura (Calcium) in three conditions (Control, DA and Wash). The same subset of 8 pixels (inside the ROI) was used to produce a spatial average in both voltage and calcium modes—traces displayed in (b). Note that during DA stimulus the dendritic transients experience amplitude reduction in calcium channel, but not so prominent in voltage channel (Antic lab, unpublished data).

9

Dopamine and Dendritic Calcium

133

(JPW3020) and one calcium-sensitive dye (bis-fura-2). It is important to note that the excitation spectra of these two dyes do not overlap. Bis-fura excitation filter is set at 380 nm, while JPW2038 excitation filter is set at 520 nm; more than 100 nm away from each other. In a typical experiment, AP-associated dendritic transients were recorded with one filter set (filter cube) and then the measurements were repeated using the same stimulation (single AP) but different filter cube. In between optical recording sweeps the X-Y position of the dendritic tree and dendritic focus were kept fixed. Using the sequential voltage-calcium recordings we found that brief dopaminergic stimulations strongly affect calcium (Fig. 5b, Calcium) but not so strongly the voltage signals (Voltage) in the same dendritic segment. 3.7. Advantages and Disadvantages

Recent studies have emphasized the advantages of two-photon microscopy for studying calcium dynamics in thin branches of CNS neurons (23). Here we point out the disadvantages of confocal and two-photon microscopy approaches, in direct comparison to our system. 1. The experimental setup described in this chapter is several times cheaper than a commercially available two-photon system. While only a selected few laboratories can afford twophoton and confocal microscope-based experimental rigs, our system can be installed on virtually any brain slice electrophysiology setup with a simple addition of one fast CCD camera, one arc lamp and one mechanical shutter. 2. The maintenance of our system is easier, cheaper and faster. The same is true for time and effort that must be invested to train users to perform experiments. 3. High-frequency optical recordings from several dozen loci at the same moment of time (Fig. 4) are currently not feasible with two-photon systems, and therefore the complex spatiotemporal patterns of synaptic integration (27, 44, 55–59), or neuromodulation (60), are difficult to study on these systems. 4. Expensive purchase of laser lines and extensive rebuilding is necessary to adapt a confocal system to perform sequential voltage-calcium recordings shown in Fig. 5. In the case of our system the user only needs to purchase one additional filter cube.

4. Notes 1. Overheating of the light source. Both 250 and 150 W xenon arc lamps made by Opti-Quip, Highland Mills, NY, USA have excellent stability of the photon output but unfortunately they

134

W.-L. Zhou et al.

lack proper cooling systems. Both lamps are prone to frequent breakdowns, which is a major setback in a research project. To prevent frequent breakdowns one should install computer fans on both ends of the lamp house (Model 770U, Opti-Quip). The bottom fan should direct air upwards into the bottom vents. The top fan should collect air coming out from top vents. In this way the lamp house is kept comfortably warm (no overheating), which dramatically extends the life of the Xenon bulb and reduces the number of malfunctions. However, the fans must not be in any physical contact with the lamp house, because the lamp house is rigidly mounted to the back of the microscope and provides the direct pathway for mechanical vibrations to get introduced into optical records. To hold the two fans in position we use metal rods attached to the wall outside the electrophysiology rig. 2. Mounting of the light source. We do not use optic guides because regardless of their quality the optic fibers significantly reduce the number of photons emanating from the light source. Instead, the arc lamp is mounted directly to the back of the microscope. Between the lamp and the microscope we position (secure) a custom-made aluminum box, which provides a narrow spacing for insertion of the optical shutter. The optical shutter must not be in any physical contact with the custom-made aluminum box, because the box is rigidly mounted to the microscope and provides a direct pathway for mechanical vibrations to get introduced into optical records. Instead, the shutter is suspended on a horizontal rod that branches off a large vertical rod, which is mounted outside the antivibration table by its bottom end. In this way mechanical vibrations caused by shutter opening are interrupted on the way to the microscope by the vibration isolation table (Subheading 2.3, item 10). 3. Cold ACSF. Note that we cut slices in the same solution (ACSF) which is used for experimental recordings (Subheading 2.1, item 1). We obtain good quality slices if we cool down the ACSF to the point where thin ice flakes are floating in the beaker. However, this initial cooling is not sufficient as ACSF rapidly warms during slicing on a tissue slicer. In order to slow down warming of the ACSF we introduce a cold aluminum block (5 cm × 3 cm × 1 cm) inside the ASCF filled cutting chamber. The aluminum block and the cutting chamber (removable parts) are both stored in the freezer between slicing. 4. Slice flipping. We use a custom-made anchor (nylon grid) to secure brain slices to the bottom of the recording chamber. Before final positioning of the anchor, the brain slice needs to be properly oriented (flipped). One side of the brain slice is

9

Dopamine and Dendritic Calcium

135

always better than the other. The good side must face the objective lens. On the good side, the apical dendrites travel parallel to the surface of the slice, or dive into the slice at a very shallow angle. If apical dendrites are coming out of the slice surface, then large portions of the apical dendritic tree have met the razor blade and were damaged in the brain slice preparation step. Neurons with severed apical dendrites are not suitable for present experiments. 5. Ideal pipette resistance. We have empirically determined that 7 MΩ pipettes are ideal for loading layer 5 pyramidal neurons with calcium-sensitive and voltage-sensitive dyes (43). Pipette with smaller tips (pipette resistance > 7 MΩ) produce inadequate loading. Pipettes with larger tips (pipette resistance < 7 MΩ) produce too much damage, especially if such neuron was meant to be repatched (43, 46). 6. Clean pipettes. To prevent pipette clogging Borosilicate electrode glass (o.d. = 1.5, i.d. = 0.86 mm) was prewashed in boiling ethanol alcohol, rinsed in acetone and dried. Both dyefree and dye-rich solutions were filtered through nylon syringe filters, pore size 0.2 μm (Nalgene 4-mm). 7. Reduce exposure to excitation light. The brightness of calcium probes (Subheading 2.1, item 3) is poor compared to other fluorescent markers for intracellular application in modern neurobiology (e.g., Rhodamine, AlexaFluor). To make things worse, the excitation-emission spectra of calcium-sensitive dyes overlaps with the brain slice-autofluorescence; to further deteriorate image quality. Considerably better images can be obtained by loading neurons with red dyes Rhodamine or AlexaFluor594. In our experiments (Figs. 2, 3, 4, and 5) neurons were loaded with a mixture containing one calciumsensitive dye and one red dye (AlexaFluor594 or JPW3028). During dye loading, positioning and focusing onto the “target” dendrites we use a filter cube for red dyes. By inserting a neutral density filter in the epi-illumination light path we reduced epi-illumination light intensity down to 5–10% of what is normally used for calcium-imaging sweeps. We used a manually controlled shutter to keep the illumination episodes very brief (2–3 s) and very seldom. In summary four steps are regularly used to minimize the photodynamic damage from calcium-sensitive dyes prior to the beginning of calcium sensitive dye measurements. Basically, in order to position and focus fluorescently labeled neurons for calcium imaging: (a) Use excitation wavelength for AlexaFluor594. (b) Reduce epi-illumination light intensity. (c) Reduce shutter open-time to less than 3 s per opening. (d) Limit the number of positioning and focusing adjustments to less than 5 per experiment.

136

W.-L. Zhou et al.

Acknowledgments This work was supported by an R01 grant from National Institutes of Health (NIH)—grant number MH063503, and the NARSAD Young Investigator Award to S.D.A. References 1. Brozoski TJ, Brown RM, Rosvold HE, Goldman PS (1979) Cognitive deficit caused by regional depletion of dopamine in prefrontal cortex of rhesus monkey. Science 205: 929–932 2. Carlsson A (1988) The current status of the dopamine hypothesis of schizophrenia. Neuropsychopharmacology 1:179–186 3. Goldman-Rakic PS, Leranth C, Williams SM, Mons N, Geffard M (1989) Dopamine synaptic complex with pyramidal neurons in primate cerebral cortex. Proc Natl Acad Sci U S A 86:9015–9019 4. Elston GN (2003) Cortex, cognition and the cell: new insights into the pyramidal neuron and prefrontal function. Cereb Cortex 13:1124–1138 5. Milojkovic BA, Radojicic MS, Goldman-Rakic PS, Antic SD (2004) Burst generation in rat pyramidal neurones by regenerative potentials elicited in a restricted part of the basilar dendritic tree. J Physiol 558:193–211 6. Polsky A, Mel BW, Schiller J (2004) Computational subunits in thin dendrites of pyramidal cells. Nat Neurosci 7:621–627 7. Larkum ME, Nevian T, Sandler M, Polsky A, Schiller J (2009) Synaptic integration in tuft dendrites of layer 5 pyramidal neurons: a new unifying principle. Science 325:756–760 8. Bito H, Deisseroth K, Tsien RW (1997) Ca2 + -dependent regulation in neuronal gene expression. Curr Opin Neurobiol 7:419–429 9. Lisman J, Malenka RC, Nicoll RA, Malinow R (1997) Learning mechanisms: the case for CaM-KII. Science 276:2001–2002 10. Zucker RS (1999) Calcium- and activitydependent synaptic plasticity. Curr Opin Neurobiol 9:305–313 11. Lohmann C (2009) Calcium signaling and the development of specific neuronal connections. Prog Brain Res 175:443–452 12. Yu LM, Goda Y (2009) Dendritic signaling and homeostatic adaptation. Curr Opin Neurobiol 19:327–335 13. Manita S, Ross WN (2009) Synaptic activation and membrane potential changes modulate the

14.

15.

16.

17.

18.

19.

20.

21.

22.

frequency of spontaneous elementary Ca2+ release events in the dendrites of pyramidal neurons. J Neurosci 29:7833–7845 Emptage N, Bliss TV, Fine A (1999) Single synaptic events evoke NMDA receptor-mediated release of calcium from internal stores in hippocampal dendritic spines. Neuron 22:115–124 Nakamura T, Barbara JG, Nakamura K, Ross WN (1999) Synergistic release of Ca2+ from IP3-sensitive stores evoked by synaptic activation of mGluRs paired with backpropagating action potentials. Neuron 24:727–737 Hagenston AM, Fitzpatrick JS, Yeckel MF (2007) MGluR-mediated calcium waves that invade the soma regulate firing in layer V medial prefrontal cortical pyramidal neurons. Cereb Cortex 18(2):407–23 Jaffe DB, Johnston D, Lasser-Ross N, Lisman JE, Miyakawa H, Ross WN (1992) The spread of Na+ spikes determines the pattern of dendritic Ca2+ entry into hippocampal neurons. Nature 357:244–246 Markram H, Helm PJ, Sakmann B (1995) Dendritic calcium transients evoked by single back-propagating action potentials in rat neocortical pyramidal neurons. J Physiol 485:1–20 Svoboda K, Denk W, Kleinfeld D, Tank DW (1997) In vivo dendritic calcium dynamics in neocortical pyramidal neurons. Nature 385:161–165 Waters J, Larkum M, Sakmann B, Helmchen F (2003) Supralinear Ca2+ influx into dendritic tufts of layer 2/3 neocortical pyramidal neurons in vitro and in vivo. J Neurosci 23:8558–8567 Regehr WG, Tank DW (1990) Postsynaptic NMDA receptor-mediated calcium accumulation in hippocampal CA1 pyramidal cell dendrites. Nature 345:807–810 Miyakawa H, Ross WN, Jaffe D, Callaway JC, Lasser-Ross N, Lisman JE, Johnston D (1992) Synaptically activated increases in Ca2+ concentration in hippocampal CA1 pyramidal cells are primarily due to voltage-gated Ca2+ channels. Neuron 9:1163–1173

9 23. Denk W, Yuste R, Svoboda K, Tank DW (1996) Imaging calcium dynamics in dendritic spines. Curr Opin Neurobiol 6:372–378 24. Mainen ZF, Malinow R, Svoboda K (1999) Synaptic calcium transients in single spines indicate that NMDA receptors are not saturated. Nature 399:151–155 25. Higley MJ, Sabatini BL (2010) Competitive regulation of synaptic Ca2+ influx by D2 dopamine and A2A adenosine receptors. Nat Neurosci 13:958–966 26. Schiller J, Major G, Koester HJ, Schiller Y (2000) NMDA spikes in basal dendrites of cortical pyramidal neurons. Nature 404:285–289 27. Milojkovic BA, Zhou WL, Antic SD (2007) Voltage and calcium transients in basal dendrites of the rat prefrontal cortex. J Physiol 585:447–468 28. Major G, Polsky A, Denk W, Schiller J, Tank DW (2008) Spatiotemporally graded NMDA spike/plateau potentials in basal dendrites of neocortical pyramidal neurons. J Neurophysiol 99:2584–2601 29. Cox CL, Denk W, Tank DW, Svoboda K (2000) Action potentials reliably invade axonal arbors of rat neocortical neurons. Proc Natl Acad Sci U S A 97:9724–9728 30. Koester HJ, Sakmann B (2000) Calcium dynamics associated with action potentials in single nerve terminals of pyramidal cells in layer 2/3 of the young rat neocortex. J Physiol 3:625–646 31. Wachowiak M, Cohen LB (2001) Representation of odorants by receptor neuron input to the mouse olfactory bulb. Neuron 32:723–735 32. Brown JE, Cohen LB, De Weer P, Pinto LH, Ross WN, Salzberg BM (1975) Rapid changes in intracellular free calcium concentration. Detection by metallochromic indicator dyes in squid giant axon. Biophys J 15:1155–1160 33. Ross WN, Arechiga H, Nicholls JG (1987) Optical recording of calcium and voltage transients following impulses in cell bodies and processes of identified leech neurons in culture. J Neurosci 7:3877–3887 34. Yuste R, Gutnick MJ, Saar D, Delaney KR, Tank DW (1994) Ca2+ accumulations in dendrites of neocortical pyramidal neurons: an apical band and evidence for two functional compartments. Neuron 13:23–43 35. Yasuda R, Nimchinsky EA, Scheuss V, Pologruto TA, Oertner TG, Sabatini BL, Svoboda K (2004) Imaging calcium concentration dynamics in small neuronal compartments. Sci STKE 2004(219):l5 36. Homma R, Baker BJ, Jin L, Garaschuk O, Konnerth A, Cohen LB, Zecevic D (2009)

37.

38.

39.

40.

41.

42. 43.

44.

45.

46.

47.

48.

Dopamine and Dendritic Calcium

137

Wide-field and two-photon imaging of brain activity with voltage- and calcium-sensitive dyes. Philos Trans R Soc Lond B Biol Sci 364:2453–2467 Lidow MS, Goldman-Rakic PS, Gallager DW, Rakic P (1991) Distribution of dopaminergic receptors in the primate cerebral cortex: quantitative autoradiographic analysis using [3H] raclopride, [3H]spiperone and [3H] SCH23390. Neuroscience 40:657–671 Goldman-Rakic PS, Muly EC 3rd, Williams GV (2000) D(1) receptors in prefrontal cells and circuits. Brain Res Brain Res Rev 31:295–301 Westenbroek RE, Hell JW, Warner C, Dubel SJ, Snutch TP, Catterall WA (1992) Biochemical properties and subcellular distribution of an N-type calcium channel alpha 1 subunit. Neuron 9:1099–1115 Kisilevsky AE, Mulligan SJ, Altier C, Iftinca MC, Varela D, Tai C, Chen L, Hameed S, Hamid J, Macvicar BA, Zamponi GW (2008) D1 receptors physically interact with N-type calcium channels to regulate channel distribution and dendritic calcium entry. Neuron 58:557–570 Gulledge AT, Stuart GJ (2003) Action potential initiation and propagation in layer 5 pyramidal neurons of the rat prefrontal cortex: absence of dopamine modulation. J Neurosci 23:11363–11372 Schultz W (2002) Getting formal with dopamine and reward. Neuron 36:241–263 Antic SD (2003) Action potentials in basal and oblique dendrites of rat neocortical pyramidal neurons. J Physiol 550:35–50 Milojkovic BA, Wuskell JP, Loew LM, Antic SD (2005) Initiation of sodium spikelets in basal dendrites of neocortical pyramidal neurons. J Membr Biol 208:155–169 Foust A, Popovic M, Zecevic D, McCormick DA (2010) Action potentials initiate in the axon initial segment and propagate through axon collaterals reliably in cerebellar Purkinje neurons. J Neurosci 30:6891–6902 Antic S, Major G, Zecevic D (1999) Fast optical recordings of membrane potential changes from dendrites of pyramidal neurons. J Neurophysiol 82:1615–1621 Zhou WL, Yan P, Wuskell JP, Loew LM, Antic SD (2008) Dynamics of action potential backpropagation in basal dendrites of prefrontal cortical pyramidal neurons. Eur J Neurosci 27(4):923–936 Canepari M, Popovic M, Vogt K, Holthoff K, Konnerth A, Salzberg BM, Grinvald A, Antic SD, Zecevic D (2010) Imaging submillisecond membrane potential changes from individual

138

49.

50.

51.

52.

53.

54.

55.

W.-L. Zhou et al. regions of single axons, dendrites and spines. In: Canepari M, Zecevic D (eds) Membrane potential imaging in the nervous system: methods and applications. Springer Science+Business Media, LLC, New York Stuart G, Schiller J, Sakmann B (1997) Action potential initiation and propagation in rat neocortical pyramidal neurons. J Physiol 505:617–632 Spruston N (2008) Pyramidal neurons: dendritic structure and synaptic integration. Nat Rev Neurosci 9:206–221 Vetter P, Roth A, Hausser M (2001) Propagation of action potentials in dendrites depends on dendritic morphology. J Neurophysiol 85:926–937 Frick A, Magee J, Johnston D (2004) LTP is accompanied by an enhanced local excitability of pyramidal neuron dendrites. Nat Neurosci 7:126–135 Tsubokawa H, Ross WN (1997) Muscarinic modulation of spike backpropagation in the apical dendrites of hippocampal CA1 pyramidal neurons. J Neurosci 17:5782–5791 Hoffman DA, Johnston D (1999) Neuromodulation of dendritic action potentials. J Neurophysiol 81:408–411 Djurisic M, Antic S, Chen WR, Zecevic D (2004) Voltage imaging from dendrites of mitral cells: EPSP attenuation and spike trigger zones. J Neurosci 24:6703–6714

56. Milojkovic BA, Radojicic MS, Antic SD (2005) A strict correlation between dendritic and somatic plateau depolarizations in the rat prefrontal cortex pyramidal neurons. J Neurosci 25:3940–3951 57. Antic SD, Acker CD, Zhou WL, Moore AR, Milojkovic BA (2007) The role of dendrites in the maintenance of the UP state. In: Timofeev I (ed) Mechanisms of spontaneous active states in the neocortex. Research Signpost, Kerala, India, pp 45–72 58. Acker CD, Antic SD (2009) Quantitative assessment of the distributions of membrane conductances involved in action potential backpropagation along basal dendrites. J Neurophysiol 101:1524–1541 59. Antic SD, Zhou WL, Moore AR, Short SM, Ikonomu KD (2010) The decade of the dendritic NMDA spike. J Neurosci Res 88:2991–3001 60. Antic SD, Acker CD, Zhou WL, Moore AR (2008) Dopaminergic modulation of dendritic excitability in neocortical pyramidal neurons. Cell Science Reviews 5:1742–8130 61. Nevian T, Sakmann B (2004) Single spine Ca2+ signals evoked by coincident EPSPs and backpropagating action potentials in spiny stellate cells of layer 4 in the juvenile rat somatosensory barrel cortex. J Neurosci 24:1689–1699

Part III Genetic Manipulation in Cells and Organisms

Chapter 10 Functional Analysis of Human D1 and D5 Dopaminergic G Protein-Coupled Receptors: Lessons from Mutagenesis of a Conserved Serine Residue in the Cytosolic End of Transmembrane Region 6 Bianca Plouffe and Mario Tiberi Abstract In mammals, dopamine G protein-coupled receptors (GPCR) are segregated into two categories: D1-like (D1R and D5R) and D2-like (D2Rshort, D2Rlong, D3R, and D4R) subtypes. D1R and D5R are primarily coupled to stimulatory heterotrimeric GTP-binding proteins (Gs/olf) leading to activation of adenylyl cyclase and production of intracellular cAMP. D1R and D5R share high level of amino acid identity in transmembrane (TM) regions. Yet these two GPCR subtypes display distinct ligand binding and G protein coupling properties. In fact, our studies suggest that functional properties reported for constitutively active mutants of GPCRs (e.g., increased basal activity, higher agonist affinity and intrinsic activity) are also observed in cells expressing wild type D5R when compared with wild type D1R. Herein, we describe an experimental method based on mutagenesis and transfection of human embryonic kidney 293 (HEK293) cells to explore the molecular mechanisms regulating ligand affinity, agonist-independent and dependent activity of D1R and D5R. We will demonstrate how to mutate one conserved residue in the cytosolic end of TM6 of D1R (Ser263) and D5R (Ser287) by modifying two or three nucleotides in the cDNA of human D1-like receptors. Genetically modified D1R and D5R cDNAs are prepared using a polymerase chain reaction method, propagated in E. coli, purified and mutations confirmed by DNA sequencing. Receptor expression constructs are transfected into HEK293 cells cultured in vitro at 37°C in 5% CO2 environment and used in radioligand binding and whole cAMP assays. In this study, we will test the effect of S263A/ G/D and S287A/G/D mutations on ligand binding and DA-dependent activation of D1R and D5R. Key words: Dopamine, GPCR, D1-like receptors, Ligand binding, cAMP, Mutagenesis, Third intracellular loop, TM6, HEK293 cells

1. Introduction Transmembrane (TM) signaling is fundamental in the homeostatic regulation of every major physiological function in eukaryotes ranging from yeast to human (1, 2). A class of integral membrane Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_10, © Springer Science+Business Media, LLC 2013

141

142

B. Plouffe and M. Tiberi

proteins known as G protein-coupled receptors (GPCRs), which harbor seven a-helical TM segments, best illustrates this notion (3). Indeed, over 800 types of GPCRs expressed throughout the body in humans are stimulated by a variety of ligands such as photons, protons, ions, odorants, lipids, biogenic amines, peptides and hormones (3). Based on primary sequence and structural similarities, GPCRs can be grouped into five families: rhodopsin (family A), secretin (family B), glutamate (family C), adhesion and Frizzled/ Taste2 (3, 4). Classically, heterotrimeric guanine-nucleotide-binding proteins (G proteins) composed of a, b, and g subunits and located at the cytoplasmic side of plasma membrane, serve as molecular switches in GPCR signaling pathways by coupling ligandinduced receptor stimulation to intracellular responses. This is essentially accomplished by receptor activation of G proteins through the catalysis of GTP for GDP exchange on Ga promoting a conformational change in GTP-bound Ga and Gbg subunits, which culminates in G protein subunit-mediated regulation of activity of different downstream effector proteins (3). Meanwhile, studies also show that GPCR signaling can be mediated in a G protein-independent manner (5, 6). Importantly, at least 50% of clinical drugs target GPCRs, hence highlighting their human therapeutic relevance (7, 8). Given the physiological and clinical importance of these integral membrane proteins, it is crucial to know how the conserved seven-a-helix bundle structure imparts subtypespecific ligand binding and activation properties to the GPCR members. Difference in the extent of constitutive activity naturally displayed by homologous GPCR subtypes was first highlighted with D1-like dopaminergic receptors (D1R and D5R), which belong to the family A GPCRs (9). In contrast to inhibitory G protein (Gi)linked D2-like subtypes (D2Rshort, D2Rlong, D3R, D4R), D1R and D5R couple to stimulatory G proteins (Gs/olf) leading to adenylyl cyclase (AC) activation and production of intracellular cAMP (10, 11). Interestingly, D5R naturally has a greater constitutive activity (i.e., increased ability to produce intracellular cAMP in the absence of agonists) when compared with D1R at similar receptor levels (9). Furthermore, pharmacological properties of ligands displayed at D5R are highly reminiscent of those described for constitutively active mutant (CAM) GPCRs (e.g., higher agonist affinity and intrinsic activity) (9, 12). The molecular and structural basis underlying D1-like subtype-specific functional properties has been difficult to address experimentally as TM regions of D1R and D5R are almost indistinguishable (>80% identity) (13). Indeed, the higher dopamine affinity of D5R relative to D1R cannot be explained by TM domains interacting with dopamine as critical amino acids implicated in catecholamine binding (e.g., TM5 serine residues) are conserved among dopaminergic and adrenergic receptors. Likewise, motifs found within cytosolic surfaces of TM

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

143

domains (e.g., “ionic lock” between TM3 and TM6) regulating receptor activation are also conserved among family A GPCRs (3). This view is further underscored by data obtained from recent crystallographic studies of inactive states of different family A GPCRs, which display a highly conserved 7TM structural design (3). Therefore, functional differences between GPCRs displaying high degree of TM identity likely implicate subtle variations in the assembly of conserved TM amino acids. These subtle variations are potentially mediated by GPCR subtype-specific conformation of remote residues such as those found in receptor intracellular regions. While amino acids located in receptor intracellular regions are unlikely to be directly involved in the binding to extracellular ligands, mutagenesis studies suggest that the third intracellular loop (IL3) and cytoplasmic tail (CT) play a major role in mediating D1-like subtype-specific constitutive activity, agonist affinity and intrinsic activity (14–18). Notably, substitution of two variant residues located in the cytosolic end of TM6 of D1R (F264 and R266) and D5R (I288 and K290) using site-directed mutagenesis show that F264 and I288 are major determinants in conferring D1-like subtype-specific agonist binding, constitutive activity and agonistdependent G protein-coupling properties (14). Interestingly, F264 (D1R) and I288 (D5R) are located three amino acids downstream of the glutamate residue, which participates in the formation of an ionic lock with the highly conserved E/DRY motif of TM3. These D1R and D5R residues are not found in other catecholaminergic receptors and hence play potentially an important role in modulating the arrangement of conserved amino acids forming the ionic lock. Herein, we describe an experimental approach to study the role of a conserved serine residue located adjacent to F264 (S263 in D1R) and I288 (S287 in D5R) (Fig. 1), which at this position does not exist in other catecholamine receptors. We test whether this conserved serine residue plays a critical role in the modulation of D1-like receptor binding and activation properties.

2. Materials 2.1. Molecular Biology Reagents

1. Custom DNA oligonucleotides (Sigma Genosys, Burlington, ON, Canada) as listed in Tables 1 and 2. Lyophilized oligonucleotides are prepared as stock solutions in sterile Milli-Q water (resistivity of 18.2 MW cm) at 25 pmol/mL for PCR primers or 2 pmol/mL for DNA sequencing primer. 2. Expand High Fidelity Taq DNA polymerase (3.5 U/mL), PCR buffer (10×), and MgCl2 (25 mM) (Roche Diagnostics, Laval, QC, Canada). Store at −20°C.

144

B. Plouffe and M. Tiberi

Fig. 1. Schematic representation of wild type hD1R and hD5R. Putative secondary structure of wild type hD1R and hD5R are indicated by circles. Distinct residues between hD1R and hD5R are depicted using black circles. Amino acid sequence of the region encompassing TM5, IL3, and TM6 is also shown. The mutated serine in IL3 of hD1R and hD5R is also indicated. hD1R human D1 receptor, hD5R human D5 receptor.

3. PCR Nucleotides (dATP, dCTP, dGTP, and dTTP) at 100 mM (Fermentas, Burlington, ON, Canada). Store at −20°C. 4. dNTP Mix: Add 10 mL dATP, 10 mL dCTP, 10 mL dGTP, and 10 mL dTTP to 60 mL sterile Milli-Q water. Store at −20°C. 5. BoxI (PshAI), BsmI (Mva1269I), DraI, Eag I (Eco52I), EcoRI, HindIII, and XbaI restriction enzymes at 10 U/mL with 10× Buffers (Y+/TangoTM or yellow; B+ or blue; R+ or red) from Fermentas. Store at −20°C. 6. Calf Intestinal Alkaline Phosphatase (CIAP) at 1 U/mL and 10× dephosphorylation buffer from Fermentas. Store at −20°C. 7. T4 DNA ligase (5 U/mL) and T4 DNA ligase buffer (10×) from Fermentas. Store at −20°C. 8. Tris–acetate–EDTA (TAE) buffer (50×): 2 M Tris–HCl, pH 8.0, 5.71% (v/v) glacial acetic acid, 0.5 M ethylenediamine tetraacetic acid (EDTA), pH 8.0 in Milli-Q water. Store at room temperature.

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

145

Table 1 Sequences of oligonucleotide primers for the construction of single-point mutations of hD1R using a PCR-based overlapping approach Construct

Primer sequence 5¢→3¢

hD1R-S263A

P1: TTcATcccAgTgcAgcTc P2: TTcTcTTTTgAATgCcATcTTAAAAgAAcTTTccgg P3: TTTAAgATgGcATTcAAAAgAgAAAcTAAAgTccTg P4: TTAggAcAAggcTggTgg P5: AcTgTgAcTccAgcc P6: ggccAggAgAggcA

hD1R-S263D

P1: TTcATcccAgTgcAgcTc P2: TTcTcTTTTAAAgTCcATcTTAAAAgAAcTTTccgg P3: TTTAAgATgGAcTTTAAAAgAgAAAcTAAAgTccTg P4: TTAggAcAAggcTggTgg P5: AcTgTgAcTccAgcc P6: ggccAggAgAggcA

hD1R-S263G

P1: TTcATcccAgTgcAgcTc P2: TTcTcTTTTAAAgCCcATcTTAAAAgAAcTTTccgg P3: TTTAAgATgGGcTTTAAAAgAgAAAcTAAAgTccTg P4: TTAggAcAAggcTggTgg P5: AcTgTgAcTccAgcc P6: ggccAggAgAggcA

For each single-point mutant, a set of six primers numbered P1–P6 is used in PCR reactions. Nucleotide changes leading to S263A, S263D and S263G mutations are indicated in bold. Nucleotide changes introducing diagnostic restriction sites through silent mutations are underlined or through the S263A mutation are underlined and bold

9. Agarose gels (1% w/v): Weigh out 0.5 g of agarose (SigmaAldrich, Oakville, ON, Canada) into a plastic 250 mL capped conical flask and add 50 mL of 1× TAE. Microwave for ~60 s with loosened cap. Gently swirl and microwave it again for 10 s and repeat 2–3 times if agarose is not completely dissolved. Keep watching agarose solution during microwaving as it can easily boil over. As the solution can become very hot, experimenter should wear gloves and hold flask at arms length. Once agarose is completely dissolved let it cool down briefly (~5 min), add 2.5 mL of ethidium bromide (10 mg/mL) and gently swirl. Gloves should be worn when handling ethidium bromide solution as it is mutagenic and to some extent toxic. Slowly pour the agarose solution into a mini plastic casting tray, insert sample comb, push away any bubbles to the side using a pipet tip and allowed gel to solidify for 30–60 min at room temperature. Rinse out the flask (see Note 1). 10. 50% (v/v) Glycerol. Sterilize through 0.22 mm filter and store at room temperature.

146

B. Plouffe and M. Tiberi

Table 2 Sequences of oligonucleotide primers for the construction of single-point mutations of hD5R using a PCR-based overlapping approach Construct

Primer sequence 5¢→3¢

hD5-S287A

P1: TAcggTgggAgg P2: gATggCAgcgcgcAgAcTggTgTcgggcgcgcAggc P3: gcgcccgAcAccAgTcTgcgcgcTGccATcAAgAAg P4: TcATgTggATgTAggcAg P5: AccTggccAAcTggA P6: TgTTcAccgTcTccA

hD5-S287D

P1: TAcggTgggAgg P2: gATgTCAgcgcgcAgAcTggTgTcgggcgcgcAggc P3: gcgcccgAcAccAgTcTgcgcgcTGAcATcAAgAAg P4: TcATgTggATgTAggcAg P5: AccTggccAAcTggA P6: TgTTcAccgTcTccA

hD5-S287G

P1: TAcggTgggAgg P2: gATgCCAgcgcgcAgAcTggTgTcgggcgcgcAggC P3: gcgcccgAcAccAgTcTgcgcgcTGGcATcAAgAAg P4: TcATgTggATgTAggcAg P5: AccTggccAAcTggA P6: TgTTcAccgTcTccA

For each single-point mutant, a set of six primers numbered P1–P6 is used in PCR reactions. For each single-point mutant, a set of six primers numbered P1-P6 is used in PCR reactions. Nucleotide changes leading to S287A, S287D and S287G mutations are indicated in bold. Nucleotide changes introducing diagnostic restriction sites through silent mutations are underlined

11. Loading Dye (10×): 0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cyanole FF (optional), 50% (v/v) glycerol in Milli-Q water. Store in aliquots at −20°C. 12. DNA Size Markers: Prepare 1× stock with 250 mL of One Kilobase Plus DNA Ladder (1 mg/mL; Invitrogen, Burlington, ON, Canada), 250 mL of 10× loading dye, and 2,000 mL Milli-Q water. Store in aliquots at −20°C. 13. Sterile Luria Broth (LB) Medium: Weigh out 12.5 g of LB base (Invitrogen) into glass bottle or Fernbach flask (for DNA maxipreps) and add 500 mL of Milli-Q water. Sterilize liquid media using autoclave. Prior to autoclaving loosen cap on glass bottle and tape aluminum foil on top of flask. Always wear heat protective gloves when handling bottle or flask at the end of autoclave cycle. Store at room temperature. If store for a long time period, check that there is no microorganism contamination prior to use.

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

147

14. Ampicillin (1,000×): 1 g of ampicillin (Sigma-Aldrich) is dissolved in Milli-Q water at 100 mg/mL and sterilized using a syringe filter (0.22 mm). Store in 0.5 mL aliquots at −20°C. 15. LB Ampicillin Plates: Weigh out 25 g of LB base (Invitrogen), 15 g of Bacto Agar (Fisher Scientific, Ottawa, ON, Canada) into glass flask (for DNA maxipreps) and add 1 L of Milli-Q water. Sterilize using autoclave. Always wear heat protective gloves when handling flask at the end of autoclave cycle. Let it cool down for 15 min at room temperature. Thaw two 0.5 mL aliquots of ampicillin (1,000×), add to molten LB-agar, swirl, and pour into sterile polystyrene petri dishes (100 × 15 mm, Fisher Scientific). Quickly pass over the petri dishes a flame from a Bunsen burner to remove bubbles. Keep moving flame while removing bubbles to avoid overheating LB-agar and break up ampicillin. 1 L of LB-agar will make 50 plates. Store inverted in plastic sleeves at 4°C for no longer than 3 months. Prior to using plates verify that there is no microorganism contamination. 16. SOB Medium: 2% (w/v) bacto-tryptone (Fisher Scientific), 0.5% (w/v) bacto-yeast extract (Fisher Scientific), and 10 mM NaCl. Sterilize by autoclaving. Store at room temperature. Prior to using SOB medium verify that there is no microorganism contamination. 17. SOC Medium: SOB medium containing 10 mM MgCl2, 10 mM MgSO4, and 20 mM glucose. Solutions of 1 M MgCl2, 1 M MgSO4, and 2 M glucose are separately made and sterilize by filtration using 0.22 mm filter. Store at room temperature. 18. XL1-Blue Electroporation-Competent Cells (Agilent Technologies, Mississauga, ON, Canada). Store at −80°C. 19. Sterile Polypropylene Capped 13 mL Tubes (100 × 16 mm; Sarstedt, St-Léonard, QC, Canada). 20. Isobutanol (Fisher Scientific). 21. Qiaex II Gel Extraction Kit (Buffer QX1, Buffer PE, Qiaex II Bead Suspension) (Qiagen, Mississauga, ON, Canada). 22. QIAprep Spin Miniprep and Plasmid Maxiprep Kits from Qiagen. 2.2. Cell Culture

1. Adenovirus type 5-transformed human embryonic kidney 293 (HEK293) cells (CRL-1573, American Tissue Type Culture Collection, Manassas, VA). 2. Minimal Essential (Invitrogen).

Medium

(MEM)

with

Earle’s

salt

3. Fetal Bovine Serum (FBS) (Invitrogen). Thaw frozen FBS bottle at 4°C overnight. The next day, warm up bottle in a 37°C water bath, heat-inactivate FBS in a 55°C water bath for

148

B. Plouffe and M. Tiberi

1 h, and sterilize through 0.22 mm filter. Store in 25 and 50 mL aliquots in sterile capped polypropylene conical tubes at −20°C. 4. Gentamicin Sulfate (10 mg/mL) (Invitrogen). 5. Trypsin (0.25%) and EDTA (0.05% (w/v)) Buffer Solution (Invitrogen). 6. Ca2+ and Mg2+-free Phosphate-Buffered Saline (PBS) (Wisent, St-Bruno, QC, Canada). 7. Tissue Culture Grade Sigma Hybri-Max Dimethylsulfoxide (DMSO) (Sigma-Aldrich). Store at room temperature. 8. BD Falcon Polystyrene 75 cm2 Flasks with 0.2 mm Vented Blue Plug Seal Cap (VWR International, Montréal, QC, Canada). 9. Sterile HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) Buffer Solution (1 M, pH 7.4) (Invitrogen). 10. 20 mM HEPES-buffered MEM: In a certified biological safety cabinet (BSC), add to bottle of MEM (500 mL), 10 mL of sterile HEPES (pH 7.4) and 0.5 mL of gentamicin (SigmaAldrich). Store at 4°C for up to 2 months. 11. 1 M HEPES (pH 7.0): 37.75 g of HEPES (Fisher Scientific) is dissolved in 100 mL Milli-Q water, adjust pH to 7.0 and complete to 150 mL with Milli-Q water. Sterilize through 0.22 mm filter. Store in tissue culture area at room temperature (see Note 2). 12. 2 M NaCl: Prepare in glass bottle containing 500 mL of Milli-Q water and sterilize by autoclaving. Store in tissue culture area at room temperature. 13. 1 M Na2HPO4: Add 35.5 g of Na2HPO4 in a beaker containing 230 mL of Milli-Q water, stir on a hot plate at low temperature (setting 2) until completely dissolved, complete to a final volume of 250 mL, and sterilize through 0.22 mm filter. Store in tissue culture area at room temperature. 14. 1 M NaH2PO4: Add 34.5 g of NaH2PO4 in a beaker containing 230 mL of Milli-Q water, stir on a hot plate at low temperature (setting 2) until completely dissolved, complete to a final volume of 250 mL, and sterilize through 0.22 mm filter. Store in tissue culture area at room temperature. 15. 1 M Na3PO4: Prepare 25 mL by adding equal volumes of sterile 1 M Na2HPO4 and 1 M NaH2PO4 solutions in a BSC. Store in tissue culture area at room temperature (see Note 3). 16. Sterile 2× HEPES-buffered saline (0.28 M NaCl, 0.05 M HEPES, and 1.5 mM Na3PO4, pH 7.1): Add 70 mL of 2 M NaCl, 25 mL of 1 M HEPES, and 750 mL of 1 M Na3PO4 to 400 mL of Milli-Q water in a glass beaker. Stir and adjust to pH 7.1 (±0.05). Complete to a final volume of 500 mL and

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

149

sterilize through 0.22 mm filter in a BSC. Store in 15 mL aliquots at −20°C. 17. Teflon cell lifters (Fisher Scientific). 2.3. Radioligand Binding and Whole Cell cAMP Assays

1. [3H]-SCH23390 and [3H]-adenine (PerkinElmer NEN, Boston, MA). Store at −20°C (see Note 4). 2. Ascorbic acid, (+)-SCH23390 hydrochloride, dopamine hydrochloride, cis-flupenthixol dihydrochloride and (+)-butaclamol hydrochloride (Sigma-Aldrich). Store at room temperature (see Note 5). 3. Lysis Buffer: 10 mM Tris–HCl, pH 7.4 and 5 mM EDTA, pH 8.0. Store at 4°C. 4. Resuspension Buffer: 62.5 mM Tris–HCl, pH 7.4 and 1.25 mM EDTA, pH 8.0. Store at 4°C. 5. Binding Buffer: 62.5 mM Tris–HCl, pH 7.4 and 1.25 mM EDTA, pH 8.0, 200 mM NaCl, 6.7 mM MgCl2, 2.5 mM CaCl2 and 8.33 mM KCl. Store at 4°C. 6. Washing Buffer (10×): 500 mM Tris–HCl, pH 7.4 and 1 M NaCl. Store at 4°C. 7. Whatman GF-C Glassfiber Filter Sheets (Brandel Inc., Gaithersburg, MD). 8. Plastic Scintillation Vials (20 mL) (Sarstedt, Newton, NC). 9. Bio-Safe II Biodegradable Scintillation Cocktail (Research Products International Corp., Mount Prospect, IL). 10. Bio-Rad Protein Assay Dye Concentrate (Bio-Rad Laboratories Inc., Mississauga, ON, Canada). Store at 4°C. 11. Bovine Serum Albumin (BSA), minimum 96% electrophoresis (Sigma-Aldrich). BSA is dissolved in sterile Milli-Q water at 1 mg/mL. Store in 1 mL aliquots at −20°C. 12. 3-isobutyl-1-methylxanthine (IBMX) (Sigma-Aldrich) stored −20°C is dissolved in DMSO at 200 mM. Store stock solution at 4°C (see Note 6). 13. Neutralizing Solution: 4.2 M KOH. Store in a glass bottle at room temperature. 14. Cyclic AMP (cAMP) (Sigma-Aldrich). Store desiccated at −20°C. 15. [14C]-cAMP (Moravek Biochemicals, Brea, CA). Store at −20°C. 16. cAMP Stop Solution: 2.5% (v/v) perchloric acid, 0.1 mM cAMP and [14C]-cAMP (~3.3 nCi/mL; ~10,000 dpm) in 1,500 mL Milli-Q water. Store at 4°C (see Note 7). 17. Dowex AG 50 W-4X Resin (hydrogen form, 200–400 dry mesh, 63–150 mm wet beads) from Bio-Rad Laboratories (Hercules, CA).

150

B. Plouffe and M. Tiberi

18. Alumina N Super I (MP Biomedicals, Montréal, QC, Canada). Store at room temperature. 19. Poly-Prep Columns (with 10 mL reservoir and graduated volume markings) from Bio-Rad Laboratories Inc. 20. Hydrochloric Acid (HCl) and Sodium Hydroxide (NaOH) at 0.1 N. 21. Imidazole (Sigma-Aldrich) is dissolved in Milli-Q water at 2 M, pH 7.5 (see Note 8). Store in plastic bottle at room temperature for up to 4 months.

3. Methods The following experimental strategy describes how to generate three single-point mutants of S263 of hD1R (S263A, S263D and S263G) and S287 of hD5R (S287A, S287D, S287G) using a PCRbased overlap extension approach, DNA ligation, and DNA automated sequencing (Fig. 2). The experimental approach entails two major steps: 1. DNA primers derived from forward and reverse nucleotide sequences coding for human D1R (hD1R) and D5R (hD5R) are designed to mutate the candidate serine into alanine, aspartate, and glycine. The hD1R and hD5R DNA templates are separately mixed with mutagenesis primers and a high proofreading DNA polymerase to produce “megaprimers” using polymerase chain reaction (PCR) and an overlap extension approach. Pairs of purified overlapping “megaprimers” are subsequently fused together by PCR and purified by silica-gel particles to obtain the different mutated receptor DNA cassettes. Mutated receptor DNA cassettes and corresponding wild type receptor DNA in the pCMV5 expression plasmid (cloning vector) are subjected to restriction digest and ligation procedures to generate different mammalian mutant receptor expression constructs. 2. Human embryonic kidney 293 (HEK293) cells are transfected with wild type and mutant receptor expression constructs using a DNA-calcium phosphate precipitation procedure, and seeded in tissue culture dishes and plates for radioligand binding and whole cell cAMP studies. Ligand binding properties of wild type and mutant receptors are measured in membrane preparations of transfected cells with saturation and competition experiments using the D1-like selective radioligand [3H]-SCH23390. Agonist-independent and dependent coupling to Gs is assessed in transfected cells metabolically labeled

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

151

Fig. 2. Schematic representation of the making of pCMV5 expression constructs for single-point mutant forms of hD1R and hD5R. The key steps involved in the preparation of hD1R (a) and hD5R (b) single-point mutant constructs in the pCMV5 expression vector are shown (see text for details).

with [3H]-adenine and the amount of intracellular cAMP production determined from cell lysates with a sequential chromatography purification procedure using Dowex and alumina columns. Radioligand binding and whole cell cAMP data are analyzed using nonlinear curve fitting program and statistical tests. Our results suggest that S263 in hD1R and S287 in hD5R play a differential role in controlling ligand affinity and DA-induced activation of AC of human D1-like receptors (see data figures).

152

B. Plouffe and M. Tiberi

3.1. Preparation of Single-Point Mutant hD1R and hD5R Cassettes by SiteDirected Mutagenesis and PCR

1. Qiagen maxipreps of wild type hD1R and hD5R subcloned in the expression vector pCMV5 are used as DNA templates (25 ng/mL) in series of two PCR rounds as depicted in Fig. 3. For each single-point mutant two PCR products (“megaprimers”), A and B, are separately amplified in the first round using specific set of P1–P2 and P3–P4 primer pairs (see Tables 1 and 2). First-round PCRs are carried out in a final volume of 50 mL containing 50 ng of DNA template (2 mL of stock), 50 pmol of forward primer (2 mL of P1 or P3 stock solution at 25 pmol/mL), 50 pmol of reverse primer (2 mL of P2 or P4 stock solution at 25 pmol/mL), 1.5 mM MgCl2 (3 mL of 25 mM stock solution), 0.2 mM dNTPs (1 mL of 10 mM stock solution), 3.5 U of Taq DNA polymerase (1 mL of stock

Fig. 3. General scheme for creating single-point mutations using PCR-based overlapping approach. Representative example of the experimental strategy used to generate the S263G and S287G mutations in hD1R-pCMV5 (a) and hD5R-pCMV5 (b) constructs is depicted. This strategy remains identical for creating other single point mutations in hD1R (S263A and S263D) and hD5R (S287A and S287D). The beginning of the polylinker region of pCMV5 (EcoRI site) is arbitrarily set to position 0. Nucleotide position of 5¢ region of PCR primers (P1-P6) annealing to DNA expression construct template is indicated. Restriction sites used for generating mutated cassette are shown. The boundaries of mutated cassettes are illustrated using brackets.

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

153

solution), 5 mL of 10× PCR buffer and of 34 mL of sterile Milli-Q water. DNA is amplified in an Eppendorf Thermal Mastercycler using the following conditions: 1 cycle (94°C for 3 min, 50°C for 1 min, 72°C for 3 min), 24 cycles (94°C for 45 s, 50°C for 1 min, 72°C for 1 min) completed by an anneal extension step (50°C for 1 min and 72°C for 8 min). Importantly, during this first PCR round, an overlapping region between A and B will be generated, which will allow amplifying the final PCR product using primers P5–P6 during the second PCR round (see Subheading 3.1.5). At the end of the final cycle, add 5.5 mL of 10× loading dye to PCR tubes. 2. Prepare 1× TAE buffer in Milli-Q water and pour a 1% (w/v) agarose minigel (see Subheading 2.1, item 8). Once agarose solidifies, put casting tray in electrophoresis apparatus, fill with 1× TAE buffer, gently remove sample comb, and load wells with PCR samples (55 mL) and one well with 4 mL of DNA size markers. Separate samples for 1 h at 80 V and visualize ethidium bromide-stained DNA bands using an UV Transilluminator equipped with a digital camera. 3. Cut off appropriate sized DNA bands (see Fig. 3) and purify agarose-embedded DNA with QIAEX beads (Qiagen) according to manufacturer’s protocol. Elute purified bands from QIAEX beads using 20 mL of sterile Milli-Q water. Add 2 mL of purified bands to 7 mL of sterile Milli-Q water, mix with 1 mL of 10× loading dye and run samples beside a well loaded with 4 mL of DNA size markers on 1% (w/v) agarose minigel for 1 h at 80 V. 4. Visualize ethidium bromide-stained DNA bands using an UV Transilluminator equipped with a digital camera and take picture to assist in the semi-quantification of purified products. 5. For the second-round PCRs, add equal volume of purified “megaprimer” A and B (~0.5 mL each) corresponding to the designated mutant receptor to a mix containing 19.5 mL of sterile Milli-Q water, 2.5 mL of 10× PCR buffer, 1.5 mM MgCl2 (1.5 mL of 25 mM stock solution), 0.2 mM dNTPs (0.5 mL of 10 mM stock solution), and 1.75 U of Taq DNA polymerase (0.5 mL of stock solution). The overlap PCR is done in a final volume of 25 mL using 1 cycle at 94°C for 3 min, 50°C for 1 min, 72°C for 10 min, and 20°C for 8 min. 6. During the 8-min period, 25 mL of a mix containing 19.5 mL of sterile Milli-Q water, 2.5 mL of 10× PCR buffer, 25 pmol of P5 forward primer (1 mL of 25 pmol/mL stock solution), 25 pmol of P6 reverse primer (1 mL of 25 pmol/mL stock solution), 1.5 mM MgCl2 (1.5 mL of 25 mM stock solution), 0.2 mM dNTPs (0.5 mL of 10 mM stock solution), and 1.75

154

B. Plouffe and M. Tiberi

U of Taq DNA polymerase (0.5 mL of stock solution) is added to overlap PCR mixture (final volume of 50 mL). PCR is then run for 25 cycles (94°C for 45 s, 50°C for 1 min, 72°C for 1 min) and completed by an anneal extension step (50°C for 1 min and 72°C for 8 min). At the end of this step, add 5.5 mL of 10× loading dye to PCR tubes. 7. Run final PCR products on 1% (w/v) agarose and appropriate mutant receptor DNA cassettes purified as described (see Subheadings 3.1.2, 3.1.3, and 3.1.4). 3.2. Preparation of Linearized Wild Type Receptor Expression Constructs and Mutated Receptor DNA Cassettes by Digestions with Restriction Enzymes

1. Set up restriction enzyme digestions of the wild type hD1RpCMV5 expression construct and purified mutated D1R DNA cassettes (849 bp) with HindIII and XbaI (see Fig. 3a), and wild type hD5R-pCMV5 expression construct and purified mutated D5R DNA cassettes (569 bp) with BsmI and EagI (see Fig. 3b) in a final volume of 30 mL using separate autoclaved 1.5 mL Eppendorf tubes. Prepare reaction tubes for wild type hD1R and hD5R-pCMV5 expression constructs, in which restriction enzymes are replaced with an equivalent amount of sterile Milli-Q water. These tubes are referred to as uncut DNA (see Note 9). 2. For hD1R DNA digestions, add to tubes 8 mL of wild type hD1R-pCMV5 (0.125 mg/mL working solution; 1 mg total) or purified mutant hD1R cassette (see Note 10), 16 mL of sterile Milli-Q water, 1.5 mL of HindIII (15 U), 1.5 mL of XbaI (15 U), and 3 mL of 10× Y+/TangoTM buffer (1× final) (see Note 10). 3. For hD5R DNA digestions, add to tubes 8 mL of wild type hD5R-pCMV5 (0.125 mg/mL working solution; 1 mg total) or purified mutant hD5R cassette, 13 mL of sterile Milli-Q water, 1.5 mL of BsmI (15 U), 1.5 mL of EagI (15 U), and 6 mL of 10× Y+/TangoTM buffer (2× final) (see Note 11). 4. Gently pipette up and down to mix and float tubes in a 37°C water bath for 1 h. At the end of incubation, put all tubes on ice (optional), add 3 mL of 10× loading dye to only the digested PCR products and mix by gently pipetting up and down. 5. Leave tubes containing digested hD1R-pCMV5 and hD5RpCMV5 DNA constructs without loading dye and prepare linearized wild type hD1R and hD5R-pCMV5 DNA samples for dephosphorylation (see Note 12).

3.3. Dephosphorylation of Linearized Wild Type hD1R and hD5R-pCMV5 DNA Constructs

1. Carry out dephosphorylation reaction in using a final volume of 50 mL. Add 14 mL of sterile Milli-Q water to tubes containing the 30 mL of restriction digestion mix of hD1R-pCMV5 (HindIII-XbaI) and hD5R-pCMV5 (BsmI-EagI) expression constructs. Then, add 5 mL of 10× dephosphorylation buffer and 1 mL of CIAP (5 U).

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

155

2. Gently pipette up and down to mix and float tubes in a 37°C water bath for 30 min. 3. At end of incubation, place dephosphorylation tubes on ice (optional), put in 5 mL of 10× loading dye and mix by gentle pipetting up and down. 3.4. Isolation of Linearized Wild Type hD1R and hD5RpCMV5 DNA Constructs and Digested Mutant Receptor DNA Cassettes

1. Digested mutant cassettes and wild type expression constructs are loaded on individual wells along with a well containing 4 mL of DNA size markers of agarose minigels as described above in Subheading 3.1 (see Subheading 3.1.2). Run hD1R and hD5R samples for 1 h at 80 V on 1% (w/v) and 1.8% (w/v) agarose minigels, respectively (see Note 13). Visualize ethidium bromide-stained agarose gel using an UV Transilluminator equipped using a digital camera and excise appropriate bands: (1) HindIII-XbaI linearized hD1R-pCMV5 (~5,400 bp), (2) HindIII-XbaI digested mutant hD1RS263G cassette (745 bp), (3) HindIII-XbaI digested mutant hD1RS263A cassette (745 bp), (4) HindIII-XbaI digested mutant hD1RS263D cassette (745 bp), (5) BsmI-EagI linearized hD5R-pCMV5 (~6,000 bp), (6) BsmI-EagI digested mutant hD5RS287G cassette (348 bp), (7) BsmI-EagI digested mutant hD5RS287A cassette (348 bp), and (8) BsmI-EagI digested mutant hD5RS287D cassette (348 bp). 2. Purify agarose-embedded DNA bands with QIAEX beads (Qiagen) according to manufacturer’s protocol. Elute purified bands from QIAEX beads using 50 mL of sterile Milli-Q water or QIAEX elution buffer. Add 2 mL of purified bands to 7 mL of sterile Milli-Q water, mix with 1 mL of 10× loading dye and run samples beside a well loaded with 4 mL of DNA size markers on 1% (w/v) agarose minigel prepared with thin sample comb for 1 h at 80 V. 3. Visualize ethidium bromide-stained DNA bands using an UV Transilluminator equipped with a digital camera and take picture to assist in the semi-quantification of purified DNAs to set up ligations.

3.5. DNA Ligation Reactions

1. Thereafter and unless stated otherwise, linearized hD1RpCMV5 (~5,400 bp band) and hD5R-pCMV5 (~6,000 bp band) will be called “vector” whereas the mutated receptor cassettes will be referred to as “inserts.” Set up control (vector alone) and test (vector + insert) ligation reactions in a final volume of 10 mL in 1.5 mL Eppendorf tubes on ice (optional). Add to test ligation tubes 0.5 mL vector, 0.5 mL insert (replace with 0.5 mL sterile Milli-Q water in control ligation tubes), 0.5 mL T4 DNA ligase (2.5 U), 1 mL 10× T4 DNA ligase buffer, and 7.5 mL sterile Milli-Q water (see Note 14).

156

B. Plouffe and M. Tiberi

2. Float tubes in a 16°C water bath overnight (see Note 15). Store ligation tubes at −20°C until use for transformation of XL-1 Blue electroporation-competent cells. 3.6. Desalting of DNA Ligation Samples

1. Add 40 mL of sterile Milli-Q water to fresh or thawed ligation tubes (10 mL) at room temperature and gently tap tubes to mix. 2. In a fume hood, add 500 mL of isobutanol to 50 mL ligation samples. 3. Mix by gently inverting tubes several times until isobutanol is fully miscible with aqueous ligation samples (no detection of isobutanol phase remnant or bubbles). 4. Spin tubes in a microfuge at 16,000 × g for 10 min at room temperature. 5. In a fume hood, decant supernatant in waste glass bottle and spin again tubes at 16,000 × g for 30 s. 6. In a fume hood, carefully remove supernatant using a P200 pipette and discard supernatant in waste glass bottle. 7. Let air dry the small DNA pellet in fume hood for 5–10 min, add 10 mL of sterile Milli-Q water to tubes and carefully resuspend DNA pellet by washing sides of tubes.

3.7. Transformation of XL1-Blue ElectroporationCompetent Cells with Ligated DNA Samples

1. Take 5 mL of desalted DNA samples from control and test ligation reactions and separately mix with 40 mL of XL1-Blue electroporation-competent cells on ice by gently pipetting up and down in 1.5 mL Eppendorf tubes. 2. Transfer 45 mL of DNA-bacteria mixtures into ice-cold electroporation cuvettes. Carefully wipe side of electroporation cuvettes to remove any condensation prior to inserting into electroporator. 3. Shock cells at 1,800 V for 5 ms (see Note 16). 4. Add 1 mL of freshly made SOC in each cuvette, gently pipette up and down and transfer 1 mL to sterile polypropylene capped 13 mL tubes (100 × 16 mm). 5. Incubate with loosened cap in a 37°C shaking incubator at a velocity of 300 rpm for 1 h. 6. Pour bacterial cultures into 1.5 mL Eppendorf tubes and spin at 6,000 × g for 30 s at room temperature. Discard ~900 mL supernatant and gently resuspend bacterial pellet with leftover supernatant (~100 mL) by pipetting up and down. 7. Spread ~100 mL of bacterial cultures on pre-warmed (37°C) LB-ampicillin plates and grow bacteria in a 37°C incubator overnight (see Note 17).

10

3.8. Preparation of Plasmid DNA Minipreps and Identification of Single-Point Mutants by Restriction Digestions

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

157

1. Prepare sterile polypropylene capped 13 mL tubes containing 5 mL of LB with 1× ampicilin (100 mg/mL). 2. Pick two isolated colonies from each test ligation plates using small sterile pipette tips and inoculate a LB-ampicillin tube with a single colony by ejecting tip in it. Incubate tubes with loosened caps in a 37°C shaking incubator at a velocity of 300 rpm overnight. 3. Prepare backup bacterial glycerol stocks with overnight cultures. Add 0.7 mL of bacterial cultures to 0.3 mL of sterile 50% (v/v) glycerol (final concentration: 15% (v/v)) in 1.5 mL Eppendorf tubes and gently mix by inverting tubes several times. Snap-freeze in liquid nitrogen. Store at −80°C (see Note 18). 4. Make miniprep DNA with the rest of bacterial cultures (~4.3 mL) using QIAprep Spin Columns according to manufacturer’s protocol (see Note 19). 5. Measure DNA concentration and purity of plasmid miniprep (total volume of 50 mL). Add 5 mL of plasmid miniprep DNAs in a final volume of 1 mL of sterile Milli-Q water in quartz UV cuvettes. Read optical density (OD) against a blank solution (1 mL of sterile Milli-Q water) at wavelengths of 260 nm and 280 nm using spectrometer (see Note 20). 6. Set up diagnostic digestion reactions in a final volume of 15 mL with and without appropriate restriction enzymes as follows (see Table 3). 7. For the screening of positive hD1R-S263A plasmid DNAs, carry out restriction digestions in 1.5 mL Eppendorf tubes containing 2 mL of plasmid DNA (0.5 mg; stock solution of 0.25 mg/mL), 1.5 mL of 10× red buffer, 0.5 mL of BsmI (5U),

Table 3 Expected band size pattern of wild type and single-point mutants of hD1R and hD5R following digestion with restriction enzymes Constructs

Restriction enzymes

Band sizes (base pairs)

hD1R-S263A

BsmI + EcoRI

Wild Type hD1R: 6025 Mutant hD1R: 804, 5221

hD1R-S263G hD1R-S263D

DraI

Wild Type hD1R: 19, 692, 1666, 3648 Mutant hD1R: 19, 692, 1112, 1666, 2536

hD5R-S287A hD5R-S287G hD5R-S287D

BoxI

Wild Type hD5R: 6322 Mutant hD5R: 612, 5710

For each single-point mutant, the diagnostic band sizes of digested positive plasmid DNAs are bold and underlined

158

B. Plouffe and M. Tiberi

0.5 mL of EcoRI (5U), and 10.5 mL of sterile Milli-Q water (11.5 mL for uncut condition). 8. For the screening of positive hD1R-S263G and hD1R-S263D plasmid DNAs, carry out restriction digestions in 1.5 mL Eppendorf tubes containing 2 mL of plasmid DNA (0.5 mg; stock solution of 0.25 mg/mL), 1.5 mL of 10× blue buffer, 0.5 mL of DraI (5U), and 11 mL of sterile Milli-Q water (11.5 mL for uncut condition). 9. For the screening of positive hD5R-S287A, hD5R-S287G and hD5R-S287D plasmid DNAs, carry out restriction digestions in 1.5 mL Eppendorf tubes containing 2 mL of plasmid DNA (0.5 mg; stock solution of 0.25 mg/mL), 1.5 mL of 10× Y+/ TangoTM buffer, 0.5 mL of BoxI (5U), and 11 mL of sterile Milli-Q water (11.5 mL for uncut condition). 10. Gently pipette up and down to mix and float tubes in a 37°C water bath for 1 h. At the end of incubation, add 1.5 mL of 10× loading dye, mix by gently pipetting up and down and run samples beside a well loaded with 4 mL of DNA size markers on 1% (w/v) agarose minigel for 1 h at 80 V. 11. Visualize ethidium bromide-stained DNA bands using an UV Transilluminator equipped with a digital camera, take picture and identify positive mutant plasmid DNAs according to expected band sizes of digested wild type and mutated DNA (see Table 3). 3.9. Preparation of Samples for Automated DNA Sequencing and Plasmid DNA Maxipreps

1. Prepare samples for automated DNA sequencing as follows (see Note 21). 2. Make working solution of plasmid DNA minipreps at a final concentration of 12.5 ng/mL with sterile Milli-Q water in autoclaved 1.5 mL Eppendorf tubes. 3. For each single-point mutants of hD1R, set up autoclaved 1.5 mL Eppendorf tubes containing 10 mL (125 ng) of plasmid DNA and 5 mL (10 pmol) of hD1R-P1 forward primer or hD1R-P4 reverse primer (see Table 1) (see Note 22). 4. For each single-point mutants of hD5R, set up autoclaved 1.5 mL Eppendorf tubes containing 10 mL (125 ng) of plasmid DNA and 5 mL (10 pmol) of hD5R-P6 reverse primer (see Table 2). 5. Align sequenced DNAs with predicted nucleotide sequences and confirm that (1) serine is mutated, (2) mutated receptor DNA cassette is ligated in-frame with the expression vector pcCMV5 containing wild type hD1R or hD5R DNA sequences, and (3) integrity of restriction sites used for subcloning the mutated hD1R (HindIII and XbaI) and hD5R (BsmI and EagI) cassettes.

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

159

6. Once positive clone identify and DNA integrity confirmed, make large-scale plasmid DNA preparations as follows. 7. Stab glycerol stocks of positive bacterial clones with a sterile straight wire and streak pre-warmed (37°C) LB-ampicillin plates. Grow bacteria in a 37°C incubator overnight. 8. The next day, prepare sterile polypropylene capped 13 mL tubes containing 1 mL of LB with 1× ampicilin (100 mg/mL) and inoculate each tube with a single colony using sterile small pipette tips as described above (see Subheading 3.8, step 2). Incubate tubes with loosened caps in a 37°C shaking incubator at a velocity of 300 rpm for 6 h. 9. Subsequently, add 10 mL of the small bacterial cultures to 1 mL of sterile LB in 1.5 mL Eppendorf tubes (see Note 23). Open aluminum foil on top of already made sterile Fernbach flasks (see Subheading 2.1, item 13) containing 500 mL of LB and 1× ampicillin with 1 mL of the diluted small bacterial cultures. Incubate flasks in a 37°C shaking incubator at a velocity of 300 rpm overnight. 10. Prepare large-scale plasmid DNAs with Maxiprep Column Kit according to Qiagen’s protocol. Measure plasmid DNA concentration and purity (see Subheading 3.8, step 5). 11. Verify integrity of plasmid receptor DNAs using digestions with restriction enzymes (see Subheading 3.8, steps 6–11). 3.10. Preparation and Transfection of HEK293 Cells

1. Make frozen stocks of HEK293 cells in Nalgene cryovials at a density of 5 × 106 cells per mL of sterile freezing medium (10% tissue culture grade DMSO, 20% FBS, 70% MEM with Earle’s salts and 40 mg/mL gentamicin) (see Note 24). 2. Grow working stocks of HEK293 cells in polystyrene 75 cm2 flasks containing 20 mL of complete MEM (10% (v/v) FBS and 40 mg/mL gentamicin) at 37°C in a humidified 5% CO2 incubator and maintain stocks as described previously (19). 3. Prepare HEK293 cells for transfection as follows. 4. Aspirate medium from 75 cm2 flasks, add 5 mL of room temperature PBS and wash cells by gently rocking flasks (see Note 25). 5. Aspirate PBS, add 1 mL trypsin to flasks, briefly incubate cells at room temperature (<1 min) (see Note 26). 6. Stop trypsinization by adding 20 mL of complete MEM per flask (see Note 27). 7. Mix cells by gentle trituration using 10 mL pipette. Pipette up and down 10–15 times to detach cells and avoid clump formation. Count cells using hemacytometer.

160

B. Plouffe and M. Tiberi

8. For the transfection procedure, seed polystyrene tissue culture dishes (100 × 20 mm) with 2 × 106 cells in final volume of 10 mL of complete MEM and grow cells until the next day (24–30 h) at 37°C in a humidified 5% CO2 incubator (see Note 28). 9. Prepare transfection solutions in sterile 13 mL polypropylene tubes (100 × 16 mm) in a BSC as follows. 10. Add a total quantity of 10 mg of different plasmid DNAs to be transfected in a final volume of 50 mL or less of Milli-Q water to 13 mL tubes (see Note 29). 11. Add sterile Milli-Q water to each tube containing plasmid DNA to final volume of 900 mL. 12. Add 100 mL of sterile 2.5 M CaCl2 to each tube (final volume 1 mL) (see Note 30). 13. Add 1 mL of sterile 2× HEPES-buffered saline solution to DNA-calcium phosphate mixture in a dropwise manner using a P1000 pipette. Mix by gentle flicking of tubes (final volume of 2 mL) (see Note 31). 14. Transfect two 100 × 20 mm dishes of HEK293 using 1 mL per dish as follows. 15. Open the lid of a dish, add dropwise 1 mL of transfection solution to the whole medium surface, close lid, and incubate HEK293 cells overnight at 37°C in humidified 5% CO2 incubator (see Note 32). 16. The next day, split cells as follows (see Note 33). 3.11. Seeding of Transfected HEK293 Cells for Radioligand Binding and Whole Cell cAMP Assays

1. For each transfection condition, aspirate culture medium from the four dishes in a BSC, add 5 mL of room temperature PBS per dish to wash cells (see previous Note 25), aspirate PBS, add 0.5 mL of 1× trypsin per dish, incubate briefly, add 10 mL of complete MEM and triturate cells using gentle pipetting up and down (see Subheading 3.10, step 7). 2. For radioligand binding studies, pool cells from four 100 × 20 mm dishes into a 150 × 25 mm dish (final volume ~40 mL) and grow transfected HEK293 cells for ~48 h at 37°C in humidified 5% CO2 incubator (see Note 34). 3. For dose–response curves using whole cell cAMP assays, pool cells from four 100 × 20 mm dishes (final volume ~40 mL) and seed two 12-well plates with 1 mL of cells per well (total volume required is 24 mL) for each experimental condition. Seed also one 100 × 20 mm dish with 10–15 mL of cells per dish for determination of receptor levels in membrane preparations from cells used for dose–response curves (see Note 35).

10

3.12. Preparation of Crude Membranes from Transfected HEK293 Cells for Saturation Studies

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

161

1. On the day of the experiment, put 150 × 25 mm dishes on ice, aspirate culture medium, add 10 mL of cold PBS to side of dishes and wash cells by gentle rocking of dishes. 2. Aspirate PBS, add 15 mL of ice-cold lysis buffer, detach cells with a cell lifter by scraping off the dish surface and transfer lysates to 50 mL polycarbonate centrifuge tubes (29 × 104 mm, Beckman Coulter). Wash dishes again with 15 mL of ice-cold lysis buffer, harvest 5 mL wash and put in centrifuge tubes. 3. Centrifuge samples at 40,000 g for 20 min at 4°C and put tubes on ice. 4. Discard supernatant, add 3 mL of ice-cold lysis buffer to centrifuge tubes, pipette up and down to detach pellets and homogenize pellets in centrifuge tubes with a Brinkmann Polytron at a velocity of 17,000 rpm for 15 s. Adjust final volume in tubes to 30 mL and centrifuge at 40,000 g for 20 min at 4°C. 5. Discard supernatant, add 3 mL of ice-cold lysis buffer to centrifuge tubes, pipette up and down to detach pellets and homogenize pellets in centrifuge tubes with a Brinkmann Polytron at a velocity of 17,000 rpm for 15 s. 6. Add 0.6 mL of membrane preparations to tubes containing 3 mL of cold resuspension buffer (dilution factor of 1:6) and leave tubes on ice until used for saturation studies. Add the remnant of membrane preparations in lysis buffer in two 1.5 mL Eppendorf tubes (~1.2 mL per tube), snap-freeze in liquid nitrogen and store at −80 C until used for competition studies.

3.13. Saturation Studies

1. Set up four three-tier polypropylene racks (6 × 12 holes per row, Fisher Scientific) containing 48 polystyrene test tubes (12 × 75 mm) to carry binding reactions in a final volume of 500 mL (see Note 36). 2. Prepare six 10× concentrations of [3H]-SCH23390 (84 Ci/ mmol) in Milli-Q water ranging from ~0.1 to 100 nM (final in assays: ~0.01–10 nM) using a dilution factor of 1:3. 3. Prepare 10× cis-flupenthixol (100 mM) in Milli-Q water using frozen stock (final in assays: 10 mM) (see Note 5). 4. Add 300 mL of binding buffer to all tubes (final in assays: 50 mM Tris–HCl, pH 7.4, 120 mM NaCl, 4 mM MgCl2, 1.5 mM CaCl2, 5 mM KCl, and 1 mM EDTA, pH 8.0). 5. Add 50 mL of Milli-Q water to total binding tubes or 50 mL of 10× cis-flupenthixol (100 mM) to nonspecific binding tubes. 6. Add 50 mL of different 10× [3H]-SCH23390 concentrations to a set of four tubes from the lowest to highest concentration (see Note 36).

162

B. Plouffe and M. Tiberi

7. Add 100 mL of membrane preparations to tubes (24 tubes for each receptor tested), shake racks to mix and incubate at room temperature (~20°C) for 90–120 min. 8. Measure protein concentration in leftover membrane preparations using Bio-Rad assay kit with BSA as standard according to manufacturer’s protocol (see Note 37). 9. Terminate binding reactions by rapid filtration through precut Whatmann GF/C glassfiber filter sheets (11.4 cm × 31.1 cm) using Brandel semi-automated harvesting system (equipped with 48 harvesting probes) and wash membranes bound to filters three times with 5 mL of cold washing buffer (tubes are subjected to three fill up and aspiration cycles). 10. Put filter circles in plastic scintillation vials, add 5 mL of scintillation cocktail and determine tritium-bound radioactivity in beta liquid scintillation counter with 30–40% counting efficiency. 11. Analyze binding isotherms with a nonlinear curve-fitting software from GraphPad Prism (GraphPad Software, San Diego, CA). Determine equilibrium dissociation constant (Kd, nM) and maximal binding capacity (Bmax, pmol/mg membrane proteins) of [3H]-SCH23390 with saturation curves (see Note 38). A representative example of saturation curves obtained in membrane preparations from transfected HEK293 cells with wild type (WT) and single-point mutant forms of hD1R and hD5R is shown in Fig. 4. Averaged Kd and Bmax values of [3H]-SCH23390 are reported in Table 4. Data are also expressed relative to hD1R-WT and hD5R-WT (Fig. 5). The S287A, S287G and S287D mutations have no significant effect on Kd of [3H]-SCH23390 while Bmax values are differentially modulated. Meanwhile, the S263G mutation in hD1R leads to a small but significant reduction of Kd of [3H]-SCH23390. This increased affinity of hD1R-S263G for [3H]-SCH23390 is not observed in cells transfected with hD1R-S263A and hD1RS263D. Moreover, while Bmax of hD1R-S263D is not changed relative to hD1R-WT, hD1R-S263G and hD1R-S263A display a reduction and augmentation of Bmax, respectively. 3.14. Competition Studies

1. Set up 16 three-tier polypropylene racks (6 × 12 holes per row, Fisher Scientific) containing 48 polystyrene test tubes (12 × 75 mm) to carry binding reactions in a final volume of 500 mL (see Note 39). 2. Thaw frozen tubes of membranes on ice and mix with 11 mL of resuspension buffer using Brinkmann Polytron at velocity of 17,000 rpm for 15 s. Leave membrane resuspension on ice until use (see Note 40). 3. Prepare a 10× concentration of [3H]-SCH23390 (84 Ci/ mmol) in Milli-Q water ranging from ~7 to 13 nM (final in assays: ~0.7–1.3 nM) (see Note 41).

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

163

Fig. 4. Saturation curves for HEK293 cells transfected with wild type and single-point mutant forms of hD1R and hD5R. Representative examples for saturation curves generating data for Table 4 are shown. Saturation curves of [3H]-SCH23390 on crude membrane preparations were determined in the presence of 10 mM cis-flupenthixol. TOTAL total binding, NS nonspecific binding, hD1R human D1 receptor, hD5R human D5 receptor, WT wild type.

0.82 (0.64–1.05)

0.64 (0.43–0.96)

0.59 (0.40–0.86)

0.69 (0.55–0.86)

1.22 (0.90–1.65)

1.03 (0.87–1.22)

0.95 (0.69–1.31)

1.32 (1.11–1.56)

hD1R-WT

hD1R-S263A

hD1R-S263G

hD1R-S263D

hD5R-WT

hD5R-S287A

hD5R-S287G

hD5R-S287D

17.6 (14.3–20.8)

12.1 (10.3–14.0)

17.6 (15.6–19.7)

13.9 (10.1–17.7)

13.4 (9.6–17.2)

7.8 (5.1–10.6)

18.7 (12.5–24.9)

13.5 (9.2–17.8)

Bmax (pmol/mg prot.)

1.08 (0.95–1.23)

0.89 (0.68–1.15)

0.91 (0.75–1.10)

0.98 (0.84–1.15)

0.70 (0.60–0.82)

0.57 (0.42–0.77)

0.66 (0.57–0.78)

0.80 (0.56–1.13)

SCH23390

Competition studies Ki (nM)

841 (636–1,113)

423 (314–572)

916 (605–1,388)

735 (542–997)

6,734 (5,369–8,446)

2,815 (2,341–3,385)

11,085 (9,273–13,252)

7,420 (4,019–13,699)

Dopamine

15.2 (13.1–17.7)

11.0 (9.27–13.0)

11.8 (9.73–14.3)

12.1 (8.69–16.9)

8.50 (6.11–11.8)

7.25 (5.49–9.58)

8.37 (6.04–11.6)

9.42 (5.89–15.1)

cis-Flupenthixol

30.0 (16.3–55.2)

23.6 (14.0–39.8)

24.7 (15.4–39.4)

29.3 (18.2–47.3)

4.73 (3.70–6.03)

4.39 (3.46–5.55)

4.98 (3.51–7.05)

5.15 (3.35–7.93)

(+)-Butaclamol

Data are expressed as geometric (Kd, Ki) and arithmetic (Bmax) means with 95% lower and upper confidence intervals from 5 to 6 experiments done in duplicate determinations. Best-fitted parameters were obtained from binding isotherms analyzed using nonlinear curve regression programs from GraphPad Prism. hD1R-WT, wild type human D1R; hD5R-WT, wild type human D5R; Kd, equilibrium dissociation constant; Bmax, maximal binding capacity; Ki, equilibrium dissociation constant of unlabeled drug

Kd (nM)

Receptor

Saturation studies [3H]-SCH23390

Table 4 Ligand binding properties of wild type and single-point mutant forms of dopamine hD1R and hD5R

164 B. Plouffe and M. Tiberi

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

165

Fig. 5. Relative equilibrium dissociation constant (Kd) and maximal binding capacity (Bmax ) values of hD1R and hD5R single-point mutants. Arithmetic means ± S.E. of Kd (a) and Bmax (b) values of hD1R and hD5R single-point mutants were calculated relative to their respective wild type receptor counterpart. *P < 0.05 when compared with a value of 1 (wild type receptor) using one-sample t test. hD1R-WT wild type human D1 receptor, hD5R-WT wild type human D5 receptor.

4. Prepare 11 increasing 10× concentrations of SCH23390 (~3 × 10−11 to 3 × 10−6 M; ~3 × 10−12 to 3 × 10−7 M final in assays), cis-flupenthixol (~4 × 10−10 to 5 × 10−5 M; ~4 × 10−11 to 5 × 10−6 M final in assays) and (+)-butaclamol (~3 × 10−10 to 3 × 10−5 M; ~3 × 10−11 to 3 × 10−6 M final in assays) in Milli-Q water using frozen stock (see Note 5). 5. Prepare a fresh 10× ascorbic acid solution (1 mM) in Milli-Q water. 6. Prepare 11 increasing 10× concentrations of dopamine (~3 × 10−8 to 1 × 10−2 M; final in assays) in 10× ascorbic acid solution (see Note 42). 7. Add 300 mL of binding buffer (final in assays: 50 mM Tris– HCl, pH 7.4, 120 mM NaCl, 4 mM MgCl2, 1.5 mM CaCl2, 5 mM KCl, and 1 mM EDTA, pH 8.0). 8. Add 50 mL of Milli-Q water or 10× ascorbic acid to total binding tubes.

166

B. Plouffe and M. Tiberi

9. Add 50 mL of different 10× concentrations of cold drugs from the lowest to highest concentration. 10. Add 50 mL of [3H]-SCH23390.

the

fixed

10×

concentration

of

11. Add 100 mL of membrane preparations to tubes (96 tubes for each receptor tested), shake racks to mix and incubate at room temperature (~20°C) for 90–120 min. 12. Terminate binding reactions by rapid filtration through precut Whatmann GF/C glassfiber filter sheets (11.4 cm × 31.1 cm) using Brandel semi-automated harvesting system and wash membranes bound to filters three times with ~5 mL of cold washing buffer (tubes are subjected to three fill up and aspiration cycles). 13. Put filter circles in plastic scintillation vials, add 5 mL of scintillation cocktail and determine tritium-bound radioactivity in beta liquid scintillation counter with 30–40% counting efficiency. 14. Analyze binding isotherms using GraphPad Prism software. Determine equilibrium dissociation constant of cold (unlabeled) drugs (Ki, nM). A representative example of competition curves obtained in membrane preparations from transfected HEK293 cells with WT and single-point mutant forms of hD1R and hD5R is depicted in Fig. 6. Averaged Ki values of unlabeled drugs are reported in Table 4. Data are also expressed relative to hD1R-WT and hD5R-WT (Fig. 7). None of the mutations has a major effect on the affinity of cis-flupenthixol and (+)-butaclamol, two inverse agonists. Interestingly, in both hD1R and hD5R, the S-to-G mutation leads to a significant increase in dopamine affinity (lower Ki value) of receptors while S-to-D mutation had no effect (Table 4). Additionally, we observe that the S-to-A mutation promotes a decrease in dopamine affinity of hD1R and hD5R, a trend that is not however statistically significant. Importantly, Ki values measured with unlabelled SCH23390 are in line with Kd values of [3H]-SCH23390 (Table 4). It is also worth mentioning that the increased affinity of hD1R-S263G for [3H]-SCH23390 was also observed with unlabelled SCH23390 using competition studies (Table 4). 3.15. Establishment of Dopamine Dose– Response Curves using Whole Cell cAMP Assays

1. Prepare MEM containing 5% (v/v) FBS and 40 mg/mL gentamicin in a BSC and add sterile [3H]-adenine (stock at 1 mCi/ mL) using a dilution factor 1:1,000 to obtain a final activity of 1 mCi/mL. 2. Aspirate medium from 12-well plates, add 1 mL of prewarmed (37°C) labeling media per well and incubate HEK293 cells with [3H]-adenine overnight at 37°C in humidified 5% CO2 incubator (see Note 43).

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

167

Fig. 6. Competition curves for HEK293 cells transfected with wild type and single-point mutant forms of hD1R and hD5R. Representative examples for competition curves generating data for Table 4 are shown. Competition curves on crude membrane preparations from wild type and single-point mutant forms of hD1R (left panels) and hD5R (right panels) were performed with 0.5–1.2 nM of [3H]-SCH23390 in the absence and presence of increasing concentrations of unlabeled drugs. Concentration of unlabeled drugs (M) is given as log values. hD1R-WT wild type human D1 receptor, hD5R-WT wild type human D5 receptor.

Fig. 7. Relative equilibrium dissociation constant of unlabeled drug (Ki) values of hD1R and hD5R single-point mutants. Arithmetic means ± S.E. of Ki values of hD1R and hD5R single-point mutants were calculated relative to their respective wild type receptor counterpart. *P < 0.05 when compared with a value of 1 (wild type receptor) using one-sample t test. hD1R-WT wild type human D1 receptor, hD5R-WT wild type human D5 receptor.

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

169

3. Prepare two scintillation vials containing 50 mL of labeling media and 5 mL of scintillation cocktail. Count in a beta liquid scintillation counter (see Note 44). 4. After an overnight labeling with [3H]-adenine, perform whole cell cAMP assays as follows. 5. Thaw 200 mM IBMX stock and prepare 20 mM HEPESbuffered MEM containing 1 mM IBMX. Keep in a 37°C water bath until use. 6. Prepare racks of polystyrene test tubes (12 × 75 mm) containing 100 mL of neutralizing solution. 7. Prepare a fresh 100× ascorbic acid solution (10 mM) in Milli-Q water. 8. Prepare seven increasing 100× concentrations of dopamine (10−9 to 10−3 M; 10−11 to 10−5 M final in assays) in 100× ascorbic acid solution using 1:10 dilution factor (see Note 42). 9. Aspirate labeling medium and add 1 mL of 20 mM HEPESbuffered MEM containing 1 mM IBMX per well. 10. Add 10 mL of 100× ascorbic acid to three wells (triplicate determinations). 11. Add 10 mL of increasing 100× concentrations of dopamine in triplicate to remaining wells of the two 12-well plates. 12. Incubate 12-well plates at 37°C for 30 min (see Note 45). 13. At the end of treatment period, put 12-well plates on ice tray, aspirate medium, add 1 mL of cAMP stop solution and incubate cells at 4°C for 30 min. 14. Transfer lysates from wells to tubes containing neutralizing solution, mix using vortex and put a parafilm sheet to seal top of tubes in racks. Store at 4°C until use (see Note 46). 15. Prepare membranes from the binding dish as follows. 16. Put dishes on ice tray, aspirate culture medium and add 5 mL of cold PBS to side of dishes and wash by gentle rocking of dishes. 17. Aspirate PBS, add 5 mL lysis buffer, detach cells with a cell lifter by scraping off the dish surface and transfer lysates to a 15 mL polycarbonate centrifuge tubes (18 × 100 mm, Beckman Coulter). Wash dishes again with 5 mL of lysis buffer and transfer whole volume to centrifuge tubes (final volume of 10 mL). Prepare membranes essentially as described in Subheading 3.12 (steps 3–6). Resuspend final pellets in 0.6–1 mL of resuspension buffer and perform binding reactions using 100 mL of membranes and a saturating concentration of [3H]-SCH23390 in the absence and presence of cold 10 mM cis-flupenthixol as described above (see Subheading 3.13). Measure protein concentrations with the Bio-Rad assay kit with BSA as standard to determine Bmax in pmol/mg membrane proteins.

170

B. Plouffe and M. Tiberi

3.16. Purification of [3H]-cAMP with Sequential Chromatography on Dowex and Alumina Columns and Quantification of Intracellular [3H]-cAMP Mediated by Dopamine

1. Mount Bio-Rad Poly-Prep columns on custom-made Plexiglas racks (100 per rack), fill separate set of columns with Dowex and alumina and establish elution profile of columns using 0.1 N NaOH/ 0.1 N HCl/distilled water (Dowex) and 0.1 M imidazole (alumina) as previously described (19). 2. Prepare KOH-neutralized cAMP samples for sequential chromatography as follows. 3. Prepare also triplicate samples of 1 mL of cAMP stop solution in tubes containing neutralizing solution, mix using vortex and store with [3H]-cAMP lysate samples at 4°C (see Note 47). 4. Centrifuge test tubes containing KOH-neutralized cAMP samples at ~500 g (1,500 rpm) in Beckman Coulter AllegraTM 6R Centrifuge for 10 min (see Note 48). 5. Load 0.85 mL of samples on separate Dowex columns and let drain the flow through fraction (see Note 49). 6. Add 1 mL of distilled water to each column, let drain the water wash fraction from Dowex and elute samples from Dowex on alumina columns using 4 mL of distilled water. 7. Let drain water wash fraction from alumina, wash with 1 mL of 0.1 M imidazole and let drain the imidazole wash fraction. 8. Elute samples from alumina columns into plastic scintillation vials using 3 mL of 0.1 M imidazole. Add 18 mL of scintillation cocktail. Shake vials vigorously to mix aqueous samples (~3 mL) with scintillation cocktail and count vials in beta liquid scintillation counter using dual 3H and 14C channels (see Note 50). 9. Take a 50 mL aliquot from leftover volume (~200 mL) of each KOH-neutralized cAMP lysate samples, add to plastic scintillation vials, fill vials with 5 mL of scintillation cocktail and count in beta liquid scintillation counter using dual 3H and 14C channels (see Note 51). 10. Enter 3H and 14C counts in Excel spreadsheet and compute intracellular [3H]-cAMP levels in the absence and presence of increasing concentrations of dopamine using formulas described in Plouffe et al. (19). 11. Compute intracellular cAMP levels expressed in arbitrary units (CA/TU × 1,000) and analyze averaged dose–response curves with a four-logistic parameter equation using a nonlinear curvefitting program from GraphPad Prism to determine effective concentration of dopamine that produces 50% of maximal stimulation (EC50, an index of potency) and dopamine-mediated maximal stimulation of AC (Emax). The best-fitted dose–response curves to dopamine obtained with cells expressing similar receptor levels are reported in Fig. 8. Our data show that S-to-A, S-to-G, and S-to-D mutations modulate EC50 and Emax.

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

171

Fig. 8. DA-induced formation of intracellular cAMP by wild type and single-point mutant forms of hD1R and hD5R in intact HEK293 cells. Cells were grown in medium containing 1 mCi/mL of [3H]-adenine overnight. Intracellular cAMP levels were determined in 12-well dishes in the absence and presence of increasing concentration of DA (10−11 to 10−5 M). Each point represents the arithmetic mean ± S.E. of three to four experiments done in triplicate determinations. (a) Dose–response curves to DA were simultaneously analyzed by a four-parameter logistic equation using GraphPad Prism version 5.03 using constrained or unconstrained parameters. Best-fitted values for effective concentration that elicits 50% of maximal stimulation (EC50, nM) with 95% lower and upper confidence intervals are as follows: 14.7 (6.87–31.4) for hD1R, 43.8 (18.5–104) for hD1R-S263A, 9.90 (5.20–18.9) for hD1R-S263G, 30.1 (13.2–68.4) for hD1R-S263D, 1.50 (0.64–3.49) for hD5R, 3.15 (1.11–8.94) for hD5R-S287A, 0.72 (0.28–1.85) for hD5R-S287D, and 1.96 (0.68–4.98) for hD5R-S287G. (b) Best-fitted values for Emax ± S.E. obtained from dose–response curves to dopamine in HEK293 cells expressing wild type and singlepoint mutant forms of hD1R and hD5R are shown. (c) EC50 shift (expressed as arithmetic mean ± S.E.) of single-point mutants were calculated relative to wild type receptor using EC50 values derived from individual fitted dose–response curves. *P < 0.05 when compared with wild type receptor. The Bmax values in pmol/mg/membrane proteins (expressed as arithmetic mean ± S.E.) are as follows: 1.04 ± 0.18 (hD1R-WT), 1.34 ± 0.32 (hD1R-S263A), 1.19 ± 0.35 (hD1R-S263G), 1.15 ± 0.20 (hD1R-S263D), 1.12 ± 0.34 (hD5R), 1.31 ± 0.28 (hD5R-S287A), 0.83 ± 0.04 (hD5R-S287G), and 1.04 ± 0.26 (hD5R-S287D). hD1R-WT wild type human D1 receptor, hD5R-WT wild type human D5 receptor.

172

B. Plouffe and M. Tiberi

Interestingly, the trend in mutation-specific effects is generally recapitulated in cells expressing hD1R and hD5R mutants (Fig. 8). Data obtained with the S-to-G mutation are in agreement with the increased dopamine affinity for hD1R-S263G and hD5R-S287G (Table 4 and Fig. 7). The S-to-G mutation may thus promote the release of molecular constraints within the cytosolic end of TM6, leading to a more “relaxed” conformation for agonist binding and agonist-dependent G protein coupling. This idea is to some extent supported by our binding data showing increased affinity of hD1R-S263G for SCH23390 (Figs. 6 and 7). Indeed, SCH23390 is a classical D1-like antagonist that behaves pharmacologically as a partial agonist in HEK293 cells (9, 20). In addition, the decreased potency (EC50 rightward shift) observed with the S-to-A mutation (Fig. 8) suggests a potential role of the hydroxyl group on the side chain of serine in the formation of hydrogen bonds, which may be only involved in agonist-dependent G protein coupling. Indeed, in contrast to S-to-G mutation, the S-to-A mutation has no effect on agonist binding (Table 4 and Fig. 7). Therefore, S-to-A mutation may impart subtle structural changes to the cytosolic end of TM6 that specifically impact the agonistdependent G protein-coupling conformation. Meanwhile, the S-to-D mutation in hD1R (and to a lesser extent in hD5R) paradoxically increases EC50 (lower potency, i.e., reduced G protein coupling) and Emax (greater dopamine efficacy) (Fig. 8). These results may imply that the phosphomimetic S-to-D mutation promotes the formation of an ionic bond between the negatively charged side chain in aspartic acid and a positively charged amino acid (arginine, histidine or lysine). Ultimately, this may lead to the formation of a new GPCR conformation with distinct G protein coupling properties when compared with wild type receptors. However, it remains to be established whether results obtained with the phosphomimetic S-to-D mutation can also be recapitulated with the phosphorylation of S263 and S287 in hD1R and hD5R, respectively. In closing, it is also worth mentioning that the role of S263 (hD1R) and S287 (hD5R) in regulating the constitutive activity will require additional studies. In fact, dose–response curves in 12-well dishes do not provide the best experimental framework to address this important question (see Note 52).

4. Notes 1. Ethidium bromide wastes (tip, gel) should be disposed as per the experimenter’s research institution rules. 2. The 1 M HEPES solution is utilized to prepare 2× HEPESbuffered saline solution, which is used in the transfection

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

173

procedure. Note that the pH of this HEPES solution is different from that of the sterile HEPES buffer solution (pH 7.4) employed in the preparation of 20 mM HEPES-buffered MEM (whole cAMP assays). 3. Na3PO4 precipitates can form over time. Precipitates can be easily dissolved by swirling solution in a 37°C water bath prior to preparing stock of 2× HEPES-buffered saline solution. 4. Handling and disposal of radioactive material should be done as per radiation safety office rules of the experimenter’s research institution. 5. Experimenter can make stock solution of cold drugs in Milli-Q water (SCH23390, cis-flupenthixol) or ethanol ((+)-butaclamol) at a final concentration of 10 mM. Store stock solution of drugs at −20°C. However, ascorbic acid and dopamine solutions are always made fresh. 6. The IBMX-DMSO stock solution will freeze at 4°C. The 200 mM IBMX is made in a 50 mL polypropylene conical tube, which facilitates rapid thawing in 37°C water bath. We recommend preparing small volume of solution, typically using 5 mL of DMSO, to avoid multiple “freeze–thaw” cycles. The IBMX stock solution should be discarded if experimenters note that it is unfrozen when stored at 4°C. 7. Typically, stop solution is prepared in large polypropylene graduated cylinder (2 L) and carefully poured in two dark glass bottles (~750 mL) fitted with a bottletop dispenser. The stop solution can be kept for 4 months at 4°C. Unlabelled cAMP is used to saturate phosphodiesterases and hence to prevent degradation of [3H]-cAMP formed by receptor stimulation. [14C]-cAMP is used as a tracer to measure [3H]-cAMP recovery during chromatography column procedure. 8. The imidazole working solution is diluted in Milli-Q water at 0.1 M. 0.1 M imidazole is stored at room temperature in a glass bottle equipped with bottletop dispenser and used during alumina column washing and elution procedure. 9. The electrophoresis of uncut DNA is recommended and useful in assessing the efficiency of digestion with selected restriction enzymes and isolation of positive clones for single-point mutants. 10. The volume of purified mutant receptor cassette can be increased to 16 mL if the recovery yield of mutant receptor cassette is low after purification using QIAEX beads. 11. Double digestions using restriction enzymes from Fermentas are typically performed in either 2× or 1× buffer Y+/TangoTM (yellow) according to optimal enzyme activity. For a double digestion using BsmI and EagI, 2× buffer Y+/TangoTM gives the highest enzyme activities.

174

B. Plouffe and M. Tiberi

12. Do not add loading dye following digestion of hD1R-pCMV5 and hD5R-pCMV5 DNA constructs because it will compromise CIAP dephosphorylation. This step is critical for the ligation reaction. It allows dephosphorylation of 5¢-ends of digested constructs DNA that have not be fully cut with both enzymes and hence reducing the self-ligation of single enzyme-digested DNA molecules during ligation reactions. Ultimately, this will decrease the number of false positive on your test ligation plates. 13. A fraction of the hD5R DNAs is partially digested with BsmI and EagI. It is thus recommended to run side by side uncut and cut samples on 1.8% (w/v) agarose gel to visualize and isolate DNA bands fully digested by BsmI and EagI. 14. We usually prepare ligation reactions using a vector and insert ratio of 1:3. The volume of vector and insert used in our ligation reactions is based on relative size and intensity between vector and insert bands. If yield of purified vector is far in excess of your purified insert, we recommend adding 0.5 mL of a 1:10 dilution of the purified vector in sterile Milli-Q water. 15. Typically, a water bath is placed in cold room or chromatography refrigerator (4°C) and temperature of water bath is set to reach 16°C. 16. Optimal desalting is critical for electroporation of bacteria. If high salt concentrations remain in your ligation reactions, this will create an electrical discharge or arcing during electroporation. Arcing will significantly decrease the viability of bacteria and ultimately lead to a poor efficiency of transformations. 17. We generally obtain less than 5 colonies following transformation with control ligations while test ligation yield between 25 and 50 colonies. The rate of positive colonies ranged between 80 and 90%. 18. The preparation of bacterial glycerol stocks of selected colonies prior to making plasmid DNA minipreps for analysis using restriction digestions and automated DNA sequencing will avoid retransforming bacteria with the positive plasmid DNA following the confirmation of the positive mutation. Overall, having readily available bacterial glycerol stocks will save the experimenter from redoing steps described in Subheading 3.7 and thus work more efficiently. 19. Alternatively, boiling methods can be used to prepare plasmid DNA minipreps. The boiling methods are perhaps more costefficient in comparison to using QIAprep Spin columns and as suitable for restriction digestions. However, in our experience, boiling methods do not give high enough DNA quality for automated DNA sequencing and thus the experimenter will have to rely on other methods to make higher quality plasmid

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

175

DNA preps. In our opinion, the cost does not justify using boiling methods. 20. Concentrations of plasmid DNA minipreps range from 0.25 to 0.5 mg/mL with a purity (OD260/OD280 ratio) of ~1.8–1.9. 21. Automated DNA sequencing protocol may vary according to company or core facility’s protocol. Make sure that method described herein is suitable with core facility or company performing automated DNA sequencing. 22. If the same oligonucleotides is used as PCR and sequencing primers, experimenter will make sure that primers are made at appropriate concentrations for PCR and sequencing reactions. 23. This step does not require the addition of ampicillin as diluted bacterial cultures are immediately inoculated into LB containing 1× ampicillin. 24. ATTC suggest that HEK293 cells be grown in 15% (v/v) horse serum. However, we have replace horse serum with heat-inactivated FBS without aberrant effect on cell growth. 25. Do not add PBS directly on cells as they may detach from flasks before adding trypsin. 26. You can gently rock flasks to accelerate trypsinization of cells. 27. Prior to begin cell culture work, put trypsin and complete MEM 37°C water bath to warm up solutions. When ready to start cell culture, trypsin and MEM bottles are wiped with 70% (v/v) ethanol and put in level 2-certified biological safety cabinet (BSC). Trypsin and MEM can be left at room temperature in BSC during cell culture procedures. 28. Dishes can also be seeded at a cell density up to 2.5 × 106 cells/ dish. If you observed that cells seeded at 2.5 × 106 cells/dish grow too fast at high number of cell passages (>P48), we recommend that dishes be seeded at a lower cell density (2 × 106 cells). In our experience, cells transfected at a density ranging from 2 to 2.5 × 106 cells does not significantly impact the D1-like receptor expression. However, our unpublished data suggest that when cells are seeded at higher cell density than 2.5 × 106 cells/dish (>3 × 106 cells/dish), there is a greater variability in receptor expression. Experimenter should bear in mind that the aforementioned guidelines are those that have been optimal for our laboratory. 29. For radioligand binding studies, we typically transfect each dish of cells with 5 mg of plasmid DNA. In our hands, this amount of DNA yields in transfected HEK293 cells the maximal achievable receptor expression as measured with [3H]-SCH23390 (~15–20 pmol/mg membrane proteins).

176

B. Plouffe and M. Tiberi

Indeed, the use of higher amounts of DNA (10–20 mg/dish) does not lead to greater expression levels of wild type (WT) or low-expressing mutant forms of hD1R or hD5R. With respect to performing dose–response curves in intact cells, we titrate the amount of receptor expression construct DNA to be used in transfection to achieve lower levels of receptor (1–3 pmol/mg membrane proteins). For instance, the amounts of receptor expression construct DNA used herein for dose–response curves are as follows: hD1R-WT (0.04–0.07 mg/dish), hD1RS263A (0.04 mg/dish), hD1R-S263G (0.08–12 mg/dish), hD1R-S263D (0.06–0.08 mg/dish), hD5R-WT (0.04– 0.08 mg/dish), hD5R-S287A (0.06 mg/dish), hD5R-S287G (0.06–0.08 mg/dish), and hD5R-S287D (0.08 mg/dish). Importantly, when using lower amount of 5 mg/dish of receptor expression construct DNAs, the total amount of plasmid DNA must be normalized at a constant amount per transfected dish (5 mg) using empty plasmid (e.g., pCMV5). This will mitigate variations in transfection efficiency between different conditions. The efficiency in HEK293 cells obtained using our transfection method will not be covered here as this issue has been previously discussed elsewhere (17, 19). 30. Remove drips left on side of tubes by gentle tapping on a hard surface inside the BSC. 31. The pH of 2× HEPES-buffered saline solution (pH to 7.1 ± 0.05) is critical for optimal transfection of plasmid DNA. Higher and lower pH will significantly impact the formation DNAcalcium phosphate precipitates. For instance, we observed a drastic reduction in receptor expression with a solution at pH of 7.3. 32. We observe that the proliferation rate of HEK293 cells slowly get higher up to 52 passages under our cell culture conditions. After 52 passages, HEK293 cells grow more rapidly and sometimes as foci. In addition, cells adhere less on dishes at older passages. In our laboratory, we generally utilize HEK293 cells between 40 and 52 passages. 33. Typically, we use four transfection dishes per experimental condition in radioligand binding and whole cell cAMP assays. Meanwhile, the number of transfected dishes can be scaled up if mutant receptors display low expression levels in transfected HEK293 cells (<1 pmol/mg membrane proteins) or if larger amounts of membrane preparations are needed for radioligand binding assays. 34. For radioligand binding studies, we normally let transfected HEK293 cells grow for 48 h following seeding in 150 × 25 mm dishes. For instance, if the transfection is performed on Tuesday, cells are split and seeded in new dishes on Wednesday and then

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

177

used to make membrane preparations on Friday. Alternatively, the experimenter can use cells on Thursday (cells grown for 24 h) as we observe that expression is high enough to detect receptor binding. 35. Cells in 100 × 20 mm dishes are used to prepare crude membranes to determine receptor levels for each transfection condition using radioligand binding. 36. In general, 4 rows of 12 holes are used to set up two saturation curves with six concentrations of [3H]-SCH23390 using duplicate determinations for total and nonspecific binding (24 tubes for each receptor tested) for a total of 48 tubes per rack. This set up is optimal for terminating reactions using our Brandel harvesting system. For the purpose of our study, four racks were used to perform saturation studies with wild type and single-point mutants of hD1R and hD5R. A set of four tubes is used for each concentration of [3H]-SCH23390 (two tubes for total binding followed by two tubes for nonspecific binding). 37. We usually carry out assays with 25–100 mL of membrane resuspension in a final volume of 800 mL of Milli-Q water in glass test tubes (10 × 75 mm) and 200 mL of protein dye. Mix well by vortexing several times. 38. Bmax value is used as an index of total receptor levels. 39. Typically, one rack (4 rows of 12 holes) allows performing two competition curves. For the purpose of the present study, each receptor construct to be tested requires two racks of 48 tubes to test four dopaminergic compounds (SCH23390, dopamine, cis-flupenthixol, (+)-butaclamol). Each competition curve is done in the absence (total binding) and presence of 11 increasing concentrations of cold drugs using duplicate determinations (24 tubes for each drug tested). 40. Volume of resuspension can be reduced to increase the specific binding signal if receptor constructs are expressed at a Bmax lower than 1 pmol/mg of membrane proteins. If so, the number of drugs to be tested will be reduced accordingly. Alternatively, to test four drugs in the same experiment, one can prepare a larger number of frozen stocks of membrane preparations of the given receptor expressed at low Bmax. 41. Competition curves are performed using a fixed concentration of [3H]-SCH23390 similar to Kd values measured at wild type and mutant receptors. 42. Competition studies and dose–response curves with dopamine are done in the presence of ascorbic acid at a final concentration of 0.1 mM in the assays to prevent dopamine oxidation.

178

B. Plouffe and M. Tiberi

43. Values obtained from dose–response curves remain essentially unchanged whether HEK923 cells are metabolically labeled with 1 or 2 mCi/mL of [3H]-adenine. Therefore, the use of 1 mCi/mL of [3H]-adenine will considerably decrease the amount of handled radioactivity and cost of experiments. Make also a note that if cells are not labeled on the following day of seeding, cells can be labeled the same day of experiment using an amount of ³2 mCi/mL of [3H]-adenine to increase signal detection. Under these circumstances, cells are metabolically labeled for a minimum of 4 h prior to doing whole cell cAMP assays. 44. These samples are used to assess the amount of [3H]-adenine in labeling media. Vials can be put aside until counting the other vials with radiolabeled eluates obtained from double column chromatography. 45. Cells can also be incubated with drugs for a shorter time period. 46. Samples can be stored at 4°C up to 5 days. In our hands, no differences have been measured between samples processed immediately for double chromatography columns and samples processed after being left at 4°C for 5 days. Samples should be frozen at −20°C if longer time of storage is required. However, experimenter should first verify that the used test tubes do not crack −20°C. 47. The average [14C]-cAMP counts from triplicate samples (column input) will be computed to determine the column recovery and establish a correction factor (CF) for each set of paired Dowex and alumina columns. The column input (C14total) is calculated from counting the radioactivity in vials containing 0.85 mL of KOH-neutralized cAMP stop solution mixed with 2 mL of 0.1 M imidazole (to make a 3 mL aqueous sample) and 18 mL of scintillation cocktail. 48. Samples are clarified using a low-speed centrifugation to pellet down potassium perchlorate salts prior to applying samples on Dowex columns. Loading salts on Dowex columns will interfere with sample elution and lead potentially to spurious radioactive counts. 49. We choose to apply a volume of 0.85 mL on Dowex columns to prevent touching and pipetting salt precipitates from the bottom of test tubes. 50. While the volumes for washes and elution of samples on Dowex and alumina columns reported herein are those routinely obtained in our laboratory, it is strongly recommended that the column profiles be thoroughly reassessed in the experimenter’s laboratory as described (19).

10

Site-Directed Mutagenesis of D1 and D5 Dopaminergic Receptors

179

51. [3H] counts measured in 50 mL aliquots will be used to determine the total amount of [3H]-adenine uptake (TU) in each well (an index of cell number). 52. The assessment of constitutive activity of wild type and mutant forms of hD1R and hD5R is beyond the scope of this study. The determination of constitutive activity of wild type and mutant forms of hD1R and hD5R is typically performed with cells seeded in 6-well plates and incubated in labeling medium containing 2 mCi/mL of [3H]-adenine overnight at 37°C in humidified 5% CO2 incubator. Generally, cells used for these assays express receptors at higher Bmax values. Methods pertaining to the study of constitutive activity of D1-like receptors and their mutants have been discussed elsewhere (19).

Acknowledgments We would to thank Dr. Kursad Turksen for his advice and Andrew Charrette for reading the manuscript. Bianca Plouffe was a recipient of a graduate scholarship from Fonds de la recherche en santé du Québec. This work was supported by an operating grant from Canadian Institutes of Health Research (MOP-81341). References 1. Arinaminpathy Y, Khurana E, Engelman DM, Gerstein MB (2009) Computational analysis of membrane proteins: the largest class of drug targets. Drug Discov Today 14:1130–1135 2. Hubert P, Sawma P, Duneau JP, Khao J, Henin J, Bagnard D, Sturgis J (2010) Single-spanning transmembrane domains in cell growth and cell-cell interactions: more than meets the eye? Cell Adh Migr 4:313–324 3. Rosenbaum DM, Rasmussen SG, Kobilka BK (2009) The structure and function of G-proteincoupled receptors. Nature 459:356–363 4. Fredriksson R, Lagerstrom MC, Lundin LG, Schioth HB (2003) The G-protein-coupled receptors in the human genome form five main families. Phylogenetic analysis, paralogon groups, and fingerprints. Mol Pharmacol 63:1256–1272 5. Pineyro G (2009) Membrane signalling complexes: implications for development of functionally selective ligands modulating heptahelical receptor signalling. Cell Signal 21:179–185 6. Ritter SL, Hall RA (2009) Fine-tuning of GPCR activity by receptor-interacting proteins. Nat Rev Mol Cell Biol 10:819–830

7. Davey J (2004) G-protein-coupled receptors: new approaches to maximise the impact of GPCRS in drug discovery. Expert Opin Ther Targets 8:165–170 8. Yildirim MA, Goh KI, Cusick ME, Barabasi AL, Vidal M (2007) Drug-target network. Nat Biotechnol 25:1119–1126 9. Tiberi M, Caron MG (1994) High agonistindependent activity is a distinguishing feature of the dopamine D1B receptor subtype. J Biol Chem 269:27925–27931 10. Missale C, Nash SR, Robinson SW, Jaber M, Caron MG (1998) Dopamine receptors: from structure to function. Physiol Rev 78:189–225 11. Beaulieu JM, Gainetdinov RR (2011) The physiology, signaling, and pharmacology of dopamine receptors. Pharmacol Rev 63: 182–217 12. Cotecchia S (2007) Constitutive activity and inverse agonism at the alpha(1)adrenoceptors. Biochem Pharmacol 73:1076–1083 13. Jarvie KR, Caron MG (1993) Heterogeneity of dopamine receptors. Adv Neurol 60:325–333 14. Charpentier S, Jarvie KR, Severynse DM, Caron MG, Tiberi M (1996) Silencing of the

180

B. Plouffe and M. Tiberi

constitutive activity of the dopamine D1B receptor. Reciprocal mutations between D1 receptor subtypes delineate residues underlying activation properties. J Biol Chem 271:28071–28076 15. Jackson A, Iwasiow RM, Tiberi M (2000) Distinct function of the cytoplasmic tail in human D1-like receptor ligand binding and coupling. FEBS Lett 470:183–188 16. Demchyshyn LL, McConkey F, Niznik HB (2000) Dopamine D5 receptor agonist high affinity and constitutive activity profile conferred by carboxyl-terminal tail sequence. J Biol Chem 275:23446–23455 17. Tumova K, Iwasiow RM, Tiberi M (2003) Insight into the mechanism of dopamine D1-like receptor activation. Evidence for a

molecular interplay between the third extracellular loop and the cytoplasmic tail. J Biol Chem 278:8146–8153 18. Tumova K, Zhang D, Tiberi M (2004) Role of the fourth intracellular loop of D1-like dopaminergic receptors in conferring subtype-specific signaling properties. FEBS Lett 576:461–467 19. Plouffe B, D’Aoust JP, Laquerre V, Liang B, Tiberi M (2010) Probing the constitutive activity among dopamine D1 and D5 receptors and their mutants. Methods Enzymol 484:295–328 20. D’Aoust JP, Tiberi M (2010) Role of the extracellular amino terminus and first membrane-spanning helix of dopamine D1 and D5 receptors in shaping ligand selectivity and efficacy. Cell Signal 22:106–116

Chapter 11 A Molecular Genetic Approach to Uncovering the Differential Functions of Dopamine D2 Receptor Isoforms Yanyan Wang, Toshikuni Sasaoka, and Mai T. Dang Abstract Alterations in the activity of the dopamine D2 receptor (D2R) have been implicated in several neurological and psychiatric disorders, including schizophrenia, Parkinson’s disease, Huntington’s disease, Tourette syndrome, attention-deficit hyperactivity disorder (ADHD), and drug addiction. Two isoforms of D2R, long form (D2LR) and short form (D2SR), have been identified. The specific function of each D2R isoform is poorly understood, primarily because isoform-selective pharmacological agents are not available. Using homologous recombination, we have generated D2LR knockout (KO) mice. D2LR KO mice are completely deficient in D2LR, but still express functional D2SR at a level similar to the total D2R level in wild-type (WT) mice. D2LR is generally the predominant isoform expressed in WT mice. We showed that D2LR KO mice displayed a number of robust behavioral phenotypes distinct from WT mice, indicating that D2LR and D2SR have differential functions. In this chapter we describe the generation and characterization of the D2LR KO mouse. This genetic approach provides a valuable research tool to investigate the functional role of individual D2R isoforms in the mammalian central nervous system (CNS). Key words: Dopamine D2 receptor, Knockout mouse, Homologous recombination, Transfection of embryonic stem cell, Genotyping

1. Introduction Dopamine (DA) receptors are classified into two categories: D1-like receptors (D1 and D5 subtypes) and D2-like receptors (D2, D3, and D4 subtypes). Additionally, two isoforms of dopamine D2 receptor (D2R) have been identified by molecular cloning in the brain of rats, mice, and humans and are termed as the long receptor form (D2LR) and short receptor form (D2SR) (1–4). D2LR and D2SR are generated by alternative splicing of the same gene

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_11, © Springer Science+Business Media, LLC 2013

181

182

Y. Wang et al.

Fig.1. Schematic illustration of the gene and mRNA structures of D2LR and D2SR. Open and filled boxes on gene structure represent exons, and lines between boxes represent introns. The Exon 6 (filled box) is 87 bp that encodes the D2LR insert region of 29 amino acids in the third intracellular loop.

(Fig.1). No selective pharmacological ligands are available for these two types of D2R, which has hampered the progress towards uncovering their specific function in the CNS. To overcome this challenge, we decided to generate mutant mice expressing only a single D2R isoform using gene-targeting technology. We made a line of mice that lack D2LR (D2LR KO mice) by selectively deleting exon 6 in the D2 gene (5). We previously showed that D2LR KO mice still express a functional D2SR isoform on the cell surface at a level similar to the native level of the total D2R in wild-type (WT) mice(5). The mice that express only D2SR allow us to examine the biological responses mediated by D2SR. Because D2LR is the predominant isoform expressed in the brain of WT mice (5, 6), the comparative studies between D2LR KO and WT mice should help elucidate the functional role of D2LR (5–8). Making a knockout mouse involves five major steps: (1) making a gene-targeting vector, (2) transfecting the targeting vector into embryonic stem (ES) cells and screening targeted ES clones by homologous recombination, (3) injecting the targeted ES cells into blastocysts and returning the blastocysts to the foster mother to generate chimeric mice, (4) breeding chimeric mice with WT mice to generate heterozygous mice, and (5) genotyping to identify germline transmitted mice and intercrossing heterozygotes to yield homozygous mice.

2. Materials 2.1. Making the Targeting Vector

1. Mouse genomic DNA library can be obtained from Agilent Technologies, Inc. (Santa Clara, CA). 2. PGK-neo cassette can be obtained from: Gene Bridges GmbH, Addgene (Cambridge, MA). 3. Cloning materials: Escherichia coli, pBluescript, Quick Ligation Kit (New England Biolabs, Ipswich, MA), an assortment of

11

Functions of D2 Receptor Isoforms

183

restriction enzymes (New England Biolabs or Invitrogen Life Science, Grand Island, NY). 4. Mini-plasmid preparation material: mini-prep solution, agarose electrophoresis gel. 5. LB solution containing antibiotic. 6. Alkaline lysis solution I: 50 mM glucose, 25 mM Tris–HCl (pH 8.0), 10 mM EDTA. 7. Alkaline lysis solution II: 0.2 M NaOH, 0.1% SDS, prepared fresh. 8. Alkaline lysis solution III: 3 M potassium, 5 M acetate. 9. Isopropanol. 10. Isobutanol. 11. Cesium chloride. 12. Ethidium bromide. 13. 1 M Tris–HCl (pH 8.0). 14. 1 M Tris–HCl (pH 8.5). 15. 0.5 M EDTA (pH 8.0). 16. 20× SCC. 17. 50× TAE (Tris–acetate–EDTA). 18. TE (pH 8.0) (10 mM Tris–HCl—1 mM EDTA). 19. 10% SDS. 20. Ethidium bromide (10 mg/mL). 21. 100× Denhardt’s solution. 22. 5 M NaCl. 23. 1 M CaCl2. 24. 1 M potassium acetate. 25. 3 M sodium acetate. 26. Tris–EDTA buffer. 27. LB solution. See “Molecular Cloning” (9) regarding how to make various buffers (see Note 1). Pre-mixed buffers are also commercially available. 2.2. Transfection of Embryonic Stem Cells

1. Embryonic stem (ES) cells: J1 ES cell established from 129 S4/SvJae provided by Dr. En Li (Novartis Institute for Biomedical Research). 2. Embryonic fibroblast (EF) cells, used to provide nourishment to ES cells (American Type Culture Collection, Manassas, VA). 3. ES medium (500 mL): 75 mL fetal calf serum (FCS), 2.5 mL 100× penicillin–streptomycin (Gibco, Invitrogen), 5 mL 100×

184

Y. Wang et al.

glutamine (Gibco, Invitrogen), 5 mL 100× nonessential amino acids (Gibco, Invitrogen), 5 mL 100× nucleosides, 5 mL 100× 2-mercaptoethanol, 400 mL DMEM; filter with 0.2 μm filter to sterilize and store at 4°. 4. 100× Nucleosides: 80 mg adenosine, 85 mg guanosine, 73 mg cytidine, 73 mg uridine, 24 mg thymidine, 100 mL Milli-Q H2O (mQH2O); dissolve for 15 min at 65°, filter (0.2 μm), aliquot and store at −20°. 5. 100× 2-mercaptoethanol (Sigma-Aldrich, St. Louis, MO): 70 μL of 2-mercaptoethanol in 100 mL PBS, filter (0.2 μm). This solution needs to be made every 2 weeks. 6. EF medium (500 mL): 50 mL FCS, 5 mL 100× penicillin– streptomycin, 5 mL 100× glutamine, 450 mL DMEM with HEPES, filter (0.2 μm) and store at 4°. 7. Plates and flasks for EF cells; treat the container with 0.1% gelatin solution for 20 min at room temperature and then aspirate the gelatin. 8. Trypsin (Gibco, Invitrogen), Geneticin (G418) (Gibco, Invitrogen), Leukemia inhibitory factor (LIF, BRL; ESGRO (mLIF), Invitrogen). 9. 2× Freezing medium: 80% FCS with 20% DMSO, filter to sterilize (0.2 μm) and store at 4°. 10. Electroporation apparatus (Bio-Rad Gene Pulser, Bio-Rad, Hercules, CA). 2.3. Injection of ES Cells into Blastocysts and Transfer of Blastocysts to the Uterus

1. Female C57BL/6 donor mice (2- to 3-month-old) (The Jackson Laboratory, Bar Harbor, ME).

2.3.1. Breeders for Harvesting Blastocysts and Foster Mother Mice

4. Vasectomized male mice (3- to 12-month-old, B6CBAF1 or ICR).

2. C57BL/6 male breeder mice (3- to 6-month-old, should be less than 1 year old) (Jackson). 3. Female foster mice (2- to 3-month-old, B6CBAF1 or ICR).

5. ES medium. 6. Hand-pulled pipette and bulb or tubing with mouth piece. 7. Sterile surgical tools: scissors and forceps. 8. Pregnant mare’s serum gonadotropin (PMSG) (SigmaAldrich), Human chorionic gonadotropin (HCG) (SigmaAldrich), Phosphate-buffered saline (PBS).

2.3.2. Blastocysts Injection

1. ES medium. 2. Small culture plates. 3. Injection and holding pipettes. 4. Microinjector apparatus (Leica Microsystems, Wetzlar, Germany).

11 2.3.3. Surgery to Return Injected Blastocysts to Foster Mother

Functions of D2 Receptor Isoforms

185

1. Sterile surgical tools: scissors (1 serrated for cutting through pelt and 1 fine for body wall), forceps, 2 Serafin clips, surgical wound clips. Hand-pulled pipette for transferring blastocyst. 2. Pentobarbital sodium (Somnopentyl injection, ScheringPlough Animal Health Corp., Union, NJ). 3. Needle and syringe for injection. 70% ethanol. 4. Heating pad.

2.4. Genotyping the Mice

1. Lysis buffer (500 mL): 50 mL of 1 M Tris–HCl (pH 8.5), 5 mL of 0.5 M EDTA (pH 8.0), 10 mL of 10% SDS, 20 mL of 5 M NaCl, 0.5 mL of 1 M CaCl2, and bring to a final volume of 500 mL with mQ H2O or equivalent. 2. Proteinase K Invitrogen).

(20

mg/mL)

(New

England

Biolabs,

3. dNTP (QIAGEN, Valencia, CA). 4. Mineral oil (Sigma). 5. AmpliTaq DNA Polymerase Stoffel Fragment (Applied Biosystems Life Technologies). 6. PCR Thermo Cycler (MJ Research, Inc./ Bio-Rad). PCR tubes (USA Scientific, Ocala, FL). 7. Agarose gel electrophoresis apparatus (Owl Scientific, Thermo Fisher, Rockford, IL). 8. Agarose (Invitrogen) 9. 1× TAE buffer. 10. Ethidium bromide (Sigma). 11. Ultraviolet transilluminator and camera (or computer) for taking the gel picture.

3. Methods 3.1. Making the Targeting Vector for D2 Long Receptor Knockout

Gene-targeting relies on homologous recombination, which is done by a cellular endogenous recombinase that recognizes the similarity between the DNA sequence fragments of the targeting vector (or construct) and that of the native gene. Recombination occurs at the sites of similarity, and the altered genetic material within those flanking sites is incorporated into the native gene. The construct design determines the type of knockout mouse. The simplest construct for producing a knockout is to replace a coding region of the gene of interest with an antibiotic resistant gene. The removal of a coding region will interfere with transcription and protein expression, disrupting the production of the protein or resulting in the translation of an unstable protein. Recombination efficiency

186

Y. Wang et al.

is sensitive to the homologous sequence. Generally, the size of the deleted segment of the gene should be close to the size of the insertion between the left homologous arm and the right homologous arm of the targeting vector, although there are successful exceptions (10). The antibiotic resistant gene most commonly used is the neomycin (neo) resistant gene driven by a ubiquitous 3-phosphoglycerate kinase (PGK) promoter, together referred to as a PGK-neo cassette (11). The PGK-neo cassette is used as a positive selection marker for homologous recombination in ES cells. Here we provide mainly general guidelines in designing and making a gene-targeting vector because this process is unique to the gene of interest. Because D2LR and D2SR are generated from the same gene by alternatively splicing, it is important to not disrupt transcription of the D2 gene; otherwise you would produce a complete D2R KO mouse. D2LR has a 29 amino acid insertion located within the third intracellular loop, which is encoded by exon 6; this insertion is absent in D2SR (Fig. 1). To generate the mice deficient in D2LR, we selectively replaced exon 6 with the PGKneo cassette. In the D2LR KO mouse, D2SR is over-expressed to a level similar to that of total D2R in WT mice. This is because our manipulation of the D2 gene eliminated the possibility of transcription of D2LR, and therefore the D2 gene was exclusively available for making D2SR. 1. Find the genomic sequence from: www.ncbi.nlm.nih.gov/ genome/guide/mouse/ 2. The mouse D2 gene containing exons 3–8 was cloned from a mouse 129/SV genomic DNA library (see Note 2). Subclone the gene into pBluescript in E. coli. 3. To verify the genomic clones, map some of the major restriction enzyme sites on the exons and introns according to the restriction map and exon/intron structure of the D2 gene obtained from the database. This mapping will also identify the orientation of the fragment. Sequence some important gene regions (see Note 3). For example, we sequenced exon 6 and the exon/intron junctions between exon 6/intron 5 and between exon 6/intron 6 on the D2 gene. 4. Replace exon 6 of the D2 gene fragment with a PGK-neo cassette (Fig. 2a). The left homology arm of this construct is 7 kb and the right homology arm is 8.5 kb, which flank the 1.8 kb PGK-neo cassette (see Note 4). 5. While making the targeting vector, you should also design and make probes for screening homologous recombination in ES cells via Southern blot analysis. Usually the sequence of the probe for Southern blot analysis should be outside of the targeting vector (Fig. 2a). The probe should be able to distinguish homologous recombination from non-homologous recombination.

11

Functions of D2 Receptor Isoforms

187

It is better to prepare two probes: one is external to the 5¢ end of the left homology arm and the other is external to the 3¢ end of the right homology arm. 6. Verify the accuracy of the targeting vector against the design with restriction mapping, nucleotide sequencing, and Southern blot analysis. 7. Produce a large-scale plasmid preparation of the finished targeting vector with cesium chloride. Aliquot the targeting vector DNA into multiple tubes and store them at −20°. 8. For standard molecular biology protocols such as cloning, plasmid DNA preparation, and Southern blot analysis, see ref. 9, 12. Miniprep Kit and Plasmid Maxi Kit for purifying DNA are also commercially available (QIAGEN). 3.2. Transfection of Embryonic Stem Cells 3.2.1. Isolation of Embryonic Fibroblasts from Mice

1. Set up timed mating between a male mouse carrying a neomycin-resistant gene and several females. 2. Remove the uterus with the fetuses and put in a petri dish (this can be done on your working bench). Aim to isolate about 5–10 of 13- to 16-day-old fetuses (see Note 5). 3. Dissect out the fetuses in the clean hood and wash 5–7 times with 50 mL PBS. 4. Cut off the heads. Remove and discard the inside organs from thorax and abdomen. 5. Wash the tissue 5 times with PBS. 6. Cut up as finely as possible with sharp scissors. 7. Add 5 mL of trypsin solution (0.25% trypsin, 1 mM EDTA, Gibco), incubate for 10 min at 37° with gentle shaking and then pipet vigorously. 8. Add 30 mL of EF medium, allow the debris to settle, and transfer the supernatant to a new tube. 9. Re-trypsinize the debris, collect the supernatant, and combine the collected supernatants. 10. Spin down the cells for 5 min at 239 × g. Resuspend the pellet in 100 mL of EF medium. 11. Plate onto three T175 flasks. This plating is considered to be passage zero (P = 0). 12. Grow to confluent culture (1–4 days) and split into ten T175 flasks (P = 1). Add G418 (150 μg/mL) to select for neomycin resistant clones and grow to confluent culture (2–3 days). 13. Split into 30 T175 flasks (P = 2) and continue the selection process with G418. Continue growing to confluent culture. 14. Trypsinize the EF cells and freeze cells from one T175 flask per tube (about 3 × 107 cells).

188

Y. Wang et al.

Fig.2. Targeted disruption of the D2L gene. (a) Targeting vector. For gene targeting of the D2L locus, exon 6 (filled box, enlarged for visualization) was replaced by a PGK-neo cassette, which was flanked by loxP (arrowhead). H, HindIII; K, KpnI; Sc, ScaI. (b) Southern blot analysis of genomic DNA from mouse tails (−/−: homozygote; +/−: heterozygote; +/+: wild-type). Tail DNA was either digested with KpnI and hybridized with the external probe A or digested with ScaI and hybridized with the external probe B. (c) Detection of two forms of D2 mRNA by RT-PCR. PCR products, differing by 87 bp and corresponding to the alternatively spliced isoforms of D2 mRNAs, were shown in lanes 2–4 (+/+ mice: 235 bp—D2S and 322 bp—D2L) and in lanes 5–7 (−/− mice). The size marker (M) was a 123 bp DNA ladder (lane 1). (d) Northern blot analysis of D2 RNA levels in striatum of +/+ (lane 1) and −/− (lane 2) mice. The probe used was a DNA fragment complementary to exon 7 of the D2 gene and thus hybridized to RNA from both WT and D2L−/− mice. Total striatal RNA was obtained from 3 to 4 mice for each genotype. The data were quantified with densitometric analysis using a scanner (UMAX Astra 2400S) and IQ Mae program. Note that D2 mRNA from mutant mice was present at the predicted smaller size (similar to D2S mRNA size). The size marker indicated by the arrows was Millennium RNA size marker (Ambion). β-Actin RNA levels were simultaneously monitored as an internal control. (This figure is reproduced from ref. 5 with permission).

11

Functions of D2 Receptor Isoforms

189

15. Grow a small amount of the cells to high confluent culture in antibiotic-free medium and check for mycoplasma contamination using the mycoplasma detection kit from Gen-Probe, Inc. Fresh medium can be used as a negative control. 3.2.2. Expanding Embryonic Fibroblasts

1. Day 0: thaw 1 tube of 3 × 107 EF cells (or feeders) at P = 2 and plate onto three T175 flasks (use about 30 mL of EF medium per flask). 2. Day 4: split the cells and plate onto ten T175 flasks. To split the cells, wash the cells with PBS, add 2 mL of trypsin, 5 min 37°, pipet up and down, and resuspend in EF medium. 3. Day 7: inactivate the cells by mitomycin C (mit C) treatment (see Note 6): (a) Dissolve 1 vial of mit C (2 mg/vial, Sigma) in 10 mL of PBS (20× solutions). (b) Feed with EF medium containing 1× mit C for 2 h at 37°. (c) Aspirate the medium and wash 3 times using PBS. (d) Trypsinize the cells and freeze at 3 × 107 cells/mL (10 tubes of 1 mL).

3.2.3. Plating of Embryonic Fibroblasts

1. Treat the plates with a 0.1% gelatin solution for at least 20 min at room temperature, and aspirate the gelatin solution. 2. Thaw a frozen stock of mit C-treated EF cells at 37°, add 10 mL EF medium, spin, and resuspend. One vial of mit C-treated EF (3 × 107 cells) should be resuspended in 30 mL of EF medium. 3. Plate EF as follows: 500 μL per 1 well of 24-well plate, 200 μL per 1 well of 96-well, 5 mL for a T25 flask, 30 mL for a T175 flask, and 9 mL for a 10 cm petri dish. 4. Wait at least 4 h to plate ES cells; do not wait longer than 3 days.

3.2.4. Plating of Embryonic Stem Cells (13)

1. Prepare a flask with mit C-treated EF feeders the day before plating the ES cells. Feed the cells with ES medium + LIF just before plating. 2. Thaw ES cells at 37°, add 10 mL of ES medium, spin, and resuspend in ES medium. About 3 × 106 cells should be plated onto a T25 flask. The cells should reach 50% confluent after 2–3 days (see Note 7). 3. ES cells are split about 1:7 (e.g., a T25 flask can be split into a T175 flask).

190

Y. Wang et al.

3.2.5. Electroporation Conditions

1. The day before electroporation, prepare ten 10 cm petri dishes with EF. 2. Linearize approximately 50 μg of the targeting vector, ethanol precipitated, wash 2 times with 70% ethanol, dry in the hood, and further dry for 30 min at 65° in a heating block with the tube cap closed. 3. Resuspend the DNA in 600 μL warm PBS. 4. Trypsinize about 5 × 107 ES cells (about 1 × 50% confluent T175), resuspend in DNA solution to a total volume of 800 μL. 5. Add the mixture to a cuvette (0.4 cm gap, #165-2088, BioRed Gene Pulser). 6. Electroporate at 3.0 μF, 800 V (Bio-Rad). These cells are considered to be at day 0. 7. Resuspend the electroporated mixture in 100 mL ES medium + LIF and plate onto ten 10 cm plates with about 5 × 106 EF previously plated. Do not add G418 to the medium on day 0. 8. On the next day, begin the selection with the antibiotic. Add 150 μg G418/mL to five plates and 125 μg G418/mL to the other five plates (see Note 8). 9. Keep changing the medium with ES medium + G418 every day (see Note 9). 10. Start picking antibiotic-resistant colonies around day 7 or 8.

3.2.6. Picking of Colonies (14)

1. The day before picking the colonies, prepare several 96-well flat-bottom plates with EF. Change the medium with ES medium + LIF before picking. 2. Pick colonies under the microscope with an Eppendorf pipetman (keep the volume smaller than 5 μL) (see Note 10). 3. Transfer the clones to 96-well U-bottom plates. Add 15 μL trypsin and incubate for 5 min at 37°. 4. Add 35 μL ES medium (use medium from the 96-well plates with EF), pipet 10 times up and down and transfer to the 96-well flat-bottom plate with EF. 5. Change the medium every day. 6. Two to three days after picking, the ES cells should be ready to split into two series of 24-well plates; one set of plates will be used for freezing, the other will be used for DNA preparation for Southern blot analysis. The plates to hold ES cells for freezing should contain EF feeders. The plates for DNA preparation do not need feeders. All plates should be gelatinized. 7. Passage method: (a) Aspirate the medium, add 50 μL trypsin, and incubate for 5 min at 37°; (b) Add 150 μL of ES medium, pipet 10 times up and down; (c) Transfer 100 μL to the 24-well

11

Functions of D2 Receptor Isoforms

191

plate without feeders and 100 μL to the 24-well plate with feeders; (d) Change the medium every day. The cells for DNA preparation can be fed with ES medium without LIF. 3.2.7. Freezing of Clones (Method 1) (See Note 11)

1. Aspirate the medium from the 24-well plates. 2. Add 1 mL of ice-cold 1× freezing medium (90% FCS + 10% DMSO, filtered (0.2 μm) and stored at 4°). 3. Seal the plate with Parafilm and put in a styrofoam box to store at −70° (see Note 12).

3.2.8. Screening of ES Cell Clones with Homologous Recombination (See Note 13)

1. For preparation of genomic DNA, aspirate the medium. Add 500 μL of lysis buffer (100 mM Tris–HCl pH 8.5, 5 mM EDTA, 0.2% SDS, 200 mM NaCl, 100 μg/mL Proteinase K); Shake overnight at 56°. 2. Put the plate on a swirling table for 15 min, add 1 volume of isopropanol, and keep swirling until the DNA has aggregated. This usually takes about 15 min. 3. Recover the DNA by lifting the aggregated precipitate from the solution using an Eppendorf tip, remove excess liquid and transfer to a pre-labeled microcentrifuge tube. 4. Spin briefly to collect the pellet at the bottom of the tube. 5. Dry for 5 min in the Speed Vac evaporator or for 1 h in a 37° warm room. 6. Add 50 μL of TE-buffer or sterile H2O and dissolve overnight at 56°. 7. Store at 4° or −20°. 8. To digest the DNA (see Note 14), prepare 10 μL DNA, 2.5 μL 10× restriction buffer, 2.5 μL 0.1 M 2-mercaptoethanol, 2.5 μL restriction enzyme, 7.5 μL mQH2O (total volume: 25 μL). 9. Mix well and digest overnight at 37°.

3.2.9. Southern Blotting

1. Take a picture of the stained gel. Cut out the region of the gel where the bands are expected. 2. Shake the gel in 0.5 M NaOH + 1.5 M NaCl for 30 min. 3. Pre-wet the nylon filter (Zeta-Probe GT, Bio-Red) in water for 5 min. 4. Transfer the gel in 0.5 M NaOH + 1.5 M NaCl overnight. 5. Rinse the filter in 2× SSC. 6. Air dry the filter, put the gel between Whatmann 3MM paper and bake at 80° under vacuum for 30 min to 1 h.

3.2.10. Hybridization

1. Hybridization mix: 1% SDS, 6× SSC, 10% dextran sulfate, and at least 100 μg salmon sperm DNA/mL. Alternatively, you can

192

Y. Wang et al.

use commercially available hybridization solution (Clonetch, Stratagene/Agilent Technologies). 2. Pre-hybridize at least 4 h at 65°, add radioactive probe and hybridize overnight. 3. Wash the filter: 1 time for 15 min with 2× SSC + 1% SDS at room temperature, and then 4 times for 15 min with 0.1× SSC + 0.1% SDS at 65°. 4. Remove the liquid, wrap in plastic wrap, and expose to X-ray film with DNA side up. 3.3. Injection of ES Cells into Blastocysts for Production of Chimeric Mice 3.3.1. Breeding of Donor Mice and Harvesting of Blastocysts

1. On day 1 at 4:00 pm, intraperitoneally (IP) inject 4–10 female donor mice with PMSG (5 IU/mouse). 2. On day 3 at 1:00 pm, inject (IP) the female donor mice with HCG (5 IU/mouse), and immediately mate the donor mice with intact male breeders (one to one). 3. On the morning of day 4, check females for semen plug, separate plugged females into a new cage from non-plugged females and hold them until day 7. 4. On day 7, collect blastocysts from the female donor mice. Remove the uterus, cut the distal ends, and flush out blastocysts by expelling ES medium from a pipette into a dish. Shake the dish gently on a shaker for a few minutes. Under a microscope, collect the blastocysts and transfer to a new petri dish containing 50 μL of ES medium underneath the light mineral oil.

3.3.2. Breeding of Foster Mothers

1. In the evening of day 4, breed 6 foster mothers (see Note 15) with vasectomized male mice. 2. On the morning of day 5, check females for semen plug, separate plugged females into a new cage from non-plugged females, and hold them until day 7.

3.3.3. Thawing of Targeted ES Clone (See Note 16)

1. Plate EF feeders in gelatinized 24-well plates. 2. For Method 1 of freezing cells, thaw the targeted ES cells in the following way: (a) Take the plate out of the −70° freezer and put in the hood. (b) Add pre-warmed medium to the well and pipet up and down with a Pasteur pipette until everything is thawed. Make sure all the cells are detached from the bottom of the well. (c) Suspend cells in 10 mL of medium, spin, resuspend in ES medium, and transfer to two 24-wells. 3. Change the medium every day and grow to 50% confluent culture. The time to confluence usually takes 2–5 days, depending on the clone.

11

Functions of D2 Receptor Isoforms

193

4. Freeze the cells from 1 well. Split the cells from the other well to 2 wells on a 12-well plate and grow to 50% confluent. 5. Freeze 1 well, split the other to 2 wells on a 6-well plate. Use also some of the cells to startup 1 or 2 wells on a 24-well plate; these cells will be used to prepare DNA to verify the clone. 6. Freeze 1 well, and split the other well into two T25 flasks. 7. Freeze cells from the two flasks in ten tubes. 8. Prepare DNA from the clone and verify by Southern blotting. 9. Select correctly targeted ES clones for injection that (a) have a close to 1:1 ratio of wild-type: knockout band on the Southern blot, (b) look healthy and undifferentiated, and (c) grow fast. 3.3.4. Blastocysts Injection (13, 15)

1. Thaw one tube of frozen ES cells 2–3 days before injection and plate onto 3 wells of a 12-well plate containing ES medium. The plate should be gelatinized but without EF. 2. In the afternoon of the injection after blastocysts have been collected, trypsinize ES cells, gently manually dissociate, spin down, resuspend in fresh ES medium, and maintain them as a single cell suspension on ice. 3. Pre-cool the injection chamber on the microscope stage. 4. Use a transfer pipette to transfer the expanded blastocysts in groups of about ten into the pre-cooled injection chamber. Then, introduce a few hundred ES cells (a “cloud” of cells, filling, at most, half the drop) into the injection chamber and allow them to settle on the bottom. 5. Using high-power magnification, select individual ES cells carefully on the basis of size (small, compared to the feeder cells) and shape (uniformly round, compared with more ragged or “rough” feeder cells). Draw about a hundred ES cells into the injection pipette and position them near the tip in a minimal amount of medium. 6. Hold a single blastocyst by applying suction to the holding pipette and move it toward the center of the microscope field. Position the embryo with the inner cell mass (ICM) at either the 6 or 12 o’clock position. 7. Adjust the focus precisely at the equator of the blastocyst. Align the tip of the injection pipette in the same focal plane as the equator of the blastocyst. 8. With a continuous movement, introduce the loaded injection pipette into the blastocyst cavity. Aim to insert the injection needle at a junction between two trophoblast cells. When a half of the needle of the injection pipette enters the blastocyst cavity, slowly expel medium into the cavity so as to expand the blastocyst and insert the pipette further into the cavity.

194

Y. Wang et al.

Take care not to touch the ICM with the injection needle. If the first attempt to penetrate the trophoblast layer is unsuccessful, and the blastocyst is not collapsed, try to insert the needle exactly at the same position again. When the blastocyst starts collapsing, expel a small amount of medium slowly to keep the blastocyst cavity expanded. 9. Slowly expel 8–15 cells into the blastocyst cavity. Take care not to insert any oil bubbles or lysed cells into the blastocyst. 10. Withdraw the injection pipette slowly. If the pressure is high inside the blastocyst cavity, keep a half of the tapered needle of the injection pipette inside the blastocyst to reduce the pressure. This is to prevent injected cells from being pushed out while the needle is being withdrawn. The blastocyst will collapse once the pipette is removed, resulting in the cells coming into close contact with the surface of the ICM. 11. Remove the injected blastocysts periodically and incubate them in a humidified incubator with 5% CO2 at 37° in microdrops of M16 or KSOM-AA medium (or ES medium). The blastocysts will re-expand after 1–3 h in culture (see Note 17). 3.3.5. Surgery to Return Injected Blastocysts to Foster Mother (13)

1. Anesthetize the foster mother mouse with pentobarbital sodium injected (IP) at a dose of 50–75 mg/kg. 2. When the foster mouse is fully anesthetized with no reflex response to pinching of paws, spray the lower back with 70% ethanol. 3. Make a horizontal incision in the flank approximately 0.5 cm away from the midline and between the natural hump of the back and the point where the rear leg joins the abdomen. 4. Remove hair by wiping the area again with ethanol. 5. Locate under the skin, the ovarian fat pad and gently nick the body wall (the peritoneum) overlying the pad. 6. Pull out the fat pad with the forceps and the uterus attached to it. Attach the serafin clips to the fat pad and lay the clips down across the animal’s body to stabilize the tissue mass. 7. Locate the uterine horns. 8. Take up 8–10 blastocysts and small air-bubbles as a marker of where the difficult-to-see blastocysts are in the pipette. 9. Insert a 26 G needle through the muscle layers of the uterus 2–3 mm distant from the utero-tubal junction. Remove the needle and keep your eye on the resulting hole in the uterine wall. 10. Insert the tip of the pipette into the lumen of the uterus through the hole. Transfer blastocysts and air-bubbles into the uterine.

11

Functions of D2 Receptor Isoforms

195

11. After removing the pipette, expel residual solution from the pipette and check under the microscope to ensure no blastocysts remained in the pipette. 12. Return the uterus and fat pad to the body cavity. 13. Repeat step 3–12 on the other side. 14. Close the wound with surgical clips. 15. Put the mouse on heating pad until it wakes up, and then return it to its home cage. 3.4. Breeding of Chimeric Mice for Determining Germline Contribution

1. Pups developed from chimeric fertilized eggs will have a mosaic coat color pattern. Mice containing a greater percentage of the ES cell strain color (i.e., agouti in the case of J1 ES cell A/A C/C) are the more useful progenies, since in these mice there is a greater likelihood that their germ cells differentiated from the targeted ES cells. In addition, since the majority of the established ES cell lines are of the XY karyotype, the gender ratio of the chimeras is usually biased towards males. 2. Mate male chimeras with mice of the strain identical to that of the host blastocysts (i.e., C57BL/6 females). Offspring of this breeding that have the color of the blastocyst host strain (i.e., non-agouti black color a/a C/C) do not carry the transgene. Pups with a homogenous hybrid color, such as agouti (A/a C/C in the case of J1 ES cell as a donor ES cell and C57BL/6 as a female strain) represent germline contribution of ES cells. Since one of two allele of the relevant gene is targeted in the ES cell, a half of ES cell-derived progeny has potential germline transmission. 3. Identify mice with germline transmission (i.e., heterozygous mice) by Southern hybridization (or blot) analysis as described in the next section. 4. Mate heterozygous males and females to yield homozygous offspring, and verify the pups genotype by Southern blot analysis (Fig. 2b.).

3.5. Genotyping to Identify Mice with Germline Transmission of the Targeted Allele

1. Cut 3–4 mm of tail and drop it into a pre-labeled 1.5 mL microcentrifuge tube.

3.5.1. Extract Genomic DNA from a Tail Tip

3. Incubate the tubes in the oven (54–56°) overnight. If possible, put tubes in a rotation rack (see Note 18) or in a foam box on a shaker to facilitate the lysis process (see Note 19).

2. Add 500 μL of lysis buffer with 2.5 μL of proteinase K into each tube containing the tail.

4. Put the tubes in a microcentrifuge and spin at 16,500 × g for 10 min. Upon completion, a pellet should be firmly aggregated at the bottom of the tubes. The DNA is in the supernatant.

196

Y. Wang et al.

5. Transfer by pouring the supernatant (about 450 μL) to a new 1.5 mL tube without disturbing the pellet. 6. Add equal volume of isopropanol (about 450 μL) to each tube containing the supernatant. Invert the tubes 10 or more times until the DNA aggregates. 7. Spin the tubes in a microcentrifuge for 5 min to collect the pellet (DNA) at the bottom of the tubes. Discard the supernatant by pouring without disturbing the pellet. 8. Add 1 mL of 70% ethanol to each tube containing the DNA and rinse briefly. 9. Spin the tubes in a microcentrifuge for 3–5 min to collect the pellet at the bottom of the tubes, and discard the ethanol by pouring without disturbing the pellet. Spin the tubes briefly again, and put the tubes with the lid open on Kim-Wipes (Kimberly-Clark Co.) in an inverted position to drain the residual ethanol. 10. Air dry the DNA for 5–10 min at room temperature. 11. Add 50 μL of TE buffer (pH 8.0), and put the tubes on a rotating rack or shaker in the oven (54–56°) overnight to dissolve the DNA (see Note 20). 12. After the DNA is dissolved, store it at 4°. 3.5.2. Southern Blot Analysis to Identify Pups with the Targeted Allele

1. Heterozygous (+/−), homozygous (−/−), and wild-type (+/+) mice should be determined by Southern hybridization analysis using appropriate DNA probes made during the construction of the targeting vector (Fig. 2b). 2. For Southern Hybridization method, see previous descriptions (Subheading 3.2).

3.5.3. RT-PCR and Northern Blot Analysis

1. To verify the D2L mRNA is deficient and determine the expression level of D2S mRNA, perform reverse transcriptase-PCR (RT-PCR) (Fig. 2c). Isolate total RNA from the striatum of D2LR KO or WT littermates (TRIzol Reagent, Molecular Res Center/ Invitrogen). Reverse-transcribed it into cDNA using oligonucleotide primers complementary to exon 8 of the D2 gene (Superscript III, Invitrogen). Use the resulting cDNA in a PCR reaction with oligonucleotide primers complementary to exon 5 (5¢-(d GTGTATCATTGCCAACCCTGCC)) and exon 7 (5¢-(d TGGTGCTTGACAGCATCTCC)) on the D2 gene. Conditions for the PCR should be 94° for 30 s, 60° for 1 min, and 72° for 3.5 min for 35 cycles. 2. Perform Northern blot analysis to quantify the expression level of D2S mRNA (Fig. 2d). 3. If you obtain the transgenic mice made by others, no Southern analysis, RT-PCR and Northern blot analyses are necessary.

11

Functions of D2 Receptor Isoforms

197

4. Receptor binding assays, receptor autoradiography, and Western blot analysis (if the specific antibody is available) can determine the expression level of receptor protein (5). 3.5.4. PCR Analysis to Identify Homozygous, Heterozygous, and Wild-Type Mice

1. Make a master mix of the following composition on ice: 28.75 μL mQ H2O, 5 μL 10× polymerase buffer, 4 μL dNTP (2.5 mM), 2 μL Primer mix (50 μM), 8 μL MgCl2 (25 mM), 0.25 μL DNA polymerase Stoffel (see Notes 21 and 22). Add 48 μL of the master mix to each PCR tube. Add 2 μL DNA (tail) to each tube and pipette up and down. Add one drop of mineral oil to each tube (optional). 2. PCR setting (see Note 23): step 1: 96° for 20 s, step 2: 60° for 1 min, step 3: 72° for 2 min. Repeat above cycle 35 times. After finishing, keep at 4° (see Note 24). 3. Identify the PCR products by running it on a 1.5% agarose gel.

4. Notes 1. Always use Milli-Q H2O (mQ H2O) or water equivalent in purity. Most buffers need to be autoclaved. For buffers that cannot be autoclaved (e.g., 10% SDS, ethidium bromide, Denhardts solution), use autoclaved mQ H2O or equivalent. Sometimes it is more appropriate to sterilize the solution with a filter (0.2 μm). 2. Genomic DNA fragments are also available from various companies (e.g., BACPAC Resource Center (BPRC), the Children’s Hospital Oakland Research Institute in Oakland, California, USA). 3. If Ensembl is used to predict the restriction enzyme sites, always confirm with your own mapping of the restriction sites. This is especially important if your gene sequence was obtained from a strain different from C57BL/6J, which is the genome represented on Ensembl (16). 4. The efficiency of homologous recombination depends on several factors. The first is similarity between the construct DNA and its corresponding endogenous counter (17) that can be best achieved using DNA for the construct that comes from the same mouse strain as the ES cells. The second is the length of the homologous arms (18) which generally should be several kilobases long on each end. Longer homologous sequences will result in recombination at high frequencies. The third is the sequence itself. Hotspots, defined as specific sequences that promote recombinase interaction with the nucleotide to induce homologous recombination, have also

198

Y. Wang et al.

been identified in some genes (19). These hotspots are difficult to predict and so as a result difficult to avoid. Keep in mind that these exist if you find that the antibiotic gene is present, but segments of your gene after homologous recombination is unexpectedly missing. 5. Fourteen-day-old fetuses are the best. 6. The EF cells can also be inactivated by irradiation with the following process: (a) Trypsinize the cell, wash, and resuspend in medium; (b) Irradiate at 6,000 rads for 1 h; (c) Spin down and freeze at 3 × 107 cells/mL (10 tubes of 1 mL). 7. Technique must be absolutely meticulous while working with ES cells to prevent contamination with any substances that could trigger the differentiation of the pluripotent cells in culture. Some investigators try to use the exact same lot of fetal calf serum that were previously used with success to generate a gene-targeted mouse line. 8. The optimal amount of G418 might vary for each cell line. 9. On day 2, small numbers of cell death can be seen. On day 3 many cells should start dying. By day 4 there should be a lot of cell death. 10. When picking antibiotic-resistant colonies, be conscientious to pick only well-shaped and undifferentiated colonies, and avoid the flat, differentiated colonies. Healthy ES colonies that have not differentiated have relatively round and well-defined edges and grow in three dimensions. Differentiated colonies usually have an irregular shape and look flat, often differentiated cells (muscle or fibroblast-like cells) can be seen. Even in a normal culture some colonies are differentiated. 11. Method 2 for freezing ES clones: (a) Aspirate the medium from the 24-well plates; (b) Add 150 μL of trypsin and incubate for 5 min at 37°; (c) Add 100 μL of ES medium, pipet up and down (10×) and transfer to chilled pre-labeled freezing tubes; (d) Add 200 μL of ice-cold 2× freezing medium (80% FCS + 20% DMSO, filter sterilized (0.2 μm) and stored at 4°; (e) Close the tubes, shake to mix, and freeze slowly at −70°; (f) Transfer tubes to liquid nitrogen the next day. 12. The plates can be kept at −70° for several months. 13. Aim to have at least 5 or 6 colonies of ES cells that have the gene targeted as expected. 14. For most enzymes, a universal buffer can be used. The composition of 10× concentrated universal buffer is as follows: 200 mM Tris–HCl (pH 7.5), 70 mM MgCl2, 500 mM KCl, 10 mM 2-me (0.875 μL 2-mercaptoethanol/mL), 0.5 mM

11

Functions of D2 Receptor Isoforms

199

bovine serum albumin/mL. For some enzyme (EcoRI), additional salt (1 M NaCl for 10× solution) can be added. 15. Use foster mothers that are experienced in supporting their litter in utero and ex utero. Overweight mice should be avoided because they tend to have large fat pads that contain blood vessels that may be easily nicked, thus making the surgery site bloody, which can block your view during the transfer of the blastocysts. 16. For Method 2 of freezing (see Note 11): thaw the tube at 37°. Suspend cells in 10 mL of medium, spin down to collect the cells, and resuspend in ES medium. Transfer the cells to two 24 wells. 17. To maximize your chances to obtain a germline transmitted pup, aim to devote at least two separate cycles of injection and embryo transfer for each colony of gene-targeted ES cells. 18. If using a rotating rack, wrap the tube with Parafilm to prevent the solution leak out. 19. If there is no rotating rack or shaker available, you can put tubes in a foam box in oven overnight. Vortex the tubes at least twice during the incubation period to help break down the tissue to release the DNA. The tail tissue should be completely dissolved with only the bones left. 20. Alternatively, you can put tubes in a foam box in the oven to warm up for at least 30 min, and then pipet up and down the DNA using a yellow pipette tip to disperse the pellet. Then, put tubes back in the oven for overnight. If necessary, pipet up and down to disperse the DNA again at a later time. 21. When preparing the master mix, it is better to prepare one extra sample for every 10–20 samples because pipetting can have error. 22. Primer mix: 4 oligonucleotide primers are used—two with sequence complementary to the D2 gene (around exon 6 regions) and two with sequence complementary to the neo gene. 23. PCR conditions may vary, depending on the gene. For example, Dr Sasaoka uses the following PCR setting: step 1: 94° 1 min, step 2: 94° 1 min, step 3: 60° 2 min, step 4: 72° 2 min, repeat steps 2–4 30 times, step 5: 72° 5 min, step 6: hold at 4° (20). 24. The DNA in PCR tubes should be denatured as soon as possible; this can be achieved by starting the program, letting the temperature rise to 96°, pausing the program, loading the PCR samples, and then resuming the program. For some genes, an extra period (20 s or longer) of denaturation at the beginning is necessary.

200

Y. Wang et al.

Acknowledgments We thank Dr. Luc Van Kaer (Vanderbilt University) for helping establish the ES cell transfection protocol. References 1. Dal Toso R, Sommer B, Ewert M, Herb A, Pritchett DB, Bach A, Shivers BD, Seeburg PH (1989) The dopamine D2 receptor: two molecular forms generated by alternative splicing. EMBO J 8:4025–4034 2. Giros B, Sokoloff P, Martres MP, Riou JF, Emorine LJ, Schwartz JC (1989) Alternative splicing directs the expression of two D2 dopamine receptor isoforms. Nature 342:923–926 3. Monsma FJ Jr, McVittie LD, Gerfen CR, Mahan LC, Sibley DR (1989) Multiple D2 dopamine receptors produced by alternative RNA splicing. Nature 342:926–929 4. Chio CL, Hess GF, Graham RS, Huff RM (1990) A second molecular form of D2 dopamine receptor in rat and bovine caudate nucleus. Nature 343:266–269 5. Wang Y, Xu R, Sasaoka T, Tonegawa S, Kung M-P, Sankoorikal E-B (2000) Dopamine D2 long receptor-deficient mice display alterations in striatum-dependent functions. J Neurosci 20:8305–8314 6. Usiello A, Baik J-H, Rouge-Pont F, Picetti R, Dierich A, LeMeur M, Piazza PV, Borrelli E (2000) Distinct functions of the two isoforms of dopamine D2 receptors. Nature 408: 199–203 7. Smith JW, Fetsko LA, Xu R, Wang Y (2002) Dopamine D2L receptor knockout mice display deficits in positive and negative reinforcing properties of morphine and in avoidance learning. Neuroscience 113:755–765 8. Xu R, Hranilovic D, Fetsko LA, Bucan M, Wang Y (2002) Dopamine D2S and D2L receptors may differentially contribute to the actions of antipsychotic and psychotic agents in mice. Mol Psychiatry 7:1075–1082 9. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning: a laboratory manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY 10. Mombaerts P, Clarke AR, Hooper ML, Tonegawa S (1991) Creation of a large genomic deletion at the T-cell antigen receptor beta-subunit locus in mouse embryonic stem cells by gene targeting. Proc Natl Acad Sci USA 88:3084–3087

11. Adra CN, Boer PH, McBurney MW (1987) Cloning and expression of the mouse pgk-1 gene and the nucleotide sequence of its promoter. Gene 60:65–74 12. Ausubel F, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, Struhl K (1995) Short protocols in molecular biology, 3rd edn. Wiley, New York 13. Nagy A, Gertsenstein M, Vintersten K, Behringer R (2003) Manipulating the mouse embryo: a laboratory manual, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor 14. Matise M, Auerbach W, Joyner AL (2000) Production of targeted embryonic stem cell clones. In: Joyner A (ed) Gene targeting: a practical approach, 2nd edn. Oxford University Press, Oxford 15. Papaioannou V, Johnson R (2000) Production of chimeras by blastocyst and morula injection of targeted ES cells. In: Joyner A (ed) Gene targeting: a practice approach, 2nd edn. Oxford University Press, Oxford 16. Dang MT, Yokoi F, Yin HH, Lovinger DM, Wang Y, Li Y (2006) Disrupted motor learning and long-term synaptic plasticity in mice lacking NMDAR1 in the striatum. Proc Natl Acad Sci USA 103:15254–15259 17. te Riele H, Maandag ER, Berns A (1992) Highly efficient gene targeting in embryonic stem cells through homologous recombination with isogenic DNA constructs. Proc Natl Acad Sci USA 89:5128–5132 18. Thomas KR, Deng C, Capecchi MR (1992) High-fidelity gene targeting in embryonic stem cells by using sequence replacement vectors. Mol Cell Biol 12:2919–2923 19. Smith GR (1994) Hotspots of homologous recombination. Experientia 50:234–241 20. Sasaoka T, Imamura M, Araishi K, Noguchi S, Mizuno Y, Takagoshi N, Hama H, Wakabayashi-Takai E, Yoshimoto-Matsuda Y, Nonaka I, Kaneko K, Yoshida M, Ozawa E (2003) Pathological analysis of muscle hypertrophy and degeneration in muscular dystrophy in gamma-sarcoglycan-deficient mice. Neuromuscul Disord 13:193–206

Chapter 12 Genomic Strategies for the Identification of Dopamine Receptor Genes in Zebrafish Wendy Boehmler, Jessica Petko, Victor A. Canfield, and Robert Levenson Abstract In this chapter, we describe the identification and cloning of D2-like dopamine receptor (DR) genes in zebrafish, a vertebrate model genetic organism. To identify DR genes, we performed searches of the zebrafish genomic sequence database that yielded contig segments of several D2-like DR genes. From these sequences, we amplified full-length cDNAs encoding three D2, one D3, and three D4 DR receptor subtypes via RT-PCR. The predicted proteins displayed 57–72% amino acid identity when compared to their human DR counterparts. To validate the identity of zebrafish DR genes, each of the genes was mapped by using the T51 radiation hybrid panel. With the exception of drd2b and drd4b, each of the zebrafish DR genes mapped to chromosomal positions that were syntenic with regions of human chromosomes containing orthologs of the zebrafish DR genes. To further validate the identity of the D2-like DR genes in zebrafish, we conducted phylogenetic analysis which supported the predicted identities of the cloned DR receptor cDNAs. Key words: Cloning, D2-like DR, RT-PCR, Phylogenetic analysis, Contig, Dopamine

1. Introduction Dopamine is a catecholamine neurotransmitter that in mammals controls a wide variety of functions including locomotion and learned behavior. Dysfunction in dopaminergic signaling is believed to underlie several neuropathologies including schizophrenia (SZ), Tourette’s syndrome, and Parkinson’s disease (PD). It has been postulated that manifestation of SZ involves dopamine hyperactivity in the brain (1) and this hyperactivity may be due to the dysregulation of dopamine receptor signaling. Much of what is known about SZ has been driven by the development of antipyschotic drugs that antagonize D2-like DRs. Even though such antipsychotic drugs effectively treat the positive symptoms of SZ, several Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_12, © Springer Science+Business Media, LLC 2013

201

202

W. Boehmler et al.

classes can cause discomforting side effects such as unwanted sedation, movement disorders including tardive dyskinesia, and pseudopregnancy. In contrast to SZ, PD is characterized by a decrease in dopaminergic signaling due to the degeneration of dopaminergic neurons in the substantia nigra (2). The lack of dopamine signaling in PD patients results in symptoms that include tremors, rigidity, and bradykinesia. To better understand DR function, we initiated studies designed to characterize D2-like DR genes in zebrafish. Zebrafish is an attractive model genetic system for the study of many biological processes. The zebrafish model system provides several advantages for identifying and characterizing genes and their function. The zebrafish embryo develops ex utero and is transparent, which allows for gene expression analysis using whole-mount in situ hybridization. Zebrafish are amenable to forward and reverse genetic techniques. Mutant zebrafish can be screened morphologically and behaviorally to identify new genes and their function. Antisense morpholino oligonucleotides make it possible to knock down expression of a specified gene during the first several days of development in order to determine its function. Zebrafish display a cohort of behaviors related to DA signaling including locomotion and conditioned place preference. Additionally, chemicals and drugs can be added directly to tank water, thus making zebrafish an attractive model system for toxicology and drug screening. We have identified and cloned the complete cohort of D2-like DR genes in zebrafish. Mining the zebrafish genomic sequence database allowed us to identify contigs containing segments of several D2-like DR genes. From these sequences, we were able to amplify full-length cDNAs encoding three D2, one D3, and three D4 DR subtypes. When these studies were originally performed, extensive searches of the expressed sequence tags (EST) database retrieved no results for zebrafish DRs. In retrospect, this probably reflected low expression of the receptors in zebrafish. In addition, the first zebrafish genome assemblies had extensive gaps that made it difficult to identify complete exons of the DR genes. Subsequent releases of preliminary genomic assemblies and tools for performing BLAST searches of the zebrafish genome (http://www.ensembl. org/Danio_rerio/blastview) have greatly facilitated identification of additional exons and assignment of exons to genes.

2. Materials 1. TRIzol® Reagent (Invitrogen, Life Technologies, Grand Island, NY). 2. SuperScript® II Reverse Transcriptase (Invitrogen).

12

Dopamine Receptors in Zebrafish

203

3. REDTaq® DNA Polymerase (Sigma-Aldrich, St. Louis, MO). 4. TOPO® TA Cloning® Kit (Invitrogen). 5. Zebrafish are sold by various vendors (e.g., SoBran, Inc., Fairfax, VA and Charles River, Wilmington, MA).

3. Methods 3.1. Cloning and Characterization of Zebrafish DR Genes

The general strategy we used to identify, clone, and validate zebrafish D2-like DR genes is outlined in Fig. 1. To identify zebrafish DR genes, we first searched the available EST database. Initial BLAST (Basic Local Alignment Search Tool) queries of the EST database using human DR nucleotide sequences failed to identify any zebrafish cDNAs with sequence homology to human

Fig. 1. Schematic representation of the strategy used to identify, clone, and validate zebrafish D2-like DR genes.

204

W. Boehmler et al.

DR genes. The Wellcome Trust Sanger Institute began sequencing the zebrafish genome in early 2001. Segments of zebrafish dopamine receptor genes were identified by using mammalian and fish polypeptide sequences as probes in low stringency SSAHA (3) and BLAST searches of the zebrafish genomic reads generated by the Sanger Center (http://www.sanger.ac.uk/Software/analysis/ SSAHA/). One of the polypeptide sequences used was the human D2 DR (DRD2; NM_000795). Specifically, this sequence was used in a TBLASTN search, which will search a translated nucleotide database using an amino acid sequence. The results of these searches allowed for the identification of portions of exons representing a minimum of three zebrafish D2-like, one D3-like, and three D4-like DR genes. All genomic sequence reads encompassed entire zebrafish DR genes except for D2b and D2c, which were missing their 5¢ and 3¢ ends, respectively. Subsequent releases of preliminary genomic assemblies and enhanced tools for performing BLAST searches (http://www.ensembl.org/Danio_rerio/ blastview) allowed for the identification of those additional exons. Polymerase chain reaction (PCR) primers that overlapped conserved initiation and termination codons were designed based on our genomic sequence reads (see Note 1). The corresponding full-length cDNAs were generated by RT (reverse transcriptase)PCR. First-strand cDNA synthesis was performed using SuperScript RT (Life Technologies) according to the manufacturer’s protocols (see Note 2). Random hexamers were used to prime cDNA synthesis and total RNA from adult zebrafish was used as template. REDTaq DNA Polymerase (Sigma) was used for subsequent amplification. For each cDNA, it was necessary to use nested primers for PCR because the first round of PCR invariably failed to amplify the transcripts. We believe this may be due to low expression of DR transcripts in whole adult zebrafish. Nested PCR is a technique that employs two primer pairs and two sequential PCR runs to amplify a single target sequence. Primers and annealing temperatures for the D2-like DR are listed in Table 1. PCR was carried out using a TD-7500 Thermal Cycler (Hybaid). An initial 4 min denaturation step at 94°C was followed by 34 cycles at 94°C for 30 s, 45–52°C for 30 s, and 72°C for 90 s. A final elongation step was carried out at 72°C for 10 min. All cDNAs were TOPO-cloned and then sequenced using an ABI 377-automated DNA sequencer. Sequences were verified by sequencing at least three independent clones and comparison to genomic sequences. 3.2. Validation of DR Gene Identity

We utilized multiple parameters to confirm the identities of the predicted DR polypeptides. These included BLAST analysis, comparison of gene structure (i.e., intron/exon organization), phylogeny, and conserved synteny.

12

Dopamine Receptors in Zebrafish

205

Table 1 PCR primers for the amplification of Zebrafish D2-like dopamine receptors Clone

Forward primer

Reverse primer

Ta (°C)

drd2a PCR 1 PCR 2

−34

1772

−3

1772

49 50

drd2b PCR 1 PCR 2

−15

1320

−7

1320

drd2c PCR 1 PCR 2

−44

drd3 PCR 1 PCR 2

−75

drd4a PCR 1 PCR 2

1

drd4b PCR 1 PCR 2

−5

1262

1

1257

drd4c PCR 1 PCR 2

−90

ggttctagggtctcagcttg−15 ctgatggaagtcttcacagcg18 cagaggatcttcatcatgcct6 cttcatcatgcctgtcctgaac15

−11

−35

1

gtgtgttcaacagtgcaggatcttgat1294 gtgtgttcaacagtgcaggatcttgat1294

gaattatgtctctcagtttcaggcttc−18 ggccacagctcatggatttc9

1363

gttacactgcatgttgtcaag−55 aaaatctgtccaccctctcc−16

1379

atggtagaggcagacatgcca21 atggtagaggcagacatgcca21 gcatcatggtcaatgtgacgcc17 atggtcaatgtgacgcccagt21

−15

caccccatacacagtaatgttatg1749 caccccatacacagtaatgttatg1749

gcagagatggaccacagtggaca−68 ggatcaagaaggacaatgtctgc8

atcctcagcagtgcaatatc1344 atcctcagcagtgcaatatc1344

1363

atatagcccatggtttagca1360 gcagctcaggattttaatga1343

1362

1157

ttagcatgctcaggctagcag1137 ctagcagcagccaggcagcgt1123

1143

caacctcaggaacgacagcagag1240 tcaggaacgacagcagagaag1237

1184

aatcagtacagtcttcagcatc1163 tatggtggatgtcagcagcagc1139

1160

50 52 50 52 45 47 54 52 54 52 52 52

Nucleotide +1 si the A of the ATG codon for the initiating methionine

3.2.1. BLAST Analysis

BLAST analysis indicates that each of the cDNAs we cloned encodes a polypeptide with a high degree of sequence similarity to mammalian dopamine receptors. Sequence comparisons indicate that the zebrafish polypeptides show 57–71% amino acid sequence identity to human D2, D3, and D4 receptors (Table 2). One zebrafish clone (drd3) shows highest identity (67%) to the human D3 DR, while the D2 zebrafish clones (drd2a, drd2b, and drd2c) share highest identity (66–71%) with the human D2 DR (Table 2). In contrast, drd3 exhibits 52% identity to zebrafish D2 DRs, similar to the 54–62% identity between mammalian D2 and D3 DRs. Sequence comparisons indicate that the zebrafish D4 DR polypeptides show 57–59% amino acid identity to mammalian D4 DRs. In contrast, the zebrafish D4 DRs share very low sequence identity with human D2 and D3 DRs (35–40%) (Table 2). Over the years, continued mining of the zebrafish genomic and EST databases has

206

W. Boehmler et al.

Table 2 Pairwise comparisons between Zebrafish and human D2-like receptors Zebrafish

D2a

D2b

D2c

D3

D4a

D4b

D4c

Human D2

71

66

71

52

38

38

37

Human D3

58

58

59

67

37

39

39

Human D4

37

40

35

39

59

57

58

Numbers represent percent amino acid identity

failed to uncover additional D2, D3, or D4 dopamine receptor genes suggesting that zebrafish likely possess three D2 DR genes, a single D3 DR gene, and three D4 DR genes (see Note 3). 3.2.2. Structural Comparisons Between Zebrafish and Human DRs

Sequence comparisons between the human, rat, and zebrafish D2, D3, and D4 DRs have been previously described (4, 5). By aligning the predicted zebrafish and human DR polypeptides, we identified seven putative transmembrane (TM) domains that are highly conserved with the TM segments of the human D2-like DRs. In addition to the TM segments, several other regions are highly conserved between mammalian and zebrafish DRs. For D2 DRs, these regions include the first and second intracellular loops, three short segments within the third intracellular loop, and the C-terminal tail. D3 receptors exhibit fewer highly conserved regions which include the first (but not the second) intracellular loop and the C-terminus. For D4 receptors, the majority of the conserved regions are the TM segments. In addition, the intron/exon organization of the zebrafish D2, D3, and D4 DR genes is virtually identical with respect to their mammalian counterparts, suggesting that the zebrafish and mammalian genes arose from a common ancestral gene. It is interesting to note that many GPCRs lack introns, however the human and zebrafish D2 and D3 DR genes each contain seven exons, while the human and zebrafish D4 DR genes each contain 4 exons. Overall, the third intracellular loop of the D2-like DRs is highly divergent between zebrafish and mammals, as well as between each of the zebrafish DR subtypes. It is possible that this sequence divergence may reflect functional differences between the various zebrafish DRs.

3.2.3. Phylogenetic Analysis

The procedures originally described for phylogenetic analysis (4, 5) were repeated, combining D4 with D2/D3 sequences, and using additional sequences not available at the time of the initial analysis.

12

Dopamine Receptors in Zebrafish

207

1. Using the human D2, D3, and D4 protein sequences as probes, we performed BLAST searches (6) to identify dopamine receptor sequences from additional species. These searches employed the blastp program to search nonredundant peptide databases in ncbi/GenBank (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Including the seven zebrafish clones and several additional sequences previously collected, this procedure yielded 31 sequences from nonmammalian vertebrates. All sequences that were not obviously incomplete were collected at this stage. Using the same procedure, we also collected eight outgroup sequences, comprising three invertebrate sequences and five human sequences representing GPCRs highly similar to D2, D3, and D4. 2. These 42 sequences were aligned using the GCG program “pileup” (7). Alternatively, the freely available ClustalW program (8) (http://www.clustal.org/) could have been used for multiple sequence alignment. Nine sequences that were truncated or contained large insertions or deletions in the alignment were discarded, and the alignment process was repeated. 3. The GCG program “pretty” was used to generate a consensus sequence for the revised alignment. Aligned sequences were inspected to locate regions in which alignments were ambiguous. These regions were excluded from phylogenetic analysis either by removing individual sequences that could not be aligned (three cases) or by using only unambiguously aligned regions. The final analysis contained 211 aligned amino acids from 30 sequences (23 vertebrate D2, D3, or D4 sequences plus 7 outgroup sequences). 4. The consensus sequence was edited to distinguish retained from excluded amino acid positions. A simple perl filter was then used to replace excluded amino acids with blanks and to reformat the file for use by the Phylip (3.68) suite of programs (9) (http://www.phylip.com). The equivalent changes could have been performed by hand using a word processing program. Note that ClustalW provides the option of writing aligned sequences in a Phylip-compatible format. 5. Aligned sequences were analyzed using Maximum Parsimony and Distance Matrix methods, using half-jacknife replications to estimate reliability of the deduced phylogenies. The sequence of programs used for the former was seqboot, protpars, then consense; while for the latter, we used seqboot, protdist, fitch, and consense. 6. Results of the phylogenetic analysis are shown in Fig. 2. D2, D3, and D4 clusters were obtained in 81, 76, and 100% of replicates, respectively (maximum parsimony) or in 94, 87, and 100%, respectively (using the distance method). Groupings

208

W. Boehmler et al.

Fig. 2. Phylogenetic analysis of zebrafish D2, D3, and D4 DRs. Consensus trees were generated using maximum parsimony (left) and distance methods (right ). Numbers represent percent of trees in which the clustering depicted was observed. Included sequences, in addition to zebrafish: hud2, human dopamine D2, NP_000786; hud3, human D3, AAB08750; hud4, human D4, AAB59386; parus, great tit (Parus major), AAY56686; chickd3 (Gallus gallus), ACR48171; turkeyd2 (Meleagris gallopavo), AAD03818; xend21 (Xenopus laevis), NP_001095212; carpd4b (Cyprinus carpio), CAA74977; troutd2, rainbow trout (Oncorhynchus mykiss), NP_001117843; tilpiad2, (Oreochromis niloticus), AAU87971; tetra_e, (Tetraodon nigroviridis), CAF97490; mulletd2 (Mugil cephalus), AAU87970; eeld2a (Anguilla anguilla), ABH06893; eeld2b (Anguilla anguilla), ABH06894; d215 (Takifugu rubripes), CAA56456; d222 (Takifugu rubripes; (17)). Outgroup sequences (only first is shown on tree): drod2 (Drosophila melanogaster), NP_001014760; Drosophila simulans, XP_002103025; human adrenergic alpha2A, AAF91441; human serotonin 5HT4, NP_001035261; human dopamine D5, NP_000789; human adrenergic alpha1D, NP_000669; human histamine H2, NP_071640. Excluded sequences: chicken (Gallus gallus), NP_001136321; Taeniopygia guttata (zebra finch), XP_002196676; Xenopus laevis, P34973; Tetraodon nigroviridis, CAF98376, CAF95731, CAG04235, CAF92229; minnow (Pimephales promelas), ABS30830; carp (Cyprinus carpio), CAA74976, CAB40081, CAA74974; Branchiostoma floridae, XP_002211994.

12

Dopamine Receptors in Zebrafish

209

within the D2 cluster were supported less strongly, with one exception, and differed somewhat between the two methods. Both methods identified zebrafish d2b as the most distant member of the D2 cluster, while the remaining D2 sequences clustered in 99% of trees with the distance method (80% with maximum parisomony). Within the D3 group, the fish and mammal/bird clusters were strongly supported. In the D4 cluster, zebrafish d4b was an outlier, although separation of this sequence from the remaining sequences was supported by only 57% (maximum parsimony) or 65% (distance method) of trees. Outgroup sequences located the root of the tree between the D4 and D2/D3 clusters, consistent with the differences in intron/exon organization, while D2 and D3 clustered together (100% of trees, both methods). The only exception among the outgroup sequences was the Drosophila D2-like sequence, which clustered with D4 sequences using the distance method (71% support). The Drosophila sequence clustered with other outgroup sequences using maximum parisomony (62% of trees), while clustering with D4 sequences occurred in only 29% of trees obtained using this method. 3.2.4. Chromosomal Mapping of Zebrafish Dopamine Receptor Genes

Each zebrafish D2-like DR gene was localized to a zebrafish chromosome via radiation hybrid mapping. This type of comparative mapping study allowed us to identify conserved synteny between zebrafish and human DR genes. Zebrafish DR genes were mapped using the Goodfellow T51 radiation hybrid (RH) panel (10). PCR products specific for each zebrafish DR gene were amplified using primers corresponding to unique sequences within each gene. PCR reactions were performed in duplicate on the RH panel using conditions optimized for each primer pair. PCR reaction products were fractionated on 2% agarose gels, and each sample scored for presence or absence of the zebrafish-specific amplicon. Linkage assignments were computed using the Zon RH mapper resource (http://zfrhmaps.tch.harvard.edu/ZonRHmapper/). Our mapping data shows that the zebrafish drd3 gene is located on chromosome 24, while the human D3 dopamine receptor DRD3 gene maps to chromosome 3 (11). Zebrafish chromosome 24 contains orthologs of additional human genes located on human chromosome 3 (Table 3). Comparative gene mapping thus provides strong additional evidence that the zebrafish drd3 DR gene and the mammalian D3 DR gene are in fact orthologous. The zebrafish drd2a and drd2c DR genes map to chromosomes 15 and 5, respectively, while the human D2 DR gene (DRD2) has been localized to chromosome 11q23 (12). Both zebrafish chromosome 15 and chromosome 5 contain multiple orthologs of human genes located on chromosome 11 (Table 3). The presence of lim1 and lim6 (zebrafish duplicates of the single human LHX1 gene) on zebrafish chromosome 15 and chromosome 5 (13) is consistent

210

W. Boehmler et al.

Table 3 Markers Syntenic with dopamine receptor genes in Zebrafish and human Zebrafish gene

Human ortholog

Name

Accession no

Map position Name

Map position

drd2a acad8 arcn1l cryab ctsc hsp47 hsp70 kiaa0102 mgc10485 mpzl3 mre11a or13.1 sesn3 tyr

AY183456 BC044203 BC050499 AF159089 AI436938 U31079 AF006006 AA494919 BC046056 BC067698 BC053202 AF012746 BC045518 NM_131013

15 15 15 15 15 15 15 15 15 15 15 15 15 15

DRD2 ACAD8 ARCN1 CRYAB CTSC CBP2 HSPA10 KIAA0102 MGC10485 MPZL3 MRE11a OR2AT4 SESN3 TYR

11q23 11 11 11q 11q14.1-q14.3 11q13.5 11 11q13.3 11q25 11q23.3 11q21 11q13.4 11q21 11q14-q21

drd2b fli1b

AY333791 BC055627

16 16

DRD2 FLI1

11q23 11q24.1-q24.3

drd2c apoa atdc htatip ins mizf slc37a2 spon1a stip1 tmprss4 wnt11

AY333792 Y13653 AI721600 AI477057 AF036326 BC057479 BC055147 AB006086 BC085642 BC048135 AF067429

5 5 5 5 5 5 5 5 5 5 5

DRD2 APOA1 ATDC HTATIP INS MIZF SLC37A2 SPON1 STIP1 TMPRSS4 WNT11

11q23 11q23-q24 11q22-q23 11q12.1 11p15.5 11q23.3 11q24.2 11p15.2 11q13 11q23.3 11q13.5

drd3 boc c3orf3b cldn11 cpb1 ek1 nktr ostalpha slc22a13 zic1

AY183455 AF461120 BI887394 AF359429 BC067637 U89295 AI584240 BC081597 AW019519 NM_130933

24 24 24 24 24 24 24 24 24 24

DRD3 BOC C3orf3 CLDN11 CPB1 EPHB3 NKTR OSTalpha SLC22A13 ZIC1

3q13.3 3q13.2 3p22.1 3q26.2-q26.3 3q24 3q21-qter 3p23-p21 3q29 3p21.3 3q24 (continued)

12

Dopamine Receptors in Zebrafish

211

Table 3 (continued) Zebrafish gene

Human ortholog

Name

Accession no

Map position Name

Map position

drd4a c11orf24 cat ckap5 fancf irf7 ldha mdkb myod nap1l4 pax6a rag1 rag2 rnh tnni2 wt1 zdhhc13

AY750152 AW154701 AJ007505 NM_001037667 CN014287 NM_200677 AF067201 BC059575 Z36945 NM_001089347 AJ507427 U71093 U71094 CF997807 BC071462 NM_131046 BC086723

25 25 25 25 25 25 25 25 25 25 25 25 25 25 25 25 25

DRD4 C11orf24 CAT CKAP5 FANCF IRF7 LDHA MDK MYOD1 NAP1L4 PAX6 RAG1 RAG2 RNH TNNI2 WT1 ZDHHC13

11p15.5 11q13 11p13 11p11.2 11p15 11p15.5 11p15.4 11p11.2 11p15.4 11p15.5 11p13 11p13 11p13 11p15.5 11p15.5 11p13 11p15.1

drd4b ms4a4a

AY750152 AW078034

DRD4 MS4A4A

11p15.5 11q12

drd4c c11orf15 ccnd1 cd81 copb1 cugbp1 dbx1a dhcr7 eif4g2a ext2 f2 fgf4 hsd17b12b htatip2 igf2 lmo1 men1 mtch2 mus81 nucb2a pax6b pc ppp1r14b psmc3 rbm4l

AY750154 BC050159 NM_131025 BC057419 AY294010 AB032726 AF030284 BC055631 BC059195 AY786508 AW115527 AF283555 BC059617 NM_131681 AF194333 AF398514 AF212919 AF176010 BC055679 NM_201493 AF061252 AF295372 BC076513 BC071390 BC067187

DRD4 C11orf15 CCND1 CD81 COPB CUGBP1 DBX1 DHCR7 EIF4G2 EXT2 F2 FGF4 HSD17B12 HTATIP2 IGF2 LMO1 MEN1 MTCH2 MUS81 NUCB2 PAX6 PC PPP14B1B PSMC3 RBM4L

11p15.5 11p15.3 11q13 11p15.5 11p15.2 11p11 11p15.1 11q13.2-q13.5 11p15 11p12-p11 11p11 11q13.3 11p11.2 11p15.1 11p15.5 11p15 11q13 11p11.2 11q13 11p15.1-p14 11p13 11q13.4–13.5 11q13 11p12–p13 11q13

4 4 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7 7

(continued)

212

W. Boehmler et al.

Table 3 (continued) Zebrafish gene

Human ortholog

Name

Accession no

Map position Name

Map position

rras2 scube2 sf1 slc17a6 st5 stk33

AI882911 NM_001014813 BC056288 AL627163 BC076471 BQ617398

7 7 7 7 7 7

11p15.2 11p15.3 11q13 11p14.3 11p15 11p15.3

RRAS2 SCUBE2 SF1 SLC17A6 ST5 STK33

with the view that these are duplicate chromosomal segments. Taken together, these data support the conclusion that drd2a and drd2c are duplicate DR genes orthologous to the single mammalian D2 DR gene. Our mapping experiments placed drd2b, the third zebrafish D2 DR gene, on chromosome 16. Although we have only been able to identify one additional gene that is syntenic between zebrafish chromosome 16 and human chromosome 11 (Table 3), improvements in the density of the zebrafish chromosome 16 map may eventually reveal whether or not these are related chromosomal segments. Our gene mapping data indicate that the zebrafish drd4a and drd4c DR genes map to chromosomes 25 and 7, respectively, whereas the human D4R DR gene (DRD4) has been localized to chromosome 11p15.5 (14). Chromosomes 25 and 7 contain multiple orthologs of human genes located on chromosome 11 (15). The fact that these zebrafish chromosomes share significant synteny with human chromosome 11 is consistent with the view that these are duplicate chromosome segments and that drd4a and drd4c are true orthologs of the single mammalian D4R DR gene. Our mapping assignments placed drd4b, the third zebrafish D4R DR gene, on chromosome 4. We were able to identify only one additional gene that is syntenic between zebrafish chromosome 4 and human chromosome 11. Again, improvements in the density of the zebrafish map may eventually reveal whether or not these are related chromosomal segments. These results are consistent with the view that zebrafish posses three paralogous D4R DR genes.

4. Notes 1. Primer Design—It is important to consider primer length(18–22 bp), GC content (<60%), and annealing temperatures (>50°C). It is also important to avoid primer secondary structure

12

Dopamine Receptors in Zebrafish

213

by avoiding repeats and runs in a sequence. There are a variety of primer design software tools available. One program that is available is OligoAnalyzer by Integrated DNA Technologies (http:// www.idtdna.com/analyzer/Applicati ons/OligoAnalyzer/ Default.aspx). 2. PCR conditions—There are several ways to optimize PCR conditions. Adding enhancing agents such as dimethyl sulfoxide (DMSO) can prevent secondary structure in difficult targets for amplification. Adjusting the amount of MgCl2 can influence the specificity of the target DNA sequence and primer interaction. The typical final concentration of MgCl2 is 1.5 mM, but ranges above and below that can work as well. 3. Functional Analysis—The homology, phylogeny, and synteny between the mammalian and zebrafish D2-like DR genes is highly supported, however this is not always the case. For example, several D2-like DR genes were identified in Drosophila with only ~33% amino acid identities shared with their human counterparts. In these particular cases, it is important to uncover the pharmacological profiles of the cloned receptors in order to verify their identities (16). References 1. Matthysse S (1974) Dopamine and the pharmacology of schizophrenia: the state of the evidence. J Psychiatr Res 11:107–113 2. Fahn S (2003) Description of Parkinson’s disease as a clinical syndrome. Ann N Y Acad Sci 991:1–14 3. Ning ZCA, Mullikin JC (2001) SSAHA: a fast search method for large DNA databases. Genome Res 11:1725–1729 4. Boehmler W, Obrecht-Pflumio S, Canfield V, Thisse C, Thisse B, Levenson R (2004) Evolution and expression of D2 and D3 dopamine receptor genes in zebrafish. Dev Dyn 230:481–493 5. Boehmler W, Carr T, Canfield V, Thisse C, Thisse B, Levenson R (2007) D4 dopamine receptor genes of zebrafish and effects of the antipsychotic clozapine on larval swimming behavior. Genes Brain Behav 6:155–166 6. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215:403–410 7. Devreux JHP, Smithies O (1984) A comprehensive set of sequence analysis programs for the VAX. Nucleic Acids Res 12:387–395 8. Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG (2007) Clustal W and

9.

10.

11.

12.

13.

14.

Clustal X version 2.0. Bioinformatics 23: 2947–2948 Felsenstein J (1981) Evolutionary trees from DNA sequences: a maximum likelihood approach. J Mol Evol 17:368–376 Kwok C, Korn RM, Davis ME, Burt DW, Critcher R, McCarthy L, Paw BH, Zon LI, Goodfellow PN, Schmitt K (1998) Characterization of whole genome radiation hybrid mapping resources for non-mammalian vertebrates. Nucleic Acids Res 26:3562–3566 Le Coniat M, Sokoloff P, Hillion J, Martres MP, Giros B, Pilon C, Schwartz JC, Berger R (1991) Chromosomal localization of the human D3 dopamine receptor gene. Hum Genet 87:618–620 Eubanks JH, Djabali M, Selleri L, Grandy DK, Civelli O, McElligott DL, Evans GA (1992) Structure and linkage of the D2 dopamine receptor and neural cell adhesion molecule genes on human chromosome 11q23. Genomics 14:1010–1018 Postlethwait JH, Woods IG, Ngo-Hazelett P, Yan YL, Kelly PD, Chu F, Huang H, HillForce A, Talbot WS (2000) Zebrafish comparative genomics and the origins of vertebrate chromosomes. Genome Res 10:1890–1902 Gelernter J, Kennedy JL, van Tol HH, Civelli O, Kidd KK (1992) The D4 dopamine receptor

214

W. Boehmler et al.

(DRD4) maps to distal 11p close to HRAS. Genomics 13:208–210 15. Woods IG, Wilson C, Friedlander B, Chang P, Reyes DK, Nix R, Kelly PD, Chu F, Postlethwait JH, Talbot WS (2005) The zebrafish gene map defines ancestral vertebrate chromosomes. Genome Res 15:1307–1314 16. Hearn MG, Ren Y, McBride EW, Reveillaud I, Beinborn M, Kopin AS (2002) A Drosophila

dopamine 2-like receptor: molecular characterization and identification of multiple alternatively spliced variants. Proc Natl Acad Sci USA 99:14554–14559 17. Macrae AD, Brenner S (1995) Analysis of the dopamine receptor family in the compact genome of the puffer fish Fugu rubripes. Genomics 25:436–446

Chapter 13 Application of Cell-Specific Isolation to the Study of Dopamine Signaling in Drosophila Eswar Prasad R. Iyer, Srividya Chandramouli Iyer, and Daniel N. Cox Abstract Dopamine neurotransmission accounts for a number of important brain functions across species including memory formation, the anticipation of reward, cognitive facilities, and drug addiction. Despite this functional significance, relatively little is known of the cellular pathways associated with drug-induced molecular adaptations within individual neurons. Due to its genetic tractability, simplicity, and economy of scale, Drosophila melanogaster has become an important tool in the study of neurological disease states, including drug addiction. To facilitate high-resolution functional analyses of dopamine signaling, it is highly advantageous to obtain genetic material, such as RNA or protein, from a homogeneous cell source. This process can be particularly challenging in most organisms including small model system organisms such as Drosophila melanogaster. Magnetic bead-based cell sorting has emerged as a powerful tool that can be used to isolate select populations of cells, from a whole organism or tissue such as the brain, for genomic as well as proteomic expression profiling. Coupled with the temporal and spatial specificity of the GAL4/UAS system, we demonstrate the application of magnetic bead-based cell sorting towards the isolation of dopaminergic neurons from the Drosophila adult nervous system. RNA derived from these neurons is of high quality and suitable for downstream applications such as microarray expression profiling or quantitative rtPCR. The versatility of this methodology stems from the fact that the cell-specific isolation method employed can be used under a variety of experimental conditions designed to survey molecular adaptations in dopamine signaling neurons including in response to drugs of abuse. Key words: Drosophila, Dopamine signaling, GAL4/UAS, Magnetic bead cell sorting, Brain, RNA isolation

1. Introduction Dopamine neurotransmission systems are found in various species where they signal the reinforcing properties of natural rewards such as food, sex, and social interaction (1–4). Genes involved directly or indirectly in the actions of drugs of abuse appear to also be conserved across evolution (5, 6), suggesting a preservation of Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_13, © Springer Science+Business Media, LLC 2013

215

216

E.P.R. Iyer et al.

dopamine mechanisms on addictive behaviors in various species. Due to its genetic tractability, simplicity, and economy of scale, Drosophila melanogaster has become an important tool in the study of neurological disease states, including drug addiction (7). Previous findings indicate that dopamine plays a key role in responses of Drosophila to addictive drugs such as ethanol, nicotine, and cocaine (5, 8–11). Magnetic bead-based cell sorting has emerged as a powerful tool that can be used to isolate select populations of cells, from a whole organism or specific tissue type such as brain, for genomic, as well as proteomic expression profiling (12, 13). Coupled with the temporal and spatial specificity of the GAL4/UAS system, we demonstrate the application of magnetic bead-based cell sorting to the specific isolation of genetically labeled, wild-type populations of dopaminergic neurons from the Drosophila adult brain. This technique provides a significant advantage over existing methodologies wherein samples derived from the brain may be of heterogeneous cell populations thereby confounding molecular analyses of dopamine signaling in either wild-type or experimentally challenged dopaminergic neurons. RNA derived from these neurons is suitable for downstream applications such as microarray expression profiling or quantitative rtPCR. The versatility of this methodology stems from the fact that cell-specific isolation can be used under a wide variety of experimental conditions designed to survey molecular signaling adaptations in dopaminergic neurons such as in response to drugs of abuse. Moreover, the availability of Drosophila strains and commercially available reagents (including antibodies, magnetic beads, and RNA purification kits optimized for recovery from relatively small, pure populations of cells at low elution volumes) provides a unique tool for investigating dopamine signaling at the level of cellular resolution.

2. Materials 2.1. Magnetic Bead Preparation

1. 10× Phosphate-buffered saline (PBS) (MP Bioproducts, Solon, OH) diluted to 1× working solution with RNase-free doubledistilled water (ddH2O) and stored at room temperature (see Note 1). 2. RNase-AWAY (Sigma-Aldrich, St. Louis, MO) stored in a spray-bottle at room temperature for rendering the working surfaces RNAse free. 3. Biotinylated Rat anti-Mouse-CD8a antibody (eBioscience, San Diego, CA) 500 μg/mL stored at 4°C. 4. Dynabeads MyOne Streptavidin T1 (Invitrogen, Life Technologies, Grand Island, NY), stored at 4°C. Once coupled

13

Cell-Specific Isolation to Study Dopamine Signaling in Drosophila

217

to antibody, the magnetic beads are stored at 4°C until needed and maximally up to 1 month. 1 mg of beads can bind up to 20 μg of biotinylated antibody. 5. (8) Rare Earth Magnet Blocks: 2 in. (length) × 0.5 in. (width) × 0.125 in. (thickness) (Magcraft, Vienna, VA). Alternatively DynaMag™-2 (Invitrogen) may be used. 6. Tabletop Vortex. 7. Crushed ice for incubating the antibody bead mixture. 2.2. Dissecting and Washing Adult Fly Heads

1. 10× Phosphate-buffered saline (PBS) diluted to 1× working solution in RNase-free ddH2O and chilled on ice (see Note 2). 2. Transgenic flies expressing UAS-mCD8::GFP (Bloomington Stock Center, stock# 30125) under the control of ple-GAL4 (Bloomington Stock Center, stock# 8848). ple-GAL4 (also known as TH-GAL4) drives the expression of GAL4 under the control of the pale (ple) promoter specifically within dopaminergic neuron clusters in the adult Drosophila brain which project to specific regions within the mushroom bodies and to the central complex (14) (Fig. 1). The Drosophila ple gene encodes the enzyme tyrosine hydroxylase, which is required for dopamine biosynthesis. 3. Glass Pasteur pipettes, fire-polished by quickly rotating the pipette tip over a hot Bunsen burner flame until the pipette edges become smooth. 4. Dissecting tools: two pairs of sharp Dumont No. 5 forceps.

2.3. Dissociating the Tissue into a Single Cell Suspension

1. 10× Phosphate-buffered saline (PBS) diluted to 1× working solution in RNase-free ddH2O and chilled on ice. 2. Kontes Glass Tissue Grinder, 7 mL working capacity, with large clearance pestle (Kimble Chase, Vineland, NJ) (see Note 3).

Fig. 1. Dopaminergic neuron clusters in adult Drosophila brain. Confocal image of whole mount adult Drosophila brain from a strain bearing the ple-GAL4 and UAS-mCD8::GFP transgenes. ple-GAL4 drives expression of UAS-mCD8::GFP specifically in dopaminergic neurons of the adult brain.

218

E.P.R. Iyer et al.

3. BSA (Bovine Serum Albumin), Fraction V. Freshly prepare 10–15 mL of 1% BSA solution in 1× PBS and store on ice (see Note 1). This is used for coating the tubes and pipettes to reduce cell loss due to adhesion to glass surfaces. 4. 30 μm cell filters, e.g., MACS Pre-Separation Filter (Miltenyi Biotec, Cambridge, MA). 5. Pipettes (P-1000, P-100, P-10) with disposable, low-retention plastic tips. 6. Pasteur pipettes with fire polished tips of standard diameter, narrowed to ~50% of standard diameter for trituration. Fire polishing and narrowing of Pasteur pipettes is achieved over a Bunsen burner flame by rotating the tip in the flame to polish edges and narrow tip diameter. 7. Fluorescent stereomicroscope equipped with GFP filter-set (a Leica MZ16FA was used in this protocol). 8. Table top micro-centrifuge (1–16,000 × g). 9. Tabletop Vortex. 10. Crushed ice. 2.4. Magnetic Bead Cell Sorting

1. 10× Phosphate-buffered saline (PBS) diluted to 1× working solution in RNase-free ddH2O and chilled on ice. 2. Anti-CD8a antibody-coated Subheading 3.1, step 4.

magnetic

beads

from

3. BSA (Bovine Serum Albumin), Fraction V. Freshly prepare 10–15 mL of 1% BSA solution in 1× PBS and store on ice. This is used for coating the tubes and pipettes to reduce cell loss due to adhesion to glass surfaces. 4. (8) Rare Earth Magnet Blocks: 2 in. (length) × 0.5 in. (width) × 0.125 in. (thickness) (Magcraft). Alternatively DynaMag™-2 (Invitrogen) may be used. 5. Pipettes (P-1000, P-100, P-10) with disposable, low retention plastic tips. 6. Standard hemocytometer to count the cells and estimate purity. 7. Pasteur pipettes with fire polished tips of standard diameter. 8. Fluorescent stereomicroscope equipped with GFP filter set (a Leica MZ16FA was used in this protocol). 9. Table top micro-centrifuge (1–16,000 × g). 10. Tabletop Vortex. 11. Crushed ice. 2.5. RNA Isolation from Magnetic Bead Sorted Cells

1. PicoPure® RNA Isolation Kit (Applied Biosystems, Life Technologies). 2. Pipettes (P-1000, P-100, P-10) with disposable, low retention plastic tips.

13

Cell-Specific Isolation to Study Dopamine Signaling in Drosophila

219

3. (8) Rare Earth Magnet Blocks: 2 in. (length) × 0.5 in. (width) × 0.125 in. (thickness) (Magcraft). Alternatively DynaMag™-2 (Invitrogen) may be used. 4. Heating block pre-heated to 42°C stored inside an incubator, pre-set to 42°C. 5. Table top micro-centrifuge (1–16,000 × g). 6. Tabletop Vortex. 7. RNAse-Free DNAse set (Qiagen, Valencia, CA) (Optional step. Needed if performing DNAse treatment prior to RNA column purification).

3. Methods To ensure high-quality RNA isolation from dopaminergic neurons, it is imperative that standard lab procedures for maintaining a clean, RNase-free environment be observed throughout this protocol in order to prevent RNA degradation (see Note 1). Preparation for this method requires isolation of age-matched Drosophila adults bearing a ple-GAL4 transgene and a UAS-mCD8::GFP transgene in order to genetically label dopaminergic neurons for visualization and CD8-mediated cell isolation via magnetic bead cell sorting. Prior to initiating dissection of adult Drosophila heads, it is important to prepare magnetic beads in advance for use in cell binding. The antibody-bound magnetic beads are then stored at 4°C until ready for cell isolation. Following dissection of heads from pleGAL4,UASmCD8::GFP adults, the tissue is dissociated and subjected to filtration into single cell suspensions. These single cell suspensions are verified by epi-fluorescent microscopy for the presence of the genetically labeled dopaminergic neurons, which uniquely express green fluorescent protein (GFP). Once verified, antibody-coated magnetic beads are added to the single cell suspensions and allowed to incubate for a limited time to promote high affinity and specificity interactions between the antibodycoated beads and the genetically labeled dopaminergic neurons. Homogenous, cell-specific isolation is achieved by placing the cell suspension in a strong magnetic field to pellet the target cells, followed by rigorous washing to remove any non-specific contaminating cells. The isolated, purified population of cells can be visually verified for specificity by epi-fluorescent assessment of GFP expression in all cells. Ultimately, the purified population of dopaminergic neurons is subjected to RNA isolation for downstream applications. Figure 2 schematically summarizes the procedures involved in this protocol and Table 1 summarizes the timing required for each step of the protocol.

220

E.P.R. Iyer et al.

Fig. 2. Schematic of procedure for magnetic bead cell sorting of Drosophila dopaminergic neurons. (a) Age-matched, intact adult Drosophila heads carrying the ple-GAL4 and UAS-mCD8::GFP transgenes are dissected. (b) The heads are transferred to a microfuge tube and washed/vortexed to remove any debris. (c) The heads are then transferred into a 7 mL dounce to homogenize the brain tissue followed by trituration to further dissociate the tissue into a cell suspension. (d, e) The homogenate is then filtered using a 30 μm cell filter to remove any large cellular debris, including the cuticle from the head and filter out the single cell suspension. The filtrate solution contains a single cell suspension of different cell types including neurons and glia. (f) Biotinylated anti-mouse-CD8-antibody-coated magnetic Streptavidin T1 Dynabead are added to the cell suspension, and incubated on ice for 30–60 min. (g) The magnetic beads specifically bind to the dopaminergic neurons that express a mouse CD8-tagged GFP fusion protein under the control of the ple-GAL4 driver. (h, i) The magnetic beadcoated cells are separated by placing the cell solution in a strong magnetic field. The supernatant is discarded, and the cells are washed three times to remove any nonspecific cells, resulting in (j) highly enrichment of specific dopaminergic neurons. Panels (b–j) are adapted in part (14).

Table 1 Summary of experimental work flow and timings Procedure

Timing

Preparing magnetic beads for binding (prepared prior to cell isolation experiment)

75–90 min

Dissecting and washing adult fly heads

10–15 min

Dissociating the tissue into a single cell suspension

18–20 min

Magnetic bead cell sorting

45–75 min

RNA isolation from magnetic bead sorted cells

60–75 min

3.1. Preparing Magnetic Beads for Binding Cells: (75–90 min)

This step must be completed prior to the start of the experiment. The prepared beads can be stored at 4°C until needed (see Note 4). 1. Wash 100 μL of Dynabeads StreptavidinT1-coated beads three times in PBS by resuspending the magnetic beads in 1 mL of fresh 1× PBS and pelleting the beads in a strong magnetic field after each wash. Discard the supernatant after each wash step.

13

Cell-Specific Isolation to Study Dopamine Signaling in Drosophila

221

2. Resuspend the beads directly in 100 μL of undiluted biotinylated rat anti-mouse-CD8a antibody (antibody concentration is 500 μg/mL). 3. Incubate the mixture for 1 h on ice with occasional mild vortexing to prevent sedimentation [Dynabeads T1 can bind 20 μg/mg of biotinylated Ig]. 4. Wash the bead–antibody mixture three times as described in step 1 above to remove any unbound excess antibody. The magnetic beads are now coated with antibody and ready to be used. Store the bead-antibody mixture in 100 μL of 1× PBS at 4°C until use. 3.2. Dissecting and Washing Adult Fly Heads: (10–15 min)

1. Dissect 30–50 age-matched adult Drosophila heads from the ple-GAL4,UAS-mCD8::GFP strain. Dissection of adult heads is achieved by using two pairs of sharp Dumont No. 5 forceps in cold 1× PBS (see Note 2). One pair of forceps is used to hold the fly in place at the thorax, while the other pair of forceps is placed around the rear side of the fly head where it meets the thorax. Using the pair of forceps placed behind the fly head, gently separate the head from the body of the fly and transfer the intact head to a 1.6 mL microfuge tube filled with 1–1.2 mL of 1× PBS on ice (see Note 5). 2. Once all fly heads have been dissected and transferred to the microfuge tube, close the tube and vortex it at the maximum setting three times for 1 s each. 3. Using a fire-polished Pasteur pipette discard the supernatant completely. Repeat the wash (Subheading 3.2, step 1) and vortex (Subheading 3.2, step 2) steps 3–4 times until the supernatant is visibly clear of any and debris.

3.3. Dissociating the Tissue Into a Single Cell Suspension: (18–20 min)

1. To avoid the cells from sticking to the glass surface of the prechilled 7 mL Kontes tissue grinder and large clearance pestle (see Note 3), pre-coat the tissue grinder and pestle with a 1% BSA in 1× PBS solution and after a brief rinse discard the BSA solution. Subsequently, using a P-1000 pipette with a cut low retention filter plastic filter tip, transfer the heads in 1 mL 1× PBS from step Subheading 3.2, step 3, and add an additional 3–4 mL of 1× PBS to the BSA-coated tissue grinder. 2. Homogenize the tissue with slow and steady strokes, avoiding frothing (approximately 20–30 strokes) (see Note 6). 3. To assess the cell dissociation levels, pipette out a small sample of the solution (up to 50 μL), and observe it under a fluorescent stereomicroscope using the GFP filter set. If one still observes intact or incompletely dissociated tissue, homogenize further. 4. Triturate the solution five times with a fire-polished Pasteur pipette narrowed to approximately 50% of the standard tip diameter (see Note 7).

222

E.P.R. Iyer et al.

5. Filter the solution through a 30-μm cell filter and collect the cell filtrate in a 1.6 mL microfuge tube. The resulting solution should consist of a single cell suspension and is now ready for magnetic cell sorting. Take a small sample of the dissociated cells and observe it under a fluorescent stereomicroscope. If cell-clumps are observed, pass the suspension again slowly through a new 30 μm filter. 3.4. Magnetic Bead Cell Sorting: (45–75 min, Depending on Antibody Incubation Time)

1. Add 15 μL of antibody-coated magnetic beads per 1 mL of cell suspension (Subheading 3.3, step 5). The remaining antibodyconjugated magnetic beads can be stored at 4°C until needed for subsequent cell isolations. 2. Incubate the cells with antibody-coated magnetic beads for 30–60 min on ice with occasional hand-mixing via inversion of the microfuge tube (see Note 8). 3. Place the microfuge tube in a strong magnetic field for 2 min. All positively selected cells along with unbound antibodycoated magnetic beads will separate to the side of the tube. 4. Slowly pipette the supernatant, making sure not to disturb the cell pellet and discard. 5. Wash the cells 3–4 times in fresh, ice-cold 1× PBS followed by magnet-induced pelleting to remove any remaining nonspecific cells. 6. Resuspend the specifically bound cells coupled to the magnetic beads in 30 μL of fresh, ice-cold 1× PBS. 7. To approximate the purity and yield of cells, pipette 5 μL of the cell suspension on the polished surface of a hemocytometer and count all the visible fluorescent cells under a fluorescent stereomicroscope equipped with a GFP filter set. Also assess the relative amount of nonfluorescent cells or any other signs of impurities. Typically the sample will be highly enriched for fluorescently labeled, GFP-positive cells (Fig. 3).

3.5. RNA Isolation from Magnetic Bead Sorted Cells: (60–75 min)

Although this protocol is focused on RNA isolation as described here, this step of the overall dopaminergic cell isolation can be replaced by other protein/DNA/chromatin isolation methods depending on nature of the study. 1. After counting, pellet the cells in a magnetic field, discard the supernatant and add 20 μL of extraction buffer from the PicoPure™ RNA Isolation Kit (Molecular Devices). Depending upon cell number one may need to add a higher volume of extraction buffer. 2. Vortex the tube at maximum speed to enable the mixing of cell pellet with extraction buffer. 3. Incubate the microfuge tube at 42°C for 30 min.

13

Cell-Specific Isolation to Study Dopamine Signaling in Drosophila

223

Fig. 3. Magnetic bead isolated dopaminergic neurons. Representative epi-fluorescent image of a field of positively selected, GFP fluorescent dopaminergic neurons isolated by magnetic bead cell sorting from the adult Drosophila brain. The purified cell population of neurons was determined to have little or no contaminating cell impurities. The variation in cell body size reflects natural variation seen in the adult brain among distinct dopaminergic neuron clusters.

4. To ensure removal of the magnetic beads prior to column purification of the RNA, the tube is briefly centrifuged at 2,000 × g for 2 min to pellet the magnetic beads. The tube is then placed in a strong magnetic field to retain the pellet, and the supernatant is transferred to a new microfuge tube. 5. Extract and column purify the RNA according to the PicoPure™ RNA extraction kit manufacturer’s instructions. DNAse treatment is optional and can be performed on column during the RNA purification according to the analysis requirement. Finally, elute the bound total RNA in a small volume (11–30 μL) of elution buffer and store at −80°C until ready for use. If desired a 1 μL aliquot may be used to assess total RNA quality on a Bioanalyzer 2100 (Agilent Technologies, Inc.) (Fig. 4).

4. Notes 1. All solutions must be prepared in RNase-free double-distilled water (not DEPC treated). BSA can be prepared prior to each experiment by dissolving the appropriate, pre-weighed quantity of BSA in PBS and vortexing at maximum speed.

224

E.P.R. Iyer et al.

Fig. 4. Bioanalyzer analysis reveals high-quality RNA purification from isolated neurons. Representative Agilent 2100 Bioanalyzer (Agilent Technologies, Inc.) electropherogram of total RNA isolated from magnetic bead sorted dopaminergic neurons. The electropherogram reveals excellent RNA quality and integrity suitable for a wide range of downstream applications including microarray analyses and qPCR as indicated by the presence of the 5.8S, 18S, and 28S rRNAs.

2. It is important to pre-cool the PBS used for dissection of fly heads. Placing the bottle of PBS covered in crushed ice for an hour before the experiment starts can do this. Alternatively, PBS can be stored at 4°C overnight. 3. Pre-cool the tissue grinder/pestle by placing it on ice for a few minutes to prevent cell damage/lysis during tissue homogenization. 4. The volume of antibody–bead mixture prepared depends on the number of CD8a-positive cells to be isolated given that 1 mg of beads can bind up to 20 mg of biotinylated antibody. Once prepared, it is recommended that the antibody–bead mixture be used within 4 weeks. 5. This protocol describes a high-throughput method of isolating specific neuronal cells from the Drosophila brain beginning with intact heads. We find that this method is highly efficient and isolation of single cell suspensions is not impeded by the head cuticle, which is filtered out when the sample is run through the 30 mm filter. Alternatively, if desired, the fly brains can be dissected according to the method described (15) prior to dissociation, however this will substantially increase the time required during the dissection stage of the protocol. 6. It is important to homogenize the tissue slowly and steadily without any sudden movements, to avoid unwanted stress and damage to the brain cells.

13

Cell-Specific Isolation to Study Dopamine Signaling in Drosophila

225

7. Trituration must be performed slowly, as forceful trituration may damage cells. 8. It is important to note that during the incubation of the antibody step, high temperatures or long incubation periods can result in a loss of antibody specificity thereby compromising purity in the isolation procedure.

Acknowledgments The authors thank the Bloomington Stock Center for providing Drosophila transgenic strains and Dr. Nadine Kabbani for contributions to this protocol. This work was supported by the Thomas F. and Kate Miller Jeffress Memorial Trust and NIH MH086928 (DNC). References 1. Kim YC, Lee HG, Han KA (2007) D1 dopamine receptor dDA1 is required in the mushroom body neurons for aversive and appetitive learning in Drosophila. J Neurosci 27:7640–7647 2. Selcho M, Pauls D, Han K-A, Stocker RF, Thum AS (2009) The role of dopamine in Drosophila larval classical olfactory conditioning. PLoS One 4:e5897. doi:10.1371/journal.pone.0005897 3. Ward A, Walker VJ, Feng Z, Xu XZS (2009) Cocaine modulates locomotion behavior in C. elegans. PLoS One 4:e5946. doi:10.1371/ journal.pone.0005946 4. Lebestky T, Chang JS, Dankert H, Zelnik L, Kim YC, Han KA, Wolf FW, Perona P, Anderson DJ (2009) Two different forms of arousal in Drosophila are oppositely regulated by the dopamine D1 receptor ortholog DopR via distinct neural circuits. Neuron 64:522–536 5. Bainton RJ, Tsai LT-Y, Singh CM, Moore MS, Neckameyer WS, Heberlein U (2000) Dopamine modulates acute responses to cocaine, nicotine and ethanol in Drosophila. Curr Biol 10:187–194 6. Blenau W, Baumann A (2001) Molecular and pharmacological properties of insect biogenic amine receptors: lessons from Drosophila melanogaster and Apis mellifera. Arch Insect Biochem Physiol 48:13–38 7. Wolf FW, Heberlein U (2003) Invertebrate models of drug abuse. J Neurobiol 54:161–178 8. McClung C, Hirsh J (1998) Stereotypic behavioral responses to free-base cocaine and the

9.

10.

11.

12.

13.

14.

15.

development of behavioral sensitization in Drosophila. Curr Biol 8:109–112 Rothenfluh A, Heberlein U (2002) Drugs, flies, and videotape: the effects of ethanol and cocaine on Drosophila locomotion. Curr Opin Neurobiol 12:639–645 Heberlein U, Wolf FW, Rothenfluh A, Guarnieri DJ (2004) Molecular genetic analysis of ethanol intoxication in Drosophila melanogaster. Integr Comp Biol 44:269–274 Heberlein U, Tsai LT, Kapfhamer D, Lasek AW (2009) Drosophila, a genetic model system to study cocaine-related behaviors: a review with focus on LIM-only proteins. Neuropharmacology 56:97–106 Iyer EPR, Iyer SC, Sulkowski MJ, Cox DN (2009) Isolation and purification of Drosophila peripheral neurons by magnetic bead sorting. J Vis Exp 34:e1599. doi: 10.3791/1599 (2009). Wang X, Bo J, Bridges T, Dugan KD, Pan T-C, Chodosh L, Montell DJ (2006) Analysis of cell migration using whole genome expression profiling of migratory cells in the Drosophila ovary. Dev Cell 10:483–495 Friggi-Grelin F, Coulom H, Meller M, Gomez D, Hirsh J, Birman S (2003) Targeted gene expression in Drosophila dopaminergic cells using regulatory sequences from tyrosine hydroxylase. J Neurobiol 54:618–627 Wu JS, Luo L (2006) A protocol for dissecting Drosophila melanogaster brains for live imaging or immunostaining. Nat Protoc 1:2110–2115

Part IV Electrochemical, Physiological, and Behavioral Analysis

Chapter 14 Regulation of Dopamine Transporter Expression by Neuronal Activity Shalini Padmanabhan, Thach Pham, and Balakrishna M. Prasad Abstract Actions of extracellular dopamine released in the central nervous system are primarily terminated by the dopamine transporter. This protein is also a target for therapeutic and abused psychostimulant drugs. Different methods used to study dopamine transporter function, its expression, and intracellular signaling in neurons are described in this chapter. Function of the dopamine transporter in mesencephalic primary cultures can be measured by dopamine uptake assay. Expression of the transporter protein and mRNA are analyzed by western blots and quantitative RT-PCR, respectively. Finally, methods to study neuronal activity-dependent changes in Ca2+⁄calmodulin-dependent protein (CaM) kinase activity in dopamine neurons are described. Key words: Mesencephalic, Dopamine, Uptake, Transporter, Western blot, Quantitative RT-PCR, Action potential, Tyrosine hydroxylase, CaM kinase, MAP kinase

1. Introduction The primary physiological function of the dopamine transporter is removal of dopamine from the extracellular space. Dopamine transporter activity can shape the anatomical distribution and duration of action of dopamine released from nerve terminals (1, 2). Blockade of dopamine transporter by psychostimulant drugs can lead to an enhanced activation of dopamine receptors and result in overt behavioral changes (3, 4). There is significant interindividual variability in dopamine transporter expression and this is associated with measures of attention in humans (5, 6). Furthermore, pharmacological and environmental factors can reversibly alter the abundance of dopamine transporter (7–9). However, mechanisms

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_14, © Springer Science+Business Media, LLC 2013

229

230

S. Padmanabhan et al.

regulating the expression of dopamine transporter are poorly understood. We discovered that neuronal activity is one of the factors that can regulate dopamine transporter expression (10, 11). Calcium and CaM kinase II signaling mediate neuronal activity-dependent changes of dopamine transporter expression (10). Primary cultures of dopamine neurons are the best experimental system to study neuronal activity-dependent regulatory mechanisms. Heterologous expression of cDNA constructs is typically used to study uptake activity of dopamine and other monoamine transporters. These expression systems offer the advantages of easy data interpretation, availability of molecular tools, and feasibility of structure–function analysis. However, the intracellular environment, electrical activity, and signaling machinery in heterologous expression systems such as HEK 293, MDCK, or HeLa cells are significantly different from that of dopamine neurons. Ex vivo preparations such as synaptosomes provide an alternative to overcome these limitations. However, ex vivo preparations are not suitable to perform longterm experiments, especially those involving gene expression changes. Thus, primary cultures are the best available option at present to reliably study changes in dopamine transporter expression.

2. Materials 2.1. Mesencephalic Primary Cultures

1. Time-mated female Sprague Dawley rats (Harlan, Indianapolis, IN) are typically obtained at 14–16 day gestation. 2. Ketamine HCl/Ketaset (10 mg/mL, Fort Dodge Laboratories, Fort Dodge, IA). 3. Dissociation medium salt concentrate (5×): 2,900 mM NaCl, 135 mM KCl, 650 mM NaHCO3, 100 mM NaH2PO4 H2O, 20 mM MgSO4, 25 mM EDTA, and 1200 mM d-glucose made in water and stored at 4°C (see Note 1). 4. Cysteine water: 1.25 mM cysteine in water supplemented with 1.9 mM CaCl2. 5. Papain (Worthington Biochemicals, Lakewood, NJ) is activated by adding it to cysteine water (20 units/mL) (see Note 2). 6. Dissociation medium is prepared by adding dissociation medium salt concentrate (1×) and kynurenate (0.5 mM) to activated papain solution. pH is adjusted to 7.2–7.4. 7. Carbogen cylinder (95% oxygen and 5% carbon dioxide). 8. Poly-D-lysine (Sigma-Aldrich, St. Louis, MO): 10 mg/mL (100×) single-use aliquots are made up in Hank’s balanced salt solution (Invitrogen, Carlsbad, CA) and stored at −20°C. 1×

14

Dopamine Transporter Expression

231

(100 μg/mL) Poly-D-lysine is made by diluting 100× stocks in tissue-culture grade water, filter-sterilized and used fresh. 9. Laminin (Sigma-Aldrich): 1 mg/mL single-use aliquots are stored at −80°C. 5 μg/mL laminin is made up in minimum essential medium (Invitrogen, Carlsbad, CA) and used fresh. 10. 12-well, 24-well, and 48-well tissue culture plates (Costar). These plates are coated with poly-d-lysine by incubating in 1× solution for 1 h at room temperature. The wells are washed twice with sterile water and laminin solution is added to the wells for 1 h. Dissociated cells are plated immediately after aspirating the laminin solution out of the plates. 48-Well plates are typically used for uptake assays, while 24-well and 12-well plates are used for western and RT-PCR assays, respectively. 11. Glial medium: Minimum essential medium with 10% new-born calf serum (Invitrogen), 0.45% D-glucose, 5 pg/mL insulin, 0.5 mM glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin. All media components are added to minimum essential medium and filter sterilized. This medium is used to maintain cultures of cortical glia and for initial plating of mesencephalic cells. 12. Neuronal medium: 50% minimum essential medium, 39% Ham’s-F12 medium, 10% heat-inactivated horse serum (Invitrogen), 1% heat-inactivated fetal bovine serum (Invitrogen), 0.45% d-glucose, 5 pg/mL insulin, and 0.1 mg/ mL apotransferrin (ICN Biomedicals, Irvine, CA). 13. Conditioned neuronal medium: 25 mL of neuronal medium is added to cortical glia grown in T75 tissue culture flasks. After overnight conditioning, the media is collected and supplemented with 1 ng/mL glial cell line-derived neurotrophic factor (1 ng/μL stock solution is made in 5% BSA water and stored as single-use aliquots at −80°C) and 500 μM kynurenate (0.5 M stock solution is made in 1 N NaOH and stored at −80°C as single-use aliquots). The medium is filter-sterilized and can be stored at −80°C, if not used immediately (see Note 3). 14. Flea micro stir bar (Fisher). 15. Sterile petri dishes (Fisher). 16. Tetrodotoxin (1 mM, PH 4–5) and 4-aminopyridine (100 mM) (Sigma-Aldrich) are made up in PBS, filter-sterilized and stored as single-use aliquots at −80°C. 2.2. Dopamine Uptake Assay

1. Ringer’s solution: 130 mM NaCl, 3 mM KCl, 1.2 mM MgSO4, 1 mM CaCl2, 10 mM d-glucose, and 1 mM ascorbic acid, pH 7.4 (one 50 mL tube warmed to 37°C and another tube chilled over ice).

232

S. Padmanabhan et al.

2. (2, 5, 6 3H)-dopamine (GE Healthcare, Piscataway, NJ) is diluted to appropriate concentration in warm Ringer’s solution before use. 3. Unlabeled dopamine stocks (10 mM) are made up in Ringer’s solution and stored at −80°C. For determining uptake kinetics, 4.05 μM dopamine is made with 20% radiolabeled and 80% unlabeled dopamine. Other dopamine concentrations are obtained by serial dilution of the 4.05 μM dopamine stock. 4. 3% trichloroacetic acid in water. 5. GBR12909 (Tocris Cookson, Ellisville, MO) is dissolved to 1 mM in water and stored at −80°C. 6. ScintiVerse scintillation liquid (Fisher Scientific, Pittsburgh, PA). 2.3. Western Blots

1. 12% Polyacrylamide gels (Bio-Rad, Hercules, CA). 2. Electrophoresis buffer: 25 mM Tris base, 192 mM glycine, and 0.5% SDS. 3. Electroblotting buffer: 20 mM Tris, 150 mM glycine, and 20% methanol. Adjust pH to 8.0 before adding methanol. 4. Sample buffer 6×: 60 mM TRIS, 1 mM EDTA, 2% SDS, 10% glycerol, 100 mM DTT, and 0.004% bromophenol blue. 5. Nitrocellulose membrane (0.45 μm NitroPure; Fisher Scientific, Pittsburgh, PA). 6. Ponceau S solution (0.1% in 5% acetic acid). 7. Prestained molecular Marker (Bio-Rad). 8. Blocking buffer: 5% nonfat dry milk in PBST. 9. Wash buffer (PBST): 1× PBS with 0.1% Tween-20. 10. SuperSignal West Pico Chemiluminescent Substrate (Pierce, Rockford, IL). 11. RIPA buffer: 50 mM Tris (pH 6.8), 150 mM NaCl, 4 mM EDTA, 100 mM sodium fluoride, 1 mM sodium orthovanadate, 1% sodium deoxycholate, 0.1% SDS, 1% Triton-100×. A Complete mini protease Inhibitor tablet (Roche Diagnostics, Indianapolis, IN) is added to 10 mL buffer, right before use. 12. Stripping buffer (Pierce). 13. Primary antibodies: (a) Rabbit polyclonal dopamine transporter antibody (10). (b) Monoclonal TH antibody (clone TH16, Sigma-Aldrich). (c) β-Actin antibody (goat polyclonal, Biotechnology, Santa Cruz, CA).

Santa

Cruz

(d) Phospho-CaM kinase II (Thr286/7) and CaM kinase II antibodies (Cell Signaling, Danvers, MA). (e) Phospho-TH (Ser19) antibody (Invitrogen).

14

Dopamine Transporter Expression

233

14. HRP-conjugated secondary antibodies: (a) Goat Anti-Mouse (Jackson ImmunoResearch Laboratories, West Grove, PA) (b) Donkey Anti-Rabbit (Jackson ImmunoResearch) 2.4. Quantitative RT-PCR

1. Trizol reagent (Invitrogen). 2. Chloroform. 3. Isopropanol. 4. 75% Ethanol. 5. RNAse-free water (Invitrogen). 6. Superscript III reverse transcriptase kit (Invitrogen). 7. SYBR® Green PCR supermix (Bio-Rad). 8. Bio-Rad iQ iCycler. 9. iQ 96-Well PCR Plates (Bio-Rad). 10. Optical Sealing Tape (Bio-Rad). 11. Primers (Integrated DNA Technologies, Coralville, IA): DAT (F)—5¢-TGGGCCTCAATGACACCTTT-3¢ DAT (R)—5¢-AGCAGAACAATGACCAGCACCA-3¢ Actin (F)—5¢-TTGCTGACAGGATGCAGAAGGAGA-3¢ Actin (R)—5¢-TAGAAGCATTTGCGGTGCACGATG-3¢

3. Methods 3.1. Mesencephalic Primary Cultures

As described above, mesencephalic primary cultures are the best available experimental system to study the regulation of dopamine transporter expression. Currently available cell-lines of dopamine neurons do not express appreciable amount of dopamine transporter. We adapted the dopamine neuronal culture methods originally described by Rayport et al. (12), for measuring dopamine transporter function and expression. 1. 2–4-day-old Sprague-Dawley rat pups are anesthetized with intraperitoneal injection of ketamine (3 mg per pup). 2. Brains are aseptically dissected out and placed in a sterile petri dish with Ringer’s solution. Approximately 3 mm thick coronal sections of mid-brain just caudal to hypothalamus are dissected. Ventral one-third of the coronal sections are cut into two pieces and placed in the dissociation medium (see Note 4). 3. Ventral mid-brain sections in dissociation medium are gently stirred using a micro-stir bar and stir plate. The vial is placed in

234

S. Padmanabhan et al. Carbogen

Vent 18g Needle Thermometer 0.2 µm Filter

Water

Hot Plate Water Micro Stir Bar

Fig. 1. Tissue dissociation setup. Carbogen is humidified via sterile water in a conical flask and passed through a 0.2 μm filter into dissociation medium. Dissociation vial with mesencephalic tissue is placed in a water bath maintained at 35–37°C. Tissue in dissociation vial is continuously bubbled with humidified carbogen and slowly stirred using fleabar for 2 h.

a 37°C water bath and tissue is constantly bubbled with carbogen (Fig. 1). 4. After 2 h of incubation, the tissue sections are transferred into a sterile 15 mL conical tube. Sections are dissociated using 10 mL plastic serological pipette, then by fire polished glass Pasteur pipette. Complete dissociation of tissue is ensured by the absence of visible tissue mass. 5. Dissociated cells are pelleted by centrifuging at 1,000 × g for 10 min at 4°C. 6. Dissociation medium is aspirated out and cells are resuspended in glial medium. Cell number is calculated using hemocytometer and the medium is diluted to obtain 300,000 cells per mL. 7. Cells are plated in 48-well plates previously coated with polyd-lysine and laminin. 150,000 cells are plated per well using repeat pipetter to minimize inter-well variability in cell density. For western blots 300,000 cells are plated per well in 24-well plates and for quantitative RT-PCR assays 600,000 cells are plated in 12-well plates. 8. An hour after plating, glial medium is aspirated and replaced with conditioned neuronal medium. 9. After 7 days in vitro, medium is changed to appropriate treatment medium (with vehicle or drugs). 10. Different assays for dopamine transporter function and abundance are performed as described below.

14

3.2. Dopamine Uptake Assay

Dopamine Transporter Expression

235

The primary function of dopamine transporter is the reuptake of extracellular dopamine. The simplest and most commonly used assay to measure dopamine transporter function is to assess the intracellular accumulation of radiolabeled dopamine. Translocation of substrates by dopamine transporter is analogous to enzymatic action of converting substrates into products. Thus, kinetics of substrate translocation are typically analyzed using Michaelis-Menten equation. Dopamine accumulation is typically measured at multiple concentrations (with the highest concentration being about tenfold greater than expected apparent affinity of dopamine, Km). 1. Cells are washed once in Ringer’s solution. 2. Radiolabeled dopamine in a total volume of 300 μL is added to wells. Five different concentrations of tritiated dopamine (0.05, 0.15, 0.45, 1.35, and 4.05) are used in duplicate wells (see Note 5). 3. After 2 min of incubation at room temperature, uptake is terminated by washing twice with 500 μL ice-cold Ringer’s solution (see Note 6). 4. Tritiated dopamine is extracted into 300 μL of 3% trichloroacetic acid by incubating 30 min at room temperature. 5. Trichloroacetic acid samples are transferred to scintillation vials and 5.0 mL of scintiverse fluid is added. 6. Radioactivity in each tube is measured by a liquid scintillation counter. 7. Nonspecific dopamine accumulation at each concentration (0.05–4.05 μM) is determined by incubating in the presence 10 μM GBR12909. Specific uptake is calculated as the difference between total (in absence of GBR12909) and nonspecific uptake. 8. Data are analyzed by the equation V = Vmax· (DA)/(Km + (DA)) using PRISM (GraphPad, La Jolla, CA). V is the velocity of uptake at a given concentration of dopamine (DA). Vmax represents maximal velocity, while Km is the apparent affinity of the substrate to transporter (see Fig. 2).

3.3. Measuring Dopamine Transporter Abundance

Dopamine neurons in culture conditions described above are tonically active and fire action potentials at an average rate of 2.3 Hz (13). Their firing can be abolished in the presence of 1 μM tetrodotoxin (TTX) (a sodium channel blocker) or increased by 1 mM 4-aminopyridine (a potassium channel blocker). In order to test the effect of neuronal activity on dopamine transporter function and abundance, we maintained mesencephalic cultures in the presence of control, TTX or 4-aminopyridine media for 5 days. Dopamine uptake data shown in Fig. 3 indicate that TTX decreases dopamine uptake while, 4-aminopyridine increases uptake.

S. Padmanabhan et al. 12

pmoles / well

10 8 6 4 2 0 0.01

0.1

1

10

Dopamine [µM] Fig. 2. Dopamine uptake kinetics in mesencephalic cultures. Picomoles of dopamine accumulated per well in 2 min at different substrate concentrations are shown. Vmax of dopamine uptake is11.6 ± 0.6 pmol/well and Km is 0.42 ± 0.08 μM. Average data from four independent experiments are shown.

4-AP 140 Dopamine Uptake (% control)

236

Control

*

TTX

120 100

*

80 60 40 20 0

3

4 30minutes

6

6 5 days

Fig. 3. The effect of neuronal activity on dopamine uptake in mesencephalic cultures. Uptake of 100 nM 3 H-dopamine in cultures treated with 1 μM TTX or 1 mM 4-aminopyridine (4-AP) for 30 min or 5 days is shown as percentage of control. n = 3–6 as indicated. *P < 0.05, compared with control. Adapted from (10) with permission.

In order to determine if these changes in dopamine uptake are accompanied by changes in dopamine transporter abundance, we analyzed cell lysates using western blots. A typical western blot assay is described below. 1. After appropriate treatments, cultures are washed once in Ringer’s solution. 2. Cells are extracted into 200 μL RIPA buffer by gentle agitation of culture plate on a shaker.

14

Dopamine Transporter Expression

237

3. Cell lysates are mixed with 6× sample buffer and run on 10% polyacrylamide gels. A molecular weight marker is run in the first lane of the gel. This will be used to ensure appropriate molecular mass of the protein of interest and also to orient the samples when they are transferred to the membrane. 4. Protein samples resolved on the gels are transferred to nitrocellulose membrane using 600 mA-h of power (15 h at 40 mA or 3 h at 200 mA). 5. Membranes with protein samples are stained with Ponceau-S by incubating in the staining solution for 5 min and then washing in water. 6. Sample lanes are labeled and the molecular weight bands are marked. 7. Membrane is incubated in blocking buffer for 30 min at room temperature. 8. Dopamine transporter antibody is diluted in the blocking buffer (1:1,000) and the membrane blot is incubated in the antibody solution for 1 h at room temperature. 9. Blot is washed four times in PBST with 5 min per wash. 10. Blot is incubated in an HRP-conjugated anti-rabbit secondary antibody (1:10,000 dilution) for 1 h at room temperature. 11. After five washes in PBST, a chemiluminescent substrate solution is added to the blot. 12. After 1 min of incubation, the chemiluminescence from the blot is captured by using an X-ray film or an imaging system (Kodak in vivo Fx Pro) (see Note 7). 13. Intensities of the bands corresponding to the dopamine transporter are analyzed by NIH image. 14. Data can be expressed as a percentage of control within each experiment. β-Actin is used as an internal control to normalize for inter-sample variability in protein concentration. 3.4. Measuring Dopamine Transporter mRNA Abundance

Data presented in Fig. 4 show that dopamine transporter abundance is altered by neuronal activity. In order to determine if these changes are accompanied by changes in dopamine transporter mRNA abundance, we used quantitative RT-PCR assay. 1. After appropriate treatments, cultures are washed once in Ringer’s solution. 2. Cells are extracted into 0.5 mL Trizol reagent by passing several times through a pipette. Samples are incubated for 5 min at room temperature (see Note 8). 3. 100 μL of chloroform is added per sample, shaken vigorously, and incubated at room temperature for 3 min.

238

S. Padmanabhan et al. 4-AP

a

Control

DAT

TH

b-Actin

TTX

Control

4-AP

Densitometric Units (% control)

b

150

Control

*

TTX

125 100

* 75 50 25 0 DAT

TH

Actin

Fig. 4. The effect of chronic changes in neuronal activity on dopamine transporter abundance. (a) Representative images of western blots from control, TTX- or 4-AP-treated cultures that were probed with dopamine transporter (DAT), tyrosine hydroxylase (TH), and β-actin antibodies. (b) Densitometric analysis of bands corresponding to DAT, TH, and β-actin. Data from NIH image analysis were normalized to respective control values (n = 5). *P < 0.05, compared with control cultures. Adapted from ref. (10) with permission.

4. Samples are centrifuged at 12,000 × g for 15 min at 4°C. Upper aqueous phase is collected into a new microfuge tube and 250 μL isopropyl alcohol is added. Samples are incubated at room temperature for 10 min. 5. Tubes are centrifuged at 12,000 × g for 10 min at 4°C. Supernatant is removed and RNA pellet is gently washed by adding 0.5 mL 75% ethanol. 6. The pellet is dissolved in RNAse-free water and RNA concentration of each sample is measured by a spectrophotometer at 260 nm wavelength. 7. 2 μg of total RNA is reverse transcribed by Superscript III assay kit, using oligo-dT primers. 8. 20 μL of RT samples are diluted to 100 μL using RNAse-free water. 9. Diluted RT samples (10 μL), SYBR GreenII mastermix (12.5 μL), and gene-specific primers (2.5 μL) are added to the wells in PCR plate. 10. PCR reactions are carried out in Bio-Rad iQ iCycler using the following parameters; Initial denaturation for 4 min at 95°C. 40 PCR cycles are performed with 95°C melting, 57°C annealing, and 72°C extension, with 30 s for each step (see Note 9). 11. Threshold cycles (CT) are calculated using baseline subtracted curves. 12. Fold change in mRNA abundance is calculated by comparative threshold cycle method (Scmittgen and Livak, 2008). Briefly,

14

b

239

3 DAT mRNA fold change

a

Dopamine Transporter Expression

* 2

1

* 4

6

4C AP on tr ol TT X

0

Fig. 5. Effects of neuronal activity on dopamine transporter mRNA abundance. (a) RT-PCR assay validation: Different volume of pooled cDNA samples that correspond to 0.05, 0.1, 0.2, 0.4, and 0.8 μg of total RNA were used to generate DAT and actin threshold cycles (CT). Standard curve graphs were plotted with RNA (μg) concentration and threshold cycles. Insets in these graphs are curves of relative fluorescence units (of cDNA samples corresponding to 0.05, 0.1, 0.2, 0.4, and 0.8 μg of RNA) plotted against PCR cycle numbers. The correlation coefficients for DAT and actin standard curves are greater than 0.99. The data are best fit to the following equations: DAT: CT = −3.189 log (μg RNA) + 20.98 and Actin: CT = −3.172 log (μg RNA) + 10.56. Amplification efficiencies defined as product increase per cycle were calculated by 10(−1⁄slope). PCR efficiency for DAT and actin were 2.058 and 2.066, respectively. These data demonstrate that amplification efficiency for DAT and actin are similar and are close to 100% (102.9 and 103.3%, respectively). (b) Changes in DAT mRNA abundance normalized to actin mRNA are expressed as fold change compared to control cultures. N values of RT-PCR assay data are shown in the graph. *P < 0.05, compared with control cultures. Adapted from ref. (10) with permission.

dopamine transporter CT values of each sample are subtracted from actin CT values to get delta CT values. The difference (delta-delta CT value) between treatment delta CT values (TTX or 4-AP) and control delta CT values is used to calculate the fold change. Fold change = 2(delta-delta CT value) (see Fig. 5, Note 10). 3.5. Measuring CaM Kinase II Activity in Dopamine Neurons

CaM kinase II and mitogen-activated protein (MAP) kinase are two major signaling molecules that mediate activity dependent changes in neurons (14, 15). Using pharmacological inhibitors we identified that CaM kinase II, but not MAP kinase is involved in activity-dependent changes in dopamine transporter expression (10). We developed assays for direct measurement of the activity of these two kinases in mesencephalic cultures (10, 11). Dopamine neurons are the only cells that express tyrosine hydroxylase in our culture conditions. Tyrosine hydroxylase can be phosphorylated at Ser19 by CaM kinase II and by MAP kinase at Ser31 (16). We used western blot assays to measure the phosphorylation state of TH at these two sites as a read out of CaM kinase and MAP kinase. Data presented in Fig. 6 show that phosphorylation of tyrosine hydroxylase (Ser19) can be used to monitor neuronal activity-dependent changes in CaM kinase II activity in dopamine neurons.

S. Padmanabhan et al.

CaM KII Activation

Phospho-CaMK II

TH Phospho-TH CaM KII Activity

p-TH

TTX

*

150 100

* 50

* 0 II

CaMKII

TTX

Control

*

200

K

p-CaMKII

Control

4-AP

aM

4-AP

250

p-

C

aM

K

II/

C

TH

/T H

CaM K II

b

pTH

a

Densitometric Units (% control)

240

Fig. 6. Effect of neuronal activity on CaM kinase II activity in dopamine neurons. (a) Representative immunoblots showing the effects of a 30 min treatment with 1 μM TTX and 1 mM 4-AP on phospho-CaM kinase II, CaM kinase II, phospho-TH, and TH. (b) Densitometric analysis shows that ratios of phospho-CaM kinase II:CaM kinase II (a measure of CaM kinase II activation) and phospho-TH:TH (a measure of CaM kinase II activity in dopamine neurons) are altered by TTX and 4-AP (n = 5). *P < 0.05, compared with control cultures. Modified from ref. (10).

4. Notes 1. Unless otherwise noted, all reagents are obtained from SigmaAldrich. 2. Papain activity expressed in units per milligram protein varies between lots. The protein content per unit volume also varies between lots. Both these factors need to be used in determining the volume of enzyme stock to be used. 3. Cortical glia is cultured from postnatal rat pups using the same protocol described for mesencephalic cultures. Neurons in these cultures are eliminated by feeding glial medium containing 100 μM l-glutamate for 24 h on the day of plating. 4. Correctly dissected mid-brain sections have a visible groove (mid-brain flexure) in the middle of the ventral surface. Proper dissection ensures the presence of sufficient dopamine neurons without any norepinephrine neurons. Presence of norepinephrine neurons can potentially be a major problem as these neurons can accumulate dopamine and contain tyrosine hydroxylase. An easy way to ensure lack of norepinephrine neurons in culture is to test the susceptibility of dopamine uptake to 100 nM GBR12909 (dopamine transporter inhibitor)

14

Dopamine Transporter Expression

241

and 100 nM desipramine (norepinephrine transporter inhibitor) (13). 5. A commonly used method to obtain different substrate concentrations for kinetic analysis is to use a low (100 nM) concentration of radiolabeled dopamine and dilute it with unlabeled dopamine to obtain desired concentration. Quantity of radiolabeled dopamine accumulated is multiplied by the dilution factor to obtain an estimate of total dopamine accumulated at each concentration. An alternative method is to use radiolabeled dopamine as a fixed proportion of the total dopamine. In the uptake assay described above 20% of total dopamine is radiolabeled. Although the latter method requires the use of more radiolabeled substrate, the data obtained are likely to be more accurate. This is because the data at the high end of concentration range are not multiplied by large dilution factors, as in the commonly used method. 6. To obtain accurate transport kinetic parameters, it is important to ensure that uptake of highest substrate concentration used is linear within the duration of uptake assay. Uptake of 4.05 μM dopamine is linear with time for at least 4 min. When single low concentration of tritiated dopamine (100 nM) is used uptake can be carried out for 4–6 min. 7. When optimizing the imaging conditions, western blots of serially diluted samples with different exposure times can be collected. Appropriate exposure timing is important to ensure that the bands corresponding to different samples are in the dynamic range of image analyzing program used. X-ray film images can be digitized by commercially available scanners. 8. RNAse-free pipette tips and nuclease-free molecular grade reagents are critical for this assay. Samples and reagent vials are to be handled using gloves. 9. Agarose gel analysis of PCR products can be used to confirm that only one product of predicted size is amplified. 10. The underlying assumption of this method is that different PCR products are amplified with equal efficiency. Amplification efficiency for each PCR product can be measured by using serial dilution of pooled cDNA samples from different cultures (Fig. 5). References 1. Gainetdinov RR, Jones SR, Fumagalli F, Wightman RM, Caron MG (1998) Re-evaluation of the role of the dopamine transporter in dopamine system homeostasis. Brain Res Rev 26:148–153 2. Benoit-Marand M, Jaber M, Gonon F (2000) Release and elimination of dopamine in vivo in

mice lacking the dopamine transporter: functional consequences. Eur J Neurosci 12:2985–2992 3. Di Chiara G, Imperato A (1988) Drugs abused by humans preferentially increase synaptic dopamine concentrations in the mesolimbic system of freely moving rats. Proc Natl Acad Sci USA 85:5274–5278

242

S. Padmanabhan et al.

4. Volkow ND, Wang G, Fowler JS, Logan J, Gerasimov M, Maynard L, Ding Y, Gatley SJ, Gifford A, Franceschi D (2001) Therapeutic doses of oral methylphenidate significantly increase extracellular dopamine in the human brain. J Neurosci 21(2):RC121, 1–5 5. Newberg A, Amsterdam J, Shults J (2007) Dopamine transporter density may be associated with the depressed affect in healthy subjects. Nucl Med Commun 28:3–6 6. Volkow ND, Wang GJ, Newcorn J, Fowler JS, Telang F, Solanto MV, Logan J, Wong C, Ma Y, Swanson JM, Schulz K, Pradhan K (2007) Brain dopamine transporter levels in treatment and drug naive adults with ADHD. Neuroimage 34:1182–1190 7. Dresel S, Krause J, Krause KH, LaFougere C, Brinkbaumer K, Kung HF, Hahn K, Tatsch K (2000) Attention deficit hyperactivity disorder: binding of (99mTc)TRODAT-1 to the dopamine transporter before and after methylphenidate treatment. Eur J Nucl Med 27:1518–1524 8. Feron FJ, Hendriksen JG, van Kroonenburgh MJ, Blom-Coenjaerts C, Kessels AG, Jolles J, Weber WE, Vles JS (2005) Dopamine transporter in attention-deficit hyperactivity disorder normalizes after cessation of methylphenidate. Pediatr Neurol 33:179–183 9. Bezard E, Dovero S, Belin D, Duconger S, Jackson-Lewis V, Przedborski S, Piazza PV, Gross CE, Jaber M (2003) Enriched environment confers resistance to 1-methyl-4-phenyl1,2,3,6-tetrahydropyridine and cocaine:

involvement of dopamine transporter and trophic factors. J Neurosci 23:10999–11007 10. Padmanabhan S, Lambert NA, Prasad BM (2008) Activity-dependent regulation of the dopamine transporter is mediated by CaM kinase signaling. Eur J Neurosci 28:2017–2027 11. Padmanabhan S, Prasad BM (2009) Sustained depolarization decreases calcium/calmodulindependent protein kinase II activity and gene expression in dopamine neurons. Neuroscience 163:277–285 12. Rayport S, Sulzer D, Shi WX, Sawasdikosol S, Monaco J, Batson D, Rajendran G (1992) Identified postnatal mesolimbic dopamine neurons in culture: morphology and electrophysiology. J Neurosci 12:4264–4280 13. Prasad BM, Amara SG (2001) The dopamine transporter in mesencephalic cultures is refractory to physiological changes in membrane voltage. J Neurosci 21:7561–7567 14. Vaillant AR, Zanassi P, Walsh GS, Aumont A, Alonso A, Miller FD (2002) Signaling mechanisms underlying reversible, activity-dependent dendrite formation. Neuron 34:985–998 15. Kelleher RJ III, Govindarajan A, Jung HY, Kang H, Tonegawa S (2004) Translational control by MAPK signaling in long-term synaptic plasticity and memory. Cell 116:467–479 16. Haycock JW, Haycock DA (1991) Tyrosine hydroxylase in rat brain dopaminergic nerve terminals. Multiple-site phosphorylation in vivo and in synaptosomes. J Biol Chem 266:5650–5657

Chapter 15 Monitoring Axonal and Somatodendritic Dopamine Release Using Fast-Scan Cyclic Voltammetry in Brain Slices Jyoti C. Patel and Margaret E. Rice Abstract Brain dopamine pathways serve wide-ranging functions including the control of movement, reward, cognition, learning, and mood. Consequently, dysfunction of dopamine transmission has been implicated in clinical conditions such as Parkinson’s disease, schizophrenia, addiction, and depression. Establishing factors that regulate dopamine release can provide novel insights into dopaminergic communication under normal conditions, as well as in animal models of disease in the brain. Here we describe methods for the study of somatodendritic and axonal dopamine release in brain slice preparations. Topics covered include preparation and calibration of carbon-fiber microelectrodes for use with fast-scan cyclic voltammetry, preparation of midbrain and forebrain slices, and procedures of eliciting and recording electrically evoked dopamine release from in vitro brain slices. Key words: Dopamine, Brain slices, Voltammetry, Carbon-fiber microelectrodes, Striatum, Substantia nigra pars compacta, Ventral tegmental area

1. Introduction The principal dopamine (DA) pathways in the brain arise from midbrain nuclei designated as the A9 and A10 cell groups, which are the substantia nigra pars compacta (SNc) and ventral tegmental area (VTA), respectively (1). Dopamine neurons in these regions project ipsilaterally via the medial forebrain bundle (MFB) to various forebrain structures (Fig. 1a). Neurons of the SNc send axon projections via the nigrostriatal pathway to the dorsolateral striatum (caudate putamen, CPu), which is critically involved in the control of movement (2, 3). Neurons of the VTA project via the mesolimbic pathway to the ventromedial striatum (nucleus accumbens, NAc), which is involved in motivation and reward (4) and

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_15, © Springer Science+Business Media, LLC 2013

243

244

J.C. Patel and M.E. Rice

a

Forebrain slice

Midbrain slice

2 mm

cc CPu

NAc ac OT u

SN

mfb

lateral 2.4 mm

b

c cc

2 mm

cc CPu

NAc (core)

SNr

NAc (shell)

SNc VTA

OTu bregma +1.0 mm

bregma –4.8 mm

Fig. 1. Rat brain sections showing major forebrain and midbrain dopamine (DA) regions. (a) Sagittal section of rat brain, stained with cresyl violet, showing main forebrain (axon terminal) and midbrain (somatodendritic) regions from which DA release can be recorded. Lateral coordinate indicates distance from rat central suture, a landmark on the skull. Dashed lines show the relative positions at which coronal forebrain or midbrain slices are typically made. (b) Typical level of a coronal forebrain slice used to study axonal DA release. At this level, it is possible to record DA release from the dorsal striatum (CPu) as well as the ventral striatum (NAc, core and shell or OTu) in the same forebrain slice. (c) Typical level of a midbrain slice to study somatodendritic DA release. At this level, it is possible to record DA release in the SNc (A9 region) as well as the VTA (A10 region). Bregma coordinates indicate distance anterior (+) and posterior (−) to this landmark on the skull. Abbreviations: ac anterior commissure, Cer cerebellum, cc corpus callosum, CPu caudate putamen, mfb medial forebrain bundle, NAc nucleus accumbens, OTu olfactory tubercle, SN substantia nigra, SNc SN pars compacta, SNr SN pars reticulata, VTA ventral tegmental area (from ref. (60) with permission from Elsevier).

to the olfactory tubercle (OTu), which is involved in olfactory processing. Additionally, VTA DA neurons project via the mesocortical pathway to the amygdala, which is involved in the control of emotion, and to prefrontal cortex, which is involved in cognition, including working memory. Consequently, dysfunction of

15

Monitoring Dopamine Release

245

DA transmission has been implicated in several significant brain disorders, including the movement deficits of Parkinson’s disease, addiction to substances like alcohol and cocaine as well as natural rewards including food and sex, and the pathophysiology of schizophrenia and depression. Studies of DA release regulation using voltammetric methods with carbon-fiber microelectrodes have made significant contributions to the present understanding of DA regulation and role of DA in normal and pathological conditions (5). A number of key mechanistic insights have been obtained using these methods in brain slices in vitro, which readily permit voltammetric detection of DA release in discrete brain regions, without complicating factors inherent to in vivo studies, including animal behavior, anesthesia, and indirect effects from distant structures via long pathways (5, 6). The use of midbrain slices permits examination of somatodendritic DA release in the SNc and VTA, whereas the use of forebrain slices permits monitoring of axonal DA release in the dorsal striatum and nucleus accumbens shell and core. These techniques are used primarily to study spontaneous or stimulated DA release, rather than basal [DA]o for two main reasons. First, the rapid sampling time and small probe size are ideally suited for very localized monitoring of transient changes in extracellular DA concentration ([DA]o). Second, basal [DA]o is in the low nM range, which is below the detection limits for most carbon-fiber electrodes. This contrasts with another popular method for monitoring [DA]o, which is microdialysis. That method involves extraction of substances, like DA, from brain extracellular fluid that are then analyzed off-line using HPLC or other analytical methods. This extraction takes time, so that microdialysis is not suited for monitoring dynamic changes in [DA]o, but can be adapted to determine basal [DA]o. Therefore these methods provide complementary information about DA transmission. Moreover, other electroactive substances found in vivo, especially DOPAC, a DA metabolite, and ascorbate, an abundant extracellular antioxidant, are readily lost from the extracellular compartment of brain slices, so do not interfere with evoked DA detection in vitro. A variety of voltammetric methods have been developed; the three most commonly used are amperometry, chronoamperometry, and fast-scan cyclic voltammetry (FCV) (5). Each involves the measurement of current generated by the electrochemical oxidation of DA at a suitable recording electrode. The primary focus of this chapter is to describe methods and methodological issues required for monitoring DA release in brain slices using FCV and carbon-fiber microelectrodes. However, the basic methods can be used to monitor the other primary biogenic amines, serotonin (5-HT), and norepinephrine (NE).

246

J.C. Patel and M.E. Rice

2. Materials 2.1. Instrumentation for FCV

1. Millar Voltammeter (obtained by special request from Dr Julian Millar at St. Bartholomew’s and The Royal London School of Medicine and Dentistry, University of London, UK). 2. Digital storage oscilloscope. 3. Analogue to Digital (A-to-D) converter (e.g., Digidata A-to-D board, Molecular Devices, Foster City, CA, USA) with Clampex Software (Molecular Devices) for data acquisition. 4. External timing unit (e.g., Master-8, A.M.P.I., Jerusalem, Israel). 5. Personal computer.

2.2. Manufacture of Carbon-Fiber Microelectrodes

1. Bundle of unsized 7–8 mm diameter carbon fibers (Goodfellow Corporation, Berwyn, PA, USA). The fibers should be stored so that they are protected from dirt and dust in the atmosphere. 2. Borosilicate glass capillaries, 10 cm length, 2.0 mm outer diameter, 1.16 mm inner diameter (Part No GC200-10; Harvard Apparatus Ltd. or Part No B200-116-10; Sutter Instrument Company). 3. Tools for electrode fabrication, including fine forceps modified by covering each tip with a plastic insulation sleeve to permit handling of carbon fibers without crushing and scissors for trimming the carbon fibers to a length of about 15 cm. 4. Clean 13 × 80 mm glass test-tube angled at 30° and held with modeling clay on a glass slide or other solid surface. 5. Acetone (General grade, Fischer Scientific, Rockford, IL). 6. Vertical microelectrode puller (e.g., Model PE-2, Narishige Scientific Instruments, Japan). 7. Microscope with 20× eyepieces and 10× objective to view and cut the carbon fiber extending from the glass pipette after pulling. 8. Apparatus for cutting carbon-fiber tips, which includes two micromanipulators set at right angles to one another; one for holding the carbon-fiber electrode and the other to hold a mounted scalpel blade. 9. Woods metal (Goodfellow, Oakdale, PA), electrically conductive silver paint (e.g., Silver Print II, GC Electronics, Rockford, IL), silver epoxy (MG Chemicals, Surrey, BC, Canada), or 4 M K-acetate/0.15 M KCl solution for electrical contact between the carbon fiber and a connection wire. 10. Insulated electrical stranded copper wire (1 mm diameter) (e.g., Extraflexible Miniature Vinyl Insulated Lead Wire, 26–28 gauge, Cooner Wire, Chatsworth, CA).

15

Monitoring Dopamine Release

247

11. Gold-plated connector pin (1 mm) (e.g., model WC1-10, Harvard Apparatus, Holliston, MA) or small alligator clip to connect the electrode wire to the voltammeter headstage. 2.3. Testing and Calibrating CarbonFiber Microelectrodes

1. Dopamine (Sigma-Aldrich, St. Louis, MO) stock solution (2 mM in 0.1 M perchloric acid, Fisher). 2. 0.1 M nitric acid solution for weekly washing of glassware; this minimizes DA oxidation in solution. Be sure to wash out the acid afterwards with deionized water. 3. Bicarbonate-buffered artificial cerebrospinal fluid (aCSF) containing 124 mM NaCl, 3.7 mM KCl, 26 mM NaHCO3, 1.3 mM MgSO4, 1.3 mM KH2PO4, 10 mM glucose, 2.4 mM CaCl2 and saturated with 95% O2/5% CO2 (Sigma-Aldrich). 4. Uncoated silver wire to make Ag/AgCl reference electrodes (0.015 in. bare Ag wire; A–M Systems, Sequm, WA); 9 V battery and 0.1 M KCl solution for chloride plating.

2.4. Preparation of Brain Slices

1. Vibratome (e.g., Ted Pellar, Inc., Redding, CA) or a vibrating blade microtome (e.g., VT1200S, Leica Microsystems, Buffalo Grove, IL). 2. Platinum-edge razor blades (Gillette, Boston, MA). 3. Appropriate stereotaxic brain atlas for rodent species examined. 4. Dissection tools including scalpel blade, rongeurs, sharp scissors, fine iris scissors, fine forceps (e.g., Fine Science Tools, Foster City, CA), small spatula, glass Petri dish, and filter paper. 5. Ice-cold HEPES-bicarbonate-buffered artificial cerebrospinal fluid (aCSF) containing in mM NaCl (120), KCl (5), NaHCO3 (20), HEPES acid (6.7), HEPES salt (3.3), MgSO4 (2), glucose (10), CaCl2 (2) and saturated with 95% O2/5% CO2. 6. Brain slice holding chamber consisting of a 15 cm disposable Petri dish with lid (Fisher). The chamber is filled with HEPESbuffered aCSF at room temperature and receives a constant supply of 95% O2/5% CO2, which gently bubbles the aCSF via a 1 mm plastic tube inserted through a small hole made in the cover or lid.

2.5. Stimulating and Recording DA Release

1. Anti-vibration table (e.g., TMC, Peabody, MA) or a solid surface. 2. Dissection microscope (e.g., Olympus, Center Valley, PA). 3. Brain slice submersion recording chamber (model RC-26G with series 20 P-1 chamber platform, Warner Instruments from Harvard Apparatus, Holliston, MA). 4. Peristaltic pump (Minipuls 3, Gilson, Middleton, WI) for controlled perfusion of the brain slice chamber with aCSF. 5. Inline heater (SH-27B, Warner Instruments) with an automatic temperature controller (TC-324B, Warner Instruments, Hamden, CT).

248

J.C. Patel and M.E. Rice

6. Fiber optic cold-light source for slice illumination (e.g., Cuda Products Co., Jacksonville, FL). 7. Two micromanipulators for placement of the carbon fiber recording electrode and stimulating electrode (e.g., Narishige from Tritech Research, Los Angeles, CA). 8. Bipolar platinum or platinum/iridium stimulating electrode (twisted-wire or concentric) (e.g., Fredrick Haer & Co. (FHC), Bowdoinham, ME). 9. Stimulus isolator (Isoflex, A.M.P.I) with stimulus pulse duration and pulse-train frequency controlled by the Master-8 pulse generator (A.M.P.I).

3. Methods 3.1. Basic Principles and Instrumentation for FCV

“Fast-scan cyclic voltammetry” is a very descriptive name for this method to monitor the release of biogenic amines. The term “fast” refers to the scan rate, typically 300–900 V/s. The standard voltage waveform is triangular (Fig. 2a), beginning with a positive ramp from a negative starting potential (we use −0.7 V vs. Ag/ Ag/Cl) to a potential that exceeds that required to oxidize DA (we scan to +1.3 V vs. Ag/AgCl), followed by a negative going ramp that returns to the starting potential, passing the potential required to reduce just-oxidized DA en route. At a scan rate of 800 V/s (7, 8), each voltage cycle, or “scan” takes ~5 ms (Fig. 2). Scans are repeated continually throughout an experiment, typically at 100 ms intervals (or 10 Hz), which maintains a stable “background” current, discussed further below. The Millar Voltammeter takes the recording electrode out of circuit between scans, whereas other voltammeters, like the Ensman EI400, keep the electrode at a negative “holding” potential. Taking the electrode out of circuit helps maintain electrode sensitivity during prolonged recording, whereas a negative holding potential may enhance sensitivity by promoting biogenic amine adsorption, however, this may also slow the clearance phase release records. The last term, “voltammetry” refers to any method in which a voltage is applied and current is measured. In the case of DA, each molecule produces two electrons during oxidation, resulting in current flow through the conductive carbon-fiber microelectrode that is proportional to the concentration of DA in a calibration solution or released into the extracellular space of brain tissue. It should be noted that other key transmitters, glutamate, GABA, and acetylcholine (ACh) are not electroactive. During the positive voltage scan, current begins to flow as the DA oxidation potential is approached (Fig. 2b). Once a potential has been reached that is sufficient to oxidize all DA molecules at the electrode surface, the

15

a

Monitoring Dopamine Release Oxidation

/s

Reduction

80 0V

Eapp (V vs. Ag/AgCl)

+1.5

249

0

–1.0

b 25 nA

c 500 nA

Eapp (V vs. Ag/AgCl) Time (ms)

0

–0.7

1.3

–0.7

2.5

5

7.5

10

Fig. 2. Fast-scan cyclic voltammetry (FCV) voltage waveform and dopamine (DA) voltammograms. (a) Typical triangular voltage waveform applied to a carbon-fiber microelectrode with FCV. (b) Voltammogram for 5 mM DA obtained by subtracting the background current in aCSF alone from that obtained in the presence of DA in panel C. Electrode sensitivity to DA in this example was 7 nA/mM; oxidation and reduction peak potentials for DA were +580 mV and −190 mV vs. Ag/AgCl, respectively. (c) Background current generated at a carbon-fiber microelectrode with FCV in the absence (solid-line) and presence (broken line) of 5 mM DA.

local concentration of unoxidized DA is rapidly depleted, resulting in a current peak that is seen at the DA oxidation potential (typically ~0.6 V vs. Ag/AgCl) (Fig. 2b). A peak is seen with FCV but not with slow voltammetric scans because the scan rates used are faster than the rate of replenishment of DA from spherical diffusion to the electrode surface is depleted. When the voltage reverses, oxidized DA is then reduced, resulting in a reduction current peak (typically ~0.2 V vs. Ag/AgCl) (Fig 2b). The oxidation and reduction peak potentials are characteristic for DA (and other catecholamines) and can therefore be used to identify it as the monitored substance, at least in a DA-rich region like dorsal striatum (6, 9, 10).

250

J.C. Patel and M.E. Rice

The characteristics of rapid monitoring and signal identification have made FCV the method of choice for many in vitro as well as in vivo studies of endogenous DA release. There are some caveats to the use of this method, as well, however. 1. There is a high non-Faradaic background current in all FCV scans (11, 12), which is directly proportional to scan rate. The Faradaic (DA) oxidation current, on the other hand, is proportional to the square root of scan rate (13). 2. The background current is usually ~1 mA (Fig. 2c), whereas a typically 1 mM increase in extracellular DA concentration ([DA]o) generates oxidation currents that are in the low nA range at most carbon-fiber microelectrodes. Therefore this large background must be subtracted (Fig. 2c). This introduces a challenge in the analysis of FCV data, which can be handled either by the voltammetric instrument used (e.g., the Millar voltammeter) or by post hoc analysis, as discussed further below. Voltammetric instrumentation required for FCV can be purchased made in-house by a good electronics workshop using published circuit diagrams (14) or generated digitally. There are a few commercially available instruments that are specifically designed for FCV, including the Millar Voltammeter (e.g., (6–8)) and the Ensman EI 400 (most recently available through ESA, now part of Dionex, but discontinued). Alternatively, voltammetric waveform generation as well as data acquisition and analysis can be carried out on a PC-based system using two multifunction data acquisition cards and software written in LabView (National Instruments, Austin, TX, USA) (15). The Millar voltammeter has a convenient feature (a “refresh” button) that stores a background voltammogram immediately before initiating a calibration or stimulation, which is then subtracted from subsequent scans, and allows direct monitoring of DA voltammograms. A schematic diagram showing the basic instrumentation used in our laboratory for determination of electrically evoked DA release using FCV with a Millar voltammeter is shown in Fig. 3. The Millar voltammeter consists of a function generator (which produces a continuous triangular voltage waveform), a waveform gate (which dissects the voltage waveform into discrete segments), a potentiostat (which applies the voltage waveform to the recording electrode and measures the current generated), and amplifiers (to amplify the current output from the recording electrode). The voltammeter can be used in three-electrode mode with a platinum wire auxiliary electrode (6, 10), but is commonly used in two-electrode mode, with only a carbon-fiber microelectrode and Ag/AgCl reference. The third auxiliary electrode provides feedback about current flow during an electrochemical measurement; this is used to maintain the applied potential when large current flow might otherwise lead to a voltage drop at the time of an oxidation peak. However, this is not a

15

Monitoring Dopamine Release

251

Oscilloscope

Millar Voltammeter

Microscope

Master-8 Pulse Generator

Solution Pump

Heater

Stimulus Isolator aCSF A-to-D Interface

Computer Cold-Light Source

Fig. 3. Schematic diagram showing the basic instrumentation for detection of dopamine release using FCV in a brain slice. A brain slice is continuously superfused with oxygenated aCSF at 32°C (arrows) via a peristaltic solution pump and in-line heater. The slice is illuminated by a cold light-source and the carbon-fiber recording electrode and bipolar stimulating electrodes positioned in a DA-rich region of the slice using two micromanipulators under microscopic visualization. A Millar voltammeter is triggered continually by a Master-8 pulse generator to apply a triangular voltage waveform every 100 ms via a headstage to the carbon-fiber microelectrode. The background non-Faradaic current detected at the surface of the electrode (versus a Ag/AgCl electrode located in aCSF near the slice) is amplified by the headstage and fed back to the voltammeter for background subtraction. Electrical stimulation is applied via a stimulus isolator controlled by the Master-8. Release of DA can also be evoked from DA somata, dendrites, and/or axons that express channelrhodopsin (e.g., 41). In that case, the Master-8 would be used to control the duration and frequency of light pulses from an LED light source or a laser and fiber optic cable. When the slice is stimulated to release DA, the Faradaic current increases at the characteristic DA oxidation and reduction peak potentials. The triangular voltage waveform and background subtracted voltammogram are both monitored on a digital storage oscilloscope and also sent to an analogue to digital (A-to-D) interface connected to a computer for data capture and off-line analysis.

concern for the usual nA–mA currents generated at carbon-fiber microelectrodes, eliminating the need for an auxiliary electrode. To monitor the time course of a DA release response, current detected at the DA oxidation peak potential can be followed using a sample-and-hold circuit (as a difference signal, this can be amplified as needed. Signal identification, however, requires evaluation of background-subtracted voltammograms of the released substance (Fig. 2c). This can be done off-line and is accomplished using a digital storage oscilloscope or data capture software with digital subtraction capabilities that permits identification of the DA signal (9, 10, 12). Labview-driven software also permits off-line subtraction of all scans. As already noted, however, the Millar Voltammeter contains circuitry that stores a background scan which is then subtracted from subsequent scans (i.e., analogue subtraction), so the Faradaic voltammogram corresponding to an increase in [DA]o can be monitored in real time.

252

J.C. Patel and M.E. Rice

3.2. Fabrication of Carbon-Fiber Microelectrodes

The earliest electrodes developed for the detection of DA release in brain tissue were made using carbon paste. These were superseded by carbon-fiber microelectrodes (16), which are now the electrode of choice in almost all in vitro or in vivo voltammetric DA release studies. A number of methods for the fabrication of carbon-fiber microelectrodes have been described (12, 16–22). In all of these, the fundamental design consideration is to get the highest signalto-noise ratio and lowest detection limit possible. The basic procedure involves inserting a carbon fiber, ~7 mm in diameter, in a glass capillary tube, then pulling the filled capillary on a conventional microelectrode puller to provide a seal and insulate the fiber with glass (Fig. 4). Every attempt is made to ensure that the seal is tight,

Fig. 4. Photographic summary of steps to make a cylindrical carbon-fiber microelectrode. (a) Loading: a glass capillary is placed in an angled test tube containing a few mL of acetone. A 7 mm carbon fiber is selected, cut, then loaded into the acetone-filled capillary. (b) Pulling: after the acetone has been removed and the loaded capillary allowed to air dry for several hours, the capillary is then placed in an electrode puller, with heat and magnet settings optimized to form a good seal between the glass and the carbon fiber, without burning the fiber. (c) Cutting: the pulled electrode is placed in an electrode holder in a micromanipulator oriented perpendicular to another manipulator with an electrode holder equipped with a scalpel blade. The optimal length of fiber extending beyond the glass seal is 30–70 mm. (d) Tip of a cylindrical carbon-fiber microelectrode after cutting; the junction at which the glass insulation ends us indicated by a white arrow ; the fiber extends ~40 mm beyond this junction (each small division is 10 mm). (e) Connecting: in this completed electrode, Woods metal was used to make electrical contact between the carbon fiber and the insulated lead wire that will be used to link the electrode to the headstage of a voltammeter. This figure is in color in the on-line version only.

15

Monitoring Dopamine Release

253

as small gaps or cracks can lead to background noise as solution is drawn into them by capillary action. The design of carbon-fiber microelectrodes used for brain slice recording must include an active surface that is completely contained within a £400 mm thick slice. Such surfaces can be prepared by beveling or cutting or by chemical or electrical etching of the carbon-fiber tip. The resulting working surface can either be a disk that is flush with surrounding insulation, a slightly extended fiber with a conical shape, or a cylinder that extends <70 mm beyond the insulation (5). Longer electrodes work well in vivo, but are not practical for use in slices. Methods for making the short-cylinder electrodes we use are given below. One final issue in making carbon-fiber microelectrodes is how to make electrical contact between a typically 5–10 mm carbon fiber and the electrode lead that will make contact with the instrumentation required for voltammetric recording. Poor contact with the fiber can be a significant source of noise. Materials used include electrically conducting silver paint or epoxy, Woods metal or conductive solutions like potassium acetate mixed with KCl (12, 16, 20, 22), each of which can link the fiber to a larger diameter wire that is then used for connection to the voltammeter headstage. The manufacture of carbon-fiber microelectrodes in the authors’ laboratory are based on the procedures described in refs. (16, 20). We make several electrodes in a given session although not all of the electrodes made will end up being suitable for experimental use. Electrodes are carefully stored in a box to protect them from mechanical or atmospheric damage at all stages after construction. 3.2.1. Loading

1. Loading a capillary tube with a carbon fiber is most easily accomplished when the tube is filled with acetone. The acetone serves two purposes; first it facilitates insertion of the carbon fiber into the capillary and second it removes any coatings or residue that may have formed on the carbon fiber surface. Using a Pasteur pipette, add a few mLs of acetone to a glass test tube £8 cm long. This should be held at a ~30° angle from horizontal, e.g., with modeling clay (Fig. 4a). 2. Place a single capillary in the test tube; it should extend ~2 cm beyond the end of the test tube (Fig. 4a). The acetone should immediately fill the glass barrel by capillary action. 3. With the capillary still in the test tube, use fine forceps with insulated tips to extract a single carbon fiber from the fiber bundle and gently insert it into the acetone-filled capillary about 1 cm at a time. A low power microscope or magnifying glass may be useful for this stage. 4. Remove the loaded capillary from the acetone, then touch one end to a tissue to remove the acetone from the capillary (without removing the fiber). As soon as the acetone is removed,

254

J.C. Patel and M.E. Rice

the carbon fiber will adhere to the inside wall of the capillary which will prevent the fiber from falling out. The protruding ends of the carbon fiber are then cut to about 1 cm beyond the ends of the glass capillary with fine scissors. Any remaining acetone should be allowed to evaporate for at least a few hours, but usually overnight, before pulling. 3.2.2. Pulling

1. Place the carbon-fiber filled capillary into an electrode puller (Fig. 4b). The combination of heat and magnet settings must be determined empirically for each puller and adjusted as the heating coil ages or when it is replaced. The ideal settings make a taper of 1–2 cm that seals the carbon fiber tightly. With a good seal, it is difficult to see where the glass ends and bare fiber begins. If the heat is too high, this can burn the fiber, which becomes wavy. If the heat is too low, the seal may not be good. Too little pull from the magnet may make the taper too short, whereas too strong a pull can result in a flare at the end of the glass where the seal should be. 2. When puller settings have been optimized, the capillary will be pulled into two halves with each tapered around the carbon fiber. The high tensile strength of the carbon fiber usually prevents it from breaking when the halves separate. Cut the fiber between the glass tapers to form two separate electrodes. Depending on the puller, the fiber may seal in only one half.

3.2.3. Cutting

1. The optimal length for slice recording is ~50 mm. Cutting the fiber is a precise and delicate process that should be done using a high-powered binocular microscope (100× to 200× total magnification). 2. Position two micromanipulators at right angles to one another to enable fine positioning of the carbon-fiber microelectrode and the cutting device (e.g., a mounted scalpel blade) (Fig. 4c). Both should be adjusted to advance a microelectrode holder with the electrode or cutting blade in a roughly horizontal plane to reach the visual field of the microscope. Attach the blade to one holder and the electrode to the other. 3. Visually inspect the glass seal, which should be tight and almost invisible (Fig. 4d); if not the electrode will likely be noisy. Then check that a sufficient length of the fiber protrudes back into the shaft of the capillary to enable electrical contact to be made. 4. Lower the electrode onto a glass slide so that the tip lightly rests on the surface of the slide. Using the edge of the mounted scalpel blade cut the carbon fiber to a length of 30–70 mm from the glass seal (Fig. 4d). In theory the second half of the pulled capillary can also be used but in practice this is rare and is usually because of a poor seal or that the fiber preferentially ends up in one half.

15 3.2.4. Connecting

Monitoring Dopamine Release

255

Electrical contact between the carbon fiber and the headstage of the voltammeter can be made using insulated, multi-stranded copper, or other flexible wire with ~1 mm total diameter that is secured to the fiber with a conductive sealant (Fig. 4e). 1. Cut the wire to a length of ~15 cm; this should be long enough to connect to the headstage, but short enough to minimize picking up electrical noise in the system which would be amplified by the headstage. Strip ~1 cm of insulation from one end of the wire and <0.5 cm from the other. 2. Solder a 1 mm gold pin to the shorter end for connection to the voltammeter headstage. This connection can be insulated with heat-shrink tubing if desired (Fig. 4e). 3. Dip the longer stripped end of the wire into electrically conductive silver paint or silver epoxy and push in as far as possible into the shank of the electrode to make contact with the carbon fiber. 4. Alternatively, low-melting point Woods metal can be used to make electrical contact. For this method, melting the metal in a beaker on a hot plate and drawing it into a plastic tube can make a Woods metal plug, 1 mm in diameter. Drop a plug that is a few millimeters in length into the open end of the electrode and then push in the stripped wire as in step 2. The metal plug is melted using a heat gun or hair dryer to connect the wire with the fiber (see Note 1).

3.3. Testing and Calibrating CarbonFiber Microelectrodes

Each carbon-fiber microelectrode must be screened for quality. This involves ensuring that the electrode has a suitable and stable background current profile, low electrical noise, and adequate sensitivity to detect low mM DA concentrations during calibration. We typically test and calibrate our electrodes in aCSF in the brain slicerecording chamber at the same temperature (~32°C) used during our experiment. With experience, one can determine very quickly whether an electrode will be viable for reliable recordings in brain tissue. It is best to test and calibrate electrodes in the aCSF solution used for brain slice recordings rather than in standard phosphate-buffered saline (PBS) because the sensitivity to DA can differ considerably between media. For example the absence of divalent Ca2+ and Mg2+ ions in PBS leads to a twofold higher peak DA oxidation current than that seen in aCSF for a given DA concentration (23). This presumably is because divalent ions associate with anionic functionalities (e.g., carboxylic acid) on the carbon surface thereby decreasing DA adsorption and thus oxidation current. As a consequence, calibration in PBS or other standard buffers that lack these divalent ions will lead to an underestimate of [DA]o in the brain microenvironment, which includes Ca2+ and Mg2+.

256

J.C. Patel and M.E. Rice

3.3.1. Setting Up Calibration in the Brain Slice Chamber

1. Prepare standard aCSF, continually bubbled with 95% O2/5% CO2. Use a peristaltic pump to flow the aCSF through a temperature controller into the brain slice recording chamber; chamber temperature should be set to ~32°C. 2. Anodize half of a 4–5 cm length of silver wire in 0.1 M KCl, using a 9 V battery to make the Ag/AgCl reference electrode. Attach the wire to be anodized to the (+) pole of the battery and another Ag (or Pt) wire to the (−) pole. A smooth, light gray AgCl coating is optimal. When this becomes black (Ag2O) and/or begins to flake off, it is time to remake. 3. For two-electrode recording with a Millar Voltammeter, make a short connection between the reference and auxiliary electrode pins on the headstage before connecting a reference or carbon fiber microelectrode to the headstage. 4. Secure the Ag/AgCl reference electrode in the corner of the chamber and connect it (using a small alligator clip attached to a flexible, insulated wire) to the appropriate input socket of the voltammeter headstage. Make sure that the electrode and clip do not touch any other metal surface. 5. Using a micromanipulator equipped with a microelectrode holder, lower a carbon-fiber electrode into the superfusing aCSF. Turn on the voltammeter and then connect the electrode to the working electrode socket of the voltammeter headstage to avoid an electrical surge that could damage the headstage or the electrode. 6. Switch on the scan trigger (or initiate control of scans from a digital timing circuit, e.g., Master-8) and monitor the voltage and current waveforms on a digital storage oscilloscope. Ensure that the starting potential and the reversal potential of the triangle voltage waveform are appropriate. Adjust the current gain so that the background current signal monitored in “Full” mode is just below the point at which the amplifiers become saturated, which can be seen when the current waveform is flattened at the top or bottom, even though the oscilloscope scale is appropriate. 7. Assess the shape of the background current. A carbon fiber in contact with an electrolyte behaves electrically as a resistor– capacitor (RC) network (24) and is further influenced by the electrolytic composition of the test solution because of surface functionalities already discussed. If the shape of the background current is rounded, much as one would expect for charging a simple RC circuit, the electrode will likely have little sensitivity for DA and should be discarded. The shape of the background current for a “good” electrode will have welldefined additional features, including a broad peak at around 0.3 V vs. Ag/Ag/Cl and some evidence of water oxidation

15

Monitoring Dopamine Release

257

near the switching potential of the input waveform (Fig. 2c). These features usually indicate a good surface for DA oxidation and predict good electrode sensitivity. 8. Assess the stability of the background current. A slow drift in the shape of the background current over a period of a few hours has a negligible effect, whereas large changes occurring over a few minutes usually indicates that the electrode is unreliable. Large changes in background current are often caused by a poor glass-carbon seal which allows electrolytes to creep up into the electrode shaft. 9. Assess electrical noise. Sufficient grounding should minimize the inherent noise of the apparatus and environment when the rig is set up and throughout an experiment. Electrical noise associated with an electrode should be minimal, as well. This can be assessed by switching to “Faradaic” mode, in which differences from a background scan can be readily seen and quantified in comparison to a built in 10 nA calibration pulse. If a calibration pulse is not available with the instrumentation used, this can be assessed from the current axis of voltammograms monitored on a PC with appropriate data acquisition software. Noisy electrodes are usually the result of poor electrical contact between the connecting wire and the carbon fiber or a poor glass-carbon seal. If the level of electrical noise is unacceptable (typically >1–2 nA) the electrode should be discarded. 3.3.2. Calibrating

1. Position the electrode at the superfusing inlet of the chamber so that the carbon-fiber tip is directly in the incoming flowstream. When the background-subtracted current is stable, switch the flow to aCSF with a known concentration of DA (e.g., 1 mM) in aCSF, remembering to refresh the background current immediately before the DA solution comes in contact with the electrode surface. It is important to match the rate of bubbling of the calibration solution with that of the background aCSF or background shifts from differences in solution pH (e.g., (12)) can distort the DA voltammogram. The appearance of a single oxidation peak (at ~+600 mV) and a single reduction peak (at ~−200 mV) can be monitored on the oscilloscope. The time that the DA is in contact with the electrode should be recorded using a stop-watch and limited to a set time between 30 and 60 s, after which the Faradaic signal should be electronically recorded for analysis and the solution changed back to control aCSF. 2. Several DA concentrations can be used to confirm the linearity of the current response over the range of [DA]o expected in a brain slice.

258

J.C. Patel and M.E. Rice

3. The detection limit can be defined as the concentration at which the Faradaic current is twice that of the background noise. Electrodes made according to these protocols have a detection limit of ~30–50 nM. 4. Although preexperiment calibration is necessary to assess electrode quality and sensitivity, calibration factors determined after the electrode has been in brain tissue are the most reliable for quantification of evoked [DA]o during an experiment. Typical postexperiment calibration factors are 5–20 nA/mM. 5. If the composition of aCSF will be changed in an experiment or if a new drug is to be used, the effect of these changes on electrode sensitivity to DA must be assessed. Whether a new drug is electroactive must also be determined. This is a threestep process. First, determine the calibration factor for 1 mM DA in aCSF alone. Second, switch the calibration solution back to aCSF alone until the background current is again stable, then switch to aCSF plus drug at the concentration to be used experimentally; be sure to “refresh” the scan immediately before drug-aCSF enters the recording chamber. Oxidation peaks if the drug is electroactive or non-Faradaic current changes (e.g., from electrode filming) can interfere with DA detection and may preclude the use of the drug. Third, to assess whether the presence of a drug alters electrode sensitivity to DA, refresh the background again in the continued presence of drug-aCSF and switch to drug-aCSF plus 1 mM DA. If the sensitivity of the electrode to DA is lost in the presence of the drug, this again precludes the use of the drug for FCV experiments. If the drug causes a consistent increase or decrease in the calibration factor from that in aCSF alone, then separate calibration factors to quantify evoked release in aCSF alone versus in drug can be used, although quantitation may be compromised. This should only be done if a similar drug that does not interfere with electrode sensitivity is not available. 3.4. Preparation of Brain Slices

Optimal conditions for preparing and maintaining viable brain slices have been developed over many years and are discussed in a comprehensive book edited by Dingledine (25). A few basic points relevant for midbrain and forebrain slices are summarized here. 1. Slices are made from deeply anesthetized animals after decapitation and cut while submerged in ice-cold aCSF or modified aCSF. For example, using an oxygenated HEPES-bicarbonatebuffered aCSF for slice preparation (26) and during the holding period (7, 8) improves slice quality by minimizing water gain (edema) (27). 2. Slice quality is best when slices are cut using a classic Vibratome or a Leica vibrating microtome (Leica Microsystems, Buffalo Grove, IL). Both instruments separate tissue into slices using a

15

Monitoring Dopamine Release

259

plane of turbulent solution that precedes a vibrating blade, thereby minimizing compression injury to the tissue. 3. Slice thickness should not exceed 400 mm, which is the maximum at which the core of the slice can still receive adequate oxygen and glucose. On the other hand, the minimal thickness of acute slices is typically considered to be 200 mm, which provides sufficient viable tissue between the cut surfaces, where there are damaged cells and severed processes. 4. Forebrain and midbrain slices are most commonly cut in the coronal plane (Fig. 1). However, horizontal, sagittal, or parasagittal slices can be cut in order to preserve a few millimeters of the DA pathway itself (28) or a non-DA input for stimulation purposes. For example horizontal slices from the small mouse brain can preserve DA connections between the midbrain and ventral striatum such that spontaneous [DA]o transients can be detected (29). 5. Once prepared, viable slices can be maintained in a holding chamber for 8–10 h at room temperature, although slice viability may not be as robust with older animals and/or in some experimental models (e.g., after DA system lesion or genetic manipulation) 3.4.1. Before Anesthetizing an Animal

1. Make sure all surgical equipment required (scalpel and razor blades, small scissors, iris scissors, small rongeurs, etc.) is available and clean. 2. aCSF (e.g., HEPES-bicarbonate buffered aCSF) is prepared and chilled on ice to ~2°C (allow 30 min for 200 mL in a beaker). 3. Clean a razor blade and the blade for the vibrating microtome with acetone, then ethanol to remove greasy lubricant manufacturers usually use to prevent rusting. This can be done by sequential immersion in each solvent. The solvents can be reused if capped to prevent evaporation, but should be changed at least bi-weekly. 4. Optimize the settings on the vibrating microtome for cutting the brain region of interest. These settings can vary among species. Parameters include the angle of the blade, the speed at which the blade advances, and amplitude at which the blade vibrates. A general rule is that slower advancement and higher vibration amplitude minimizes tissue damage, with the caveat that taking too long to slice can compromise slice viability. 5. Prepare a tissue holding chamber (e.g., a large plastic Petri dish) containing the desired aCSF at room temperature, with gentle bubbling of 95% O2/5% CO2 that is sufficient to maintain oxygenation and proper pH (7.4–7.6), but not so vigorous that the tissue moves around the container.

260

J.C. Patel and M.E. Rice

3.4.2. Dissecting

1. Anesthetize an animal with an appropriate anesthetic until pedal reflexes are absent (e.g., sodium pentobarbital, 50 mg/ Kg/i.p). 2. Using a scalpel, make an incision along the midline of the scalp from neck to snout of the anesthetized animal. Pull back the skin flaps and remove underlying connective tissue to expose the skull surface. As soon as the skull is exposed, use a Pasteur pipette to bathe it with a few milliliters of ice-cold oxygenated HEPES-buffered aCSF. 3. Insert the lower blade of sharp, pointed scissors beneath the bone at the back of the skull and make a short cut along the midline. Bathe exposed tissue with cold aCSF. Remove the bone on each side of the cut using fine rongeurs and repeat the procedure until all the bone posterior to Bregma is removed. Tips of scissors and rongeurs should be pointed upwards to prevent tissue damage. Indeed, care is more important than speed for this and all other steps, although efficiency will come with experience. 4. Cut and remove the dural membrane using fine iris scissors and fine forceps and continue removing bone until olfactory bulbs are revealed. 5. Gently invert the skull and then release the brain into a beaker of ice-cold oxygenated aCSF by carefully inserting a small spatula between the brain and the base of the skull to sever the cranial and optic nerves. 6. Allow the brain to chill 1–2 min to minimize on-going metabolism and the consequences of anoxia. Chilling procedure also makes the brain firmer for blocking and cutting.

3.4.3. Blocking and Slicing

1. To cut coronal slices, remove the chilled brain from the beaker of aCSF and place it ventral side-up on an inverted glass Petri dish (or other surface) covered with filter paper soaked with ice-cold aCSF. 2. Remove any remaining membranes that could snag the blade during slicing. Using a cleaned razor blade cut a block of tissue that is 3–7 mm in anterior-to-posterior length that incorporates the brain region of interest (e.g., striatum in the forebrain or substantia nigra in the midbrain). 3. Affix one cut surface of the tissue block to the specimen plate of the tissue slicer using cyanoacrylate glue (Krazy glue). For forebrain slices, glue the posterior surface so that slices are cut from anterior-to-posterior locations; orient the block so that the dorsal surface faces the blade for dorsal-to-ventral slicing. For midbrain slices, glue the cut anterior surface so that slices are cut from posterior-to-anterior locations; orient the midbrain block so that the ventral surface faces the blade for ventral-to-dorsal slicing. These procedures have been found

15

Monitoring Dopamine Release

261

empirically to produce the best slices for voltammetric recording. 4. Fit the specimen plate into the buffer well of the tissue slicer as quickly as possible after gluing, and fill the buffer well with oxygenated ice-cold aCSF. 5. Attach a clean slicing blade to the vibrating microtome. 6. Position the starting point of the blade to allow ~500 mm of tissue to be cut from the block to leave a flat, horizontal plane for subsequent slicing; discard this section. 7. Begin slicing of tissue sections of the desired thickness (200– 400 mm). Using a small spatula or a glass pipette, carefully transfer each slice as it is made to the tissue holding chamber. The number of slices prepared also depends on the region of interest and the species examined. For example, more slices can be made from striatum than from the smaller midbrain DA cell body regions, and more from either region using a guinea pig than the smaller mouse brain. 8. Allow at least 1 h for recovery in the holding chamber at room temperature before experimentation. 3.5. Stimulating and Recording DA Release in Brain Slices

In the striatum, in vivo, rapid, spontaneous [DA]o transients can be detected using FCV, which reflect axonal DA release in response to presumed bursting of DA neurons in the midbrain (30). Most in vitro brain slices preparations, however, contain either DA axons (forebrain slices) or DA neurons (midbrain slices), so such spontaneous transients are not seen in striatal slices. Moreover, DA neurons in vitro display steady pacemaker activity rather than bursting behavior (31), so that somatodendritic [DA]o transients are also not seen in slices. However, axonal and somatodendritic DA release can be readily evoked using local electrical stimulation (10, 32–36) both of which are sensitive to Na+ channel blockade, indicating action-potential dependence, and are Ca2+ dependent. An insulated bipolar stimulating electrode is used to evoke release, which can be either concentric or parallel in style. Stimulating electrodes are typically made from tungsten, platinum, or platinum–iridium and can be constructed locally or purchased from a commercial supplier. For example, an effective twisted wire electrode can be constructed from insulated narrow gauge wire (e.g., 75 mm diameter wire only), with tip separation of ~100 mm. Methods in this section will focus on local electrical stimulation, but the basic procedures for voltammetric recording in slices can be used with other stimulation methods, as well. For example, superfusion or local pressure ejection of depolarizing agents, like high K+ or veratridine, can also be use to elicit DA release (21, 26, 37), with the caveat that these can cause large background shifts from changes in extracellular Ca2+ and pH. The effect of other releasing agents like amphetamine can also be examined using FCV (38–40). Most recently, optogenetic techniques that permit selective

262

J.C. Patel and M.E. Rice

activation of neuronal populations have been introduced and applied to study DA transmission. This methodology involves transfecting DA neurons with photo-sensitive channelrhodopsin using a viral vector injected into the SNc or VTA of DAT-Cre or TH-Cre transgenic mice. Once the virus has fully penetrated and has been transported throughout the DA neuron, including axon terminals (usually after a few weeks), slices are prepared as usual and a blue laser or LED is used to elicit DA release selectively (41). 3.5.1. Brain Slice Recording

For voltammetric recording of DA release using carbon-fiber microelectrodes in brain slices, it is preferable to use a submersion chamber in which slices (and electrode tips) can be submerged in aCSF at all times, rather than an interface chamber in which the slice surface is exposed to a humidified atmosphere of 95% O2/5% CO2 into which the electrode must be drawn after recording. Keeping the carbon-fiber microelectrode in solution at all times when not in tissue helps maintain a constant electrode surface state and thereby optimal reliability of electrochemical measurements. Background currents are also kept as constant as possible by continuous triggering of the FCV voltage waveform from the time the electrode is first positioned in the chamber for preexperiment calibration until the end of the experimental day. A few basic points about brain slice recording in a submersion chamber are summarized here. 1. Slices are superfused at a rapid but constant rate (typically >1–2 mL/min) with freshly prepared oxygenated bicarbonatebuffered aCSF. 2. To maintain slice viability, aCSF used for in vitro brain slice recording differs from CSF in vivo in that it is hyperoxic, when equilibrated with 95% O2/5% CO2, and hyperglycemic, with 10 mM glucose. Both conditions are necessary to provide sufficient O2 and glucose to maintain the viability of cells below the slice surface, as superficial cells consume these metabolic substrates. To preserve O2 and CO2 saturation of aCSF en route to a slice, the inlet tubing should be made of gas-impermeable material, although this is not always done. 3. Tubing should also be narrow and kept as short as practical to ensure rapid delivery of oxygenated aCSF. This also facilitates efficient solution changes for drug studies as well as for electrode calibration. 4. The slice recording environment is slightly hypothermic, typically ~32°C, which helps preserve viability for several hours of recording, and also decreases temperature-dependent DA uptake via the DA transporter (DAT), enhancing [DA]o during stimulation (6). 5. The aCSF used for recording usually has slightly elevated concentrations of K+ and/or Ca2+ compared to those in CSF in vivo to enhance slice excitability and Ca2+-dependent DA release.

15 3.5.2. Brain Slice Orientation and Electrode Placement

Monitoring Dopamine Release

263

1. Once the brain slice recording chamber is set up with aCSF (at ~32°C) flowing through at a rate of at least 1.2 mL/min and the Ag/AgCl reference electrode secured in place, as described in Subheading 3.3, carefully transfer a suitable slice from the holding chamber to the recording chamber using a spoon-like spatula or a glass pipette with an open tip greater than the dimensions of the slice. Keep the slice immersed in aCSF and flat at all times during transfer to prevent mechanical trauma. 2. Orient the slice in the recording chamber to allow optimal placement of the stimulating and recording electrodes (more below). Secure the slice by placing 3 or 4 small lengths (~2–4 mm) of bare silver wire around the edges of the slice to hold it down without blocking the flow of aCSF over the slice surface or interfering with electrode placement. 3. Position a fiber optic (cold) light source below the slice to allow transillumination of the tissue for viewing with a dissection microscope. Under these illumination conditions gray matter appears translucent, while the white matter containing myelinated fiber bundles appears dark. 4. Connect the stimulating electrode to a stimulus isolator controlled by a Master-8 or other pulse-timing device. The use of a stimulus isolator avoids interference with the ground of the voltammeter. 5. Allow 20–30 min for equilibration with the recording medium before stimulating the tissue. The voltammetric electrode can be positioned in the tissue at any time during this equilibration period to facilitate background current stabilization; scan trigger is on. Insert the electrode tip 50–100 mm below the slice surface under the control of a micromanipulator. The stimulating electrode can be positioned using a micromanipulator at this time as well, with the electrode tip(s) just resting on the surface of the slice. An appropriate rodent stereotaxic atlas can be used for guidance in optimal electrode orientation, although this must be confirmed empirically. 6. To monitor evoked axonal DA release in the striatal complex (Fig. 1b), the carbon-fiber microelectrode is typically positioned ~100 mm dorsolateral to the stimulating electrode. Both recording and stimulating electrodes should be placed so as to avoid the non-dopaminergic myelinated fiber bundles that run through the striatum. In the dorsolateral (motor) striatum and nucleus accumbens shell and core, which can all be contained in a coronal plane of section (see Fig. 1b), reproducible DA release can be evoked in the same site for several hours at 2–5 min intervals for single-pulse stimulation or 5–10 min for pulse-train stimulation. Single-pulse stimulation is ~1 mM in the guinea-pig dorsal striatum but is lower in the NAc core and even lower in the NAc shell.

264

J.C. Patel and M.E. Rice

7. To monitor somatodendritic DA release in the SNc and VTA, optimal release is usually seen when the recording electrode is positioned between the poles of a non-concentric bipolar stimulated electrode. The location of these cell body regions at similar midbrain levels make it possible to record from both regions in a single coronal slice (see Fig. 1b). Moreover, the lateral projecting dendrites of SNc DA neurons lie in this plane. In the SNc, both poles of the stimulating electrode should be placed within this band of DA somata, with the carbon-fiber tip equidistant between them. Note that this band is not pigmented in rodents (as it is in humans), so that a brain atlas can be very useful for proper electrode placement. Detection of DA release provides confirmation of location. In both SNc and VTA multiple-pulse stimulation is usually required to detect DA release, which is typically much smaller (<0.5 mM) than that evoked in dorsal striatum. 3.5.3. Stimulating and Recording Axonal DA Release (Forebrain Slice)

1. After equilibration in the recording chamber, check the shape and stability of the background current. The shape should be similar to that seen in calibration. Stability is indicated by little drift in background-subtracted scans monitored on a digital oscilloscope or data acquisition program over a few seconds, i.e., the usual recording time for DA release measurements. 2. Set stimulus pulse duration on the timing system (e.g., Master-8); this is often 0.1 ms, but should ideally be £1 ms to remain shorter than the duration of action potentials initiated by each pulse. The same pulse duration should be used throughout an experimental series, and ideally used for both single-pulse and pulse-train stimulation in a given experiment. 3. The timing of stimulus pulse should be synchronized with the on-going FCV scan trigger, but adjusted to avoid “collision” of a stimulus pulse with an FCV scan. This can be achieved by delaying the stimulus trigger by 7–8 ms after initiation of a scan. This brief delay also can allow a stimulus artifact to be captured with each voltammogram to ensure that the stimulus was turned on, etc. This works perfectly for single-pulse or 10-Hz stimulation when FCV sampling interval is 100 ms, but may need adjustment for other pulse-train frequencies. 4. Set the stimulus isolator to apply current pulses. Although either current or voltage can be used, current pulses provide more consistent stimulation than voltage pulses because of possible voltage drop at the electrode surface from tissue buildup after exposure to slices over many days of experimentation. 5. Using the data acquisition software described (see Subheading 3.3), set parameters to capture the input voltage waveform and output current for each voltammetric scan. For single-pulse stimulation or pulse trains of up to 30 pulses, set the timing

15

Monitoring Dopamine Release

265

circuit to trigger data acquisition to provide ~2 s of baseline records before the stimulus pulse or train is triggered. Allow a ~9 s time window for data acquisition; with a scan trigger rate of 10 scans/s, this records 90 voltammograms. Excess baseline or recovery data can be omitted when making figures from release records, as appropriate. 6. Using single-pulse stimulation, determine the lowest stimulus intensity (amplitude) that gives the highest peak [DA]o. Determination of this “perimaximal” stimulus intensity is best done in dorsolateral striatum, which has the largest single-pulse evoked peak [DA]o of any dopaminergic region. Be sure to “refresh” the background immediately before stimulation and turn on the 10 nA calibration pulse. Start with a low current amplitude, e.g., 0.3 mA. With most stimulating electrodes, 0.3 mA will be subthreshold for initiating action potentialdependent DA release. Increment this sequentially using 0.05– 0.1 mA steps. As long as the intensity is subthreshold (i.e., no DA release is evoked), another measurement can be taken immediately. Once DA release is seen, however, wait 5 min before examining the next current increment. Usually after 2–5 more increments, no further increase in peak [DA]o is seen. Lower the current setting to the first setting that gave this maximal [DA]o. The rationale is that supramaximal stimulation can mask regulatory processes that might decrease DA release, e.g., from DA D2 autoreceptors that activate a hyperpolarizing K+ channel. Moreover, larger currents can cause electrolytic damage to the tissue (6) (see Note 2). 7. Single-pulse stimulation can be sampled at different sites of the slice or can be sampled at regular intervals at the same site. Peak [DA]o evoked by the first stimulus pulse in a slice is usually higher than subsequent pulses. However, this is usually constant (±10%) after 2–3 stimulations repeated at a constant interval of 4 or 5 min. Shorter interstimulus intervals may take slightly longer to become constant. However, this characteristic can be used deliberately to compare DA release “sustainability” among groups (42). Regardless of the stimulus-interval, release in some slices or recording sites can result in a progressive decrement in peak [DA]o that never becomes constant. Conducting experiments in these sites should be avoided. Single-pulse stimulation evokes DA release that is independent of regulation by transmitters released by the pulse, including glutamate and GABA, as well as DA itself (10, 32, 33, 43, 44). This is because the DA release event has already occurred by the time receptor-dependent regulation might have influenced it. One caveat is that in the striatal complex single-pulse DA release is influenced by ACh release from cholinergic interneurons (29, 45). Single-pulse stimulation is well-suited for

266

J.C. Patel and M.E. Rice

comparison of absolute DA release levels and/or DA uptake between groups of transgenic mice (e.g., (40, 42, 46)), determination of whether a drug has a direct effect on axonal DA release (e.g., nicotine, (29, 45), or to compare D2 autoreceptor sensitivity among groups (40, 46). For example, to assess D2 autoreceptor sensitivity, single-pulse stimulation is repeated until 5 records with peak evoked [DA]o that differ by £10% have been obtained, then the superfusing aCSF switched to aCSF plus a low concentration of a selective D2 agonist, e.g., quinpirole. Once a maximal effect on peak [DA]o is seen (~30– 40 min), a slightly higher concentration of the drug is applied and so on until responses in several concentrations are obtained. The efficacy of the drug (i.e., concentration of quinpirole at which a half maximal effect is seen can then be extracted from the sigmoidal concentration–response curve (40, 46). 8. Pulse-train stimulation. The number of pulses in the train and the frequency depend on the questions asked. With shorter pulse trains (4–5 pulses), a 5 min interval between stimulation is usually sufficient to maintain stable release after the first 2–3 stimulations. With longer trains (10–50 pulses), a 10 min interval is required. The simplest use of pulse-train stimulation is to assess the frequency dependence of DA release using constant pulse number and varying frequency in a range from 5 to 100 Hz (32, 33, 45, 46). Pulse-train stimulation can be used to assess how endogenous transmitters, including DA, glutamate, and GABA, influence DA released by subsequent pulses in a train; this contrasts with single-pulse evoked [DA]o which is independent of regulation by concurrently released transmitters. For this analysis, train stimulation with constant pulse number and frequency (e.g., 30 pulses at 10 Hz) is repeated until three records with peak evoked [DA]o that differ by £10% have been obtained, then the superfusing aCSF switched to aCSF plus a selective antagonist, e.g., for glutamatergic AMPA receptors (28, 47), and the effect on peak [DA]o assessed after a maximal effect is seen (30–60 min) (see Note 3). 9. Regional dependence of single-pulse and pulse-train DA release characteristics. The amplitude of single-pulse evoked [DA]o varies among striatal subregions with highest peak single-pulse [DA]o in dorsal striatum, intermediate in NAc core, and lowest in NAc shell (Fig. 5a–c). The shape of the release response also differs among these regions. In dorsal striatum, peak [DA]o is seen within 2–3 stimulus pulses, then declines during continued stimulation because of D2 DA autoreceptor activation and DAT-dependent uptake (Fig. 5a). By contrast, in NAc shell, a progressive increase in [DA]o is usually seen throughout a pulse train (Fig. 5c), so that although peak single-pulse [DA]o is much lower in NAc shell than in dorsal striatum, peak pulse-train

15

b

Dorsal striatum

0.6

c

NAc core

0.5

[DA]o (µM)

1.4 1.2 1.0 0.8 0.6 0.4 0.2 0

[DA]o (µM)

[DA]o (µM)

a

Monitoring Dopamine Release

0.4 0.3 0.2 0

[DA]o (µM)

0.6

1.6 1.2 0.8

0 10 Hz, 3 s 1P

10 Hz, 3 s 1P

e

SNc

0.5

0.6

[DA]o (µM)

d

0.4 0.3 0.2

NAc shell

2.0

0.4

0.1 10 Hz, 3 s 1P

2.4

267

VTA

0.5 0.4 0.3 0.2 0.1

0.1

0

0 10 Hz, 3 s

10 Hz, 3 s

Fig. 5. Comparison of DA release records obtained using FCV in different regions within forebrain and midbrain slices. (a–c) Representative [DA]o versus time records showing axonal DA release evoked by single-pulse stimulation (1P; gray traces) or pulse-train stimulation (30 pulses at 10 Hz; black traces) in the mouse dorsal striatum (CPu) (a), NAc core (b), and NAc shell (c) in a forebrain slice. (d–e) Representative [DA]o versus time records showing somatodendritic DA release evoked by pulse-train stimulation (30 pulses at 10 Hz) in the SNc and VTA in a guinea-pig midbrain slice.

[DA]o can be similar in these regions. The response in NAc core is intermediate between dorsal striatum and NAc shell (Fig. 5b), reflecting mixed DA input from SNc and VTA (48). 10. FCV can also be used to examine DA release in other axonterminal regions, like the olfactory tubercle (33), amygdala (49, 50), prefrontal cortex (51), and subthalamic nucleus (52). The small size of some of these structures as well as sparse DA innervation and significant NE input can present challenges, however. Confirmation that the monitored substance is DA rather than NE contributions must be made using pharmacological and anatomical methods to complement voltammetric data (49). 3.5.4. Stimulating and Recording Somatodendritic DA Release (Midbrain Slice)

Initial steps for stimulating and recording somatodendritic DA release in the SNc and VTA are identical to steps 1–5 for assessing axonal DA release in the previous section. Step 6, determining perimaximal stimulation intensity, is also important for assessing DA release from somata and dendrites. However, the typically small amplitude of single-pulse evoked [DA]o in these regions (~100 nM) make this tricky to assess. Consequently, stimulus parameters determined in dorsal striatum with the electrode orientation for midbrain recording can be used as a starting point. It should be noted that guinea pigs are the species of choice for

268

J.C. Patel and M.E. Rice

studies of electrically evoked somatodendritic DA release in the SNc because FCV measurements detect only DA, whereas stimulation of the SNc in rats and mice evokes concurrent release of 5-HT (26, 53–55). In VTA, however, DA is the only electroactive substance detected in the VTA of rats and mice (34, 55) as well as guinea pigs (35, 54). On the other hand, DA release in SNc is entirely somatodendritic, based on anatomical studies showing no synaptic DA input, whereas somatodendritic DA release in the VTA is mixed with synaptic DA release from its own axon collaterals as well as synaptic input from SNc (36). 1. Single-pulse stimulation. Like axonal DA release in the striatal complex, single-pulse evoked somatodendritic DA release in the midbrain is free from regulation by concurrently released transmitters (44). However, as already noted, midbrain DA release levels are often much lower than in striatum. To improve detection of single-pulse-evoked DA release in midbrain, [DA]o can be amplified by using a stimulus pulse duration of 1 ms, blocking D2 DA autoreceptors and inhibiting DA uptake (36, 44), with the caveat that this may also alter DA neuron excitability. 2. Pulse-train stimulation. As a consequence of the low levels of evoked [DA]o in SNc and VTA without pharmacological manipulation, pulse-train stimulation is most commonly used for studies of somatodendritic DA release (34–36, 54–57). Higher peak [DA]o is seen with longer trains (e.g., 30 pulses), so these are often used. The concentration–time profiles of pulse-train evoked [DA]o differs again from axonal release in forebrain structures (Fig. 5d, e). Indeed, [DA]o typically continues to rise after the stimulation train ends, consistent with limited DA uptake in these regions compared to striatum (57). Perhaps as a consequence of lower reuptake, somatodendritic DA release is not as robust as axonal release in striatum, with a progressive decrease in evoked [DA]o seen with repetitive stimulation. However, DA release regulation in the SNc can still be assessed by recording evoked [DA]o in 2–3 sites, then comparing these with a drug response in paired sites in the opposite hemisphere. Although this paradigm lacks the advantages of same-site controls, including requiring larger sample sizes because of greater variability, valuable information can still be obtained (8, 44, 56). Release in the VTA can be elicited repetitively in the same site (34), possibly reflecting the mix of axonal and somatodendritic release in this region, although paired measurements can also be used effectively in VTA. As in striatal regions, the use of selective receptor antagonists can be used to indicate the influence of concurrently release glutamate and GABA on somatodendritic DA release in both SNc and VTA (8, 56) (see Note 4).

15 3.5.5. Data Analysis

Monitoring Dopamine Release

269

1. The simplest form to data analysis is to express data as absolute levels of peak [DA]o obtained by converting the current detected into a DA concentration using the post-experimental calibration factor obtained for a given electrode. This is particularly useful for comparing release levels between brain regions in the same species, release in the same brain region between species, or release in the same brain region between a transgenic and non-transgenic mouse. 2. To examine the effect of a drug on evoked DA release, data are often expressed as a percentage of the control response, where control responses are set as 100%. This analysis is also useful to compare if a drug alters the time course of the response or if the profile differs between brain regions. 3. Data can also be expressed as a ratio of peak [DA]o evoked by a pulse-train stimulation relative to that evoked by a singlepulse stimulation when they are evoked in the same recording site. This analysis can provide an indication of the sensitivity of release to tonic versus phasic stimulation and can vary throughout the striatal complex (e.g., (10, 33)) or in the presence of drugs including D2 autoreceptor antagonists (e.g., (10, 33)) or drugs that act at striatal nicotinic receptors (45, 46). 4. Evoked [DA]o data can be used to evaluate the kinetics of DA release and uptake in axon terminal and cell body regions. A number of studies determine Michaelis-Menten uptake parameters using appropriate curve-fitting procedures (42, 59). In this analysis, evoked DA release (using electrical stimulation) is assumed to provide a homogeneous increase in extracellular concentration, such that diffusion plays no role in DA clearance. Typically, the Michaelis-Menten constant, Km, which indicates the [DA]o at which uptake is half-maximal, is set from literature values with experimental assessment of the maximal rate, Vmax, and the increase in [DA]o per stimulus pulse. This method can be used to examine regional or species variation in DA dynamics, to examine differences in transgenic versus nontransgenic mice or to characterize the effect of drugs acting at the DAT, like cocaine or amphetamine.

4. Notes 1. Individual users may chose to use a fresh electrode for each experimental day because of loss of sensitivity and possible increase in noise. However, we tend to re-use ours if they are still in good condition and the response kinetics of the surface has not deteriorated. Given the initial loss of sensitivity that can occur upon initial electrode exposure to brain tissue, the calibration factor of re-used electrodes can be relatively stable.

270

J.C. Patel and M.E. Rice

After use in tissue, the surface can be washed; further washing can take place when water is run through the chamber to remove aCSF after an experiment. Optimal methods are user dependent and should be determined empirically. 2. Perimaximal stimulation intensity should be assessed initially for every slice tested, because this can vary with the distance between stimulating and working electrode, as well as recording electrode depth, etc. Once an experimenter is proficient and consistent in electrode placement, perimaximal intensity will be consistent, as well. Nevertheless, this should still be assessed regularly, because the stimulus electrode surface and pole spacing can change over time, influencing current density in the surrounding tissue. Perimaximal stimulus intensity should also be assessed whenever a new stimulating electrode is used, a new slice thickness is used or when a slice from a different species is used. Another important test when setting up a new voltammetry system is to confirm that the stimulus used produces action-potential dependent DA release using 0.1–1 mM tetrodotoxin (TTX) (6, 36). Large gauge stimulating electrodes (³150 mm diameter) can produce TTX-insensitive release (35), which is not ideal. If the distance between the stimulating and recording electrode is too large, current levels ³1 mA may be required to elicit DA release, which as mentioned above can lesion to the tissue and result in a progressive decrease in peak evoked [DA]o (35). If the distance is too short, however, the stimulation voltammogram may be dominated by stimulation artifact (i.e., large background current shifts that do not correspond to DA oxidation or reduction peaks). Background-dominated shifts can also be seen when the carbon-fiber microelectrode tip is too close to one pole of the stimulating electrode. 3. The basic procedure for assessing the effects of receptor agonists or antagonists can be used with either single- or multiplepulse stimulation to address the effect of any drug or other agent on DA release. Ideally, the effects observed should reverse upon drug washout for ~30 min, unless a drug is hydrophobic or is known to have an irreversible action. 4. One final point about voltammetric recordings is that background shifts during stimulation can also arise from nonelectroactive interferents, especially Ca2+ and pH (H+), which can change dynamically with depolarizing stimuli (see ref. (5), for review). The potentials of these background shifts can readily be distinguished from the DA oxidation peak at ~+0.6 V vs. Ag/AgCl) (12, 58); however, this can interfere with quantitation in regions with low evoked [DA]o like the amygdala (58) as well as SNc and VTA. If necessary a method that uses principal component regression coupled with appropriate calibration

15

Monitoring Dopamine Release

271

data can be used to resolve substances that give overlapping voltammetric waveforms, including DA and pH (15).

Acknowledgments We are grateful for support from NIH/NINDS grant NS036362 and the Attilio and Olympia Ricciardi Research Fund (M.E.R). We also appreciate critical reading of the manuscript by Melissa A. Stouffer and Harry Xenias. References 1. Dahlström A, Fuxe K (1964) Evidence of the existence of monoamine-containing neurons in the central nervous system. I: demonstration of monoamines in the cell bodies of brain stem neurons. Acta Physiol Scand 62:1–55 2. Carlsson A (2002) Treatment of Parkinson’s with L-DOPA. The early discovery phase, and a comment on current problems. J Neural Transm 109:777–787 3. Evans AH, Lees AJ (2004) Dopamine dysregulation syndrome in Parkinson’s disease. Curr Opin Neurol 17:393–398 4. Cannon CM, Bseikri MR (2004) Is dopamine required for natural reward? Physiol Behav 81:741–748 5. Patel JC, Rice ME (2006) Dopamine in brain slices. In: Grimes CA, Dickey EC, Pishko MV (eds) Encyclopedia of sensors, vol 6. American Scientific Publishers, Stevenson Ranch, California, USA, pp 313–334 6. Bull DR, Palij P, Sheehan MJ, Millar J, Stamford JA, Kruk ZL, Humphrey PP (1990) Application of fast cyclic voltammetry to measurement of electrically evoked dopamine overflow from brain slices in vitro. J Neurosci Methods 32:37–44 7. Chen BT, Avshalumov MV, Rice ME (2001) H2O2 is a novel, endogenous modulator of synaptic dopamine release. J Neurophysiol 85:2468–2476 8. Patel JC, Witkovsky P, Avshalumov MV, Rice ME (2009) Mobilization of calcium from intracellular stores facilitates somatodendritic dopamine release. J Neurosci 29:6568–6579 9. Millar J, Stamford JA, Kruk ZL, Wightman RM (1985) Electrochemical, pharmacological and electrophysiological evidence of rapid dopamine release and removal in the rat caudate nucleus following electrical stimulation of the median forebrain bundle. Eur J Pharmacol 109:341–348

10. Patel J, Trout SJ, Kruk ZL (1992) Regional differences in evoked dopamine efflux in brain slices of rat anterior and posterior caudate putamen. Naunyn Schmiedebergs Arch Pharmacol 346:267–276 11. Armstrong-James M, Millar J, Kruk ZL (1980) Quantification of noradrenaline iontophoresis. Nature 288:181–183 12. Rice ME, Nicholson C (1989) Measurement of nanomolar dopamine diffusion using low-noise perfluorinated ionomer coated carbon fiber microelectrodes and high-speed cyclic voltammetry. Anal Chem 61:1805–1810 13. Adams RN (1969) Electrochemistry at solid electrodes. Marcel Dekker, Inc., New York, p 124 14. Millar J, Barnett T (1988) Basic instrumentation for fast cyclic voltammetry. J Neurosci Methods 25:91–95 15. Heien ML, Johnson MA, Wightman RM (2004) Resolving neurotransmitters detected by fast-scan cyclic voltammetry. Anal Chem 76:5697–5704 16. Armstrong-James M, Millar J (1979) Carbon fibre microelectrodes. J Neurosci Methods 1:279–287 17. Kawagoe KT, Zimmerman JB, Wightman RM (1993) Principles of voltammetry and microelectrode surface states. J Neurosci Methods 49:225–240 18. Cahill PS, Walker QD, Finnegan JM, Mickelson GE, Travis ER, Wightman RM (1996) Microelectrodes for the measurement of catecholamines in biological systems. Anal Chem 68:3180–3186 19. Koh DS, Hille B (1999) Rapid fabrication of plastic-insulated carbon-fiber electrodes for micro-amperometry. J Neurosci Methods 88:83–91 20. Millar J, Pelling CW (2001) Improved methods for construction of carbon fibre electrodes

272

21.

22.

23.

24.

25. 26.

27.

28.

29.

30.

31.

32.

J.C. Patel and M.E. Rice for extracellular spike recording. J Neurosci Methods 110:1–8 Gerhardt GA, Hoffman AF (2001) Effects of recording media composition on the responses of Nafion-coated carbon fiber microelectrodes measured using high-speed chronoamperometry. J Neurosci Methods 109:13–21 Clark JJ, Sandberg SG, Wanat MJ, Gan JO, Horne EA, Hart AS, Akers CA, Parker JG, Willuhn I, Martinez V, Evans SB, Stella N, Phillips PE (2010) Chronic microsensors for longitudinal, subsecond dopamine detection in behaving animals. Nat Methods 7:126–129 Kume-Kick J, Rice ME (1998) Dependence of dopamine calibration factors on media Ca2+ and Mg2+ at carbon-fiber microelectrodes used with fast-scan cyclic voltammetry. J Neurosci Methods 84:55–62 Fox K, Armstrong-James M, Millar J (1980) The electrical characteristics of carbon fibre microelectrodes. J Neurosci Methods 3:37–48 Dingledine R (1984) Brain slices. Plenum Publishing Corporation, New York Rice ME, Richards CD, Nedergaard S, Hounsgaard J, Nicholson C, Greenfield SA (1994) Direct monitoring of dopamine and 5-HT release from substantia nigra and ventral tegmental area in vitro. Exp Brain Res 100: 395–406 MacGregor DG, Chesler M, Rice ME (2001) HEPES prevents edema in rat brain slices. Neurosci Lett 303:141–144 Avshalumov MV, Patel JC, Rice ME (2008) AMPA receptor-dependent H2O2 generation in striatal medium spiny neurons, but not dopamine axons: one source of a retrograde signal that can inhibit dopamine release. J Neurophysiol 100:1590–1601 Zhou FM, Liang Y, Dani JA (2001) Endogenous nicotinic cholinergic activity regulates dopamine release in the striatum. Nat Neurosci 4:1224–1229 Sombers LA, Beyene M, Carelli RM, Wightman RM (2009) Synaptic overflow of dopamine in the nucleus accumbens arises from neuronal activity in the ventral tegmental area. J Neurosci 29:1735–1742 Grace AA, Onn S-P (1989) Morphology and electrophysiological properties of immunocytochemically identified rat dopamine neurons recorded in vitro. J Neurosci 9:3463–3481 Limberger N, Trout SJ, Kruk ZL, Starke K (1991) “Real time” measurement of endogenous dopamine release during short trains of pulses in slices of rat neostriatum and nucleus accumbens: role of autoinhibition. Naunyn Schmiedebergs Arch Pharmacol 344:623–629

33. Trout SJ, Kruk ZL (1992) Differences in evoked dopamine efflux in rat caudate putamen, nucleus accumbens and tuberculum olfactorium in the absence of uptake inhibition: influence of autoreceptors. Br J Pharmacol 106:452–458 34. Iravani MM, Muscat R, Kruk ZL (1996) Comparison of somatodendritic and axon terminal dopamine release in the ventral tegmental area and the nucleus accumbens. Neuroscience 70:1025–1037 35. Rice ME, Cragg SJ, Greenfield SA (1997) Characteristics of electrically evoked somatodendritic dopamine release in substantia nigra and ventral tegmental area in vitro. J Neurophysiol 77:853–862 36. Chen BT, Rice ME (2001) Novel Ca2+ dependence and time course of somatodendritic dopamine release: substantia nigra vs. striatum. J Neurosci 21:7841–7847 37. Elverfors A, Jonason J, Jonason G, Nissbrandt H (1997) Effects of drugs interfering with sodium channels and calcium channels on the release of endogenous dopamine from superfused substantia nigra slices. Synapse 26:359–369 38. Iravani MM, Kruk ZL (1995) Effects of amphetamine on carrier-mediated and electrically stimulated dopamine release in slices of rat caudate putamen and nucleus accumbens. J Neurochem 64:1161–1168 39. Schmitz Y, Lee CJ, Schmauss C, Gonon F, Sulzer D (2001) Amphetamine distorts stimulation-dependent dopamine overflow: effects on D2 autoreceptors, transporters, and synaptic vesicle stores. J Neurosci 21: 5916–5924 40. Patel J, Mooslehner KA, Chan PM, Emson PM, Stamford JA (2003) Presynaptic control of striatal dopamine neurotransmission in adult vesicular monoamine transporter 2 (VMAT2) mutant mice. J Neurochem 85: 898–910 41. Tecuapetla F, Patel JC, Xenias H, English D, Tadrosi I, Shah F, Berlin J, Deisseroth K, Rice ME, Tepper JM, Koos T (2010) Glutamatergic signaling by mesolimbic dopamine neurons in the nucleus accumbens. J Neurosci 30: 7105–7110 42. Li X, Patel JC, Wang J, Avshalumov MV, Nicholson C, Buxbaum JD, Elder GA, Rice ME, Yue Z (2010) Enhanced striatal dopamine transmission and motor performance with LRRK2 overexpression in mice is eliminated by familial Parkinson’s disease mutation G2019S. J Neurosci 30:1788–1797 43. Phillips PEM, Hancock PJ, Stamford JA (2002) Time window of autoreceptor-mediated

15 inhibition of limbic and striatal dopamine release. Synapse 44:15–22 44. Chen BT, Moran KA, Avshalumov MV, Rice ME (2006) Limited regulation of somatodendritic dopamine release by voltage-sensitive Ca channels contrasted with strong regulation of axonal dopamine release. J Neurochem 96: 645–655 45. Rice ME, Cragg SJ (2004) Nicotine amplifies reward-related dopamine signals in striatum. Nat Neurosci 7:583–584 46. Bao L, Patel JC, Walker RH, Shashidharan P, Rice ME (2010) Dysregulation of striatal dopamine release in a mouse model of dystonia. J Neurochem 114:1781–1791 47. Avshalumov MV, Chen BT, Marshall SP, Peña DM, Rice ME (2003) Glutamate-dependent inhibition of dopamine release in striatum is mediated by a new diffusible messenger, H2O2. J Neurosci 23:2744–2750 48. Voorn P, Vanderschuren LJMJ, Groenewegen HJ, Robbins TR, Pannartz CMA (2004) Putting the spin on the dorsal-ventral divide of the striatum. Trends Neurosci 27: 468–474 49. Bull DR, Bakhtiar R, Sheehan MJ (1991) Characterization of dopamine autoreceptors in the amygdala: a fast cyclic voltammetric study in vitro. Neurosci Lett 134:41–44 50. Jones SR, Garris PA, Kilts CD, Wightman RM (1995) Comparison of dopamine uptake in the basolateral amydaloid nucleus, caudate-putamen, and nucleus accumbens of the rat. J Neurochem 64:2581–2589 51. Mundorf ML, Joseph JD, Austin CM, Caron MG, Wightman RM (2001) Catecholamine release and uptake in the mouse prefrontal cortex. J Neurochem 79:130–142 52. Cragg SJ, Baufreton J, Xue Y, Bolam JP, Bevan MD (2004) Synaptic release of dopamine in

53.

54.

55.

56.

57.

58.

59.

60.

Monitoring Dopamine Release

273

the subthalamic nucleus. Eur J Neurosci 20:1788–1802 Iravani MM, Kruk ZL (1997) Real-time measurement of stimulated 5-hydroxytryptamine release in rat substantia nigra pars reticulata brain slices. Synapse 25:93–102 Cragg SJ, Hawkey CR, Greenfield SA (1997) Comparison of serotonin and dopamine release in substantia nigra and ventral tegmental area: region and species differences. J Neurochem 69:2378–2386 John CE, Budygin EA, Mateo Y, Jones SR (2006) Neurochemical characterization of the release and uptake of dopamine in ventral tegmental area and serotonin in substantia nigra of the mouse. J Neurochem 96:267–282 Chen BT, Rice ME (2002) Synaptic regulation of somatodendritic dopamine release by glutamate and GABA differs between substantia nigra and ventral tegmental area. J Neurochem 8:158–169 Cragg SJ, Rice ME, Greenfield SA (1997) Heterogeneity of electrically-evoked dopamine release and uptake in substantia nigra, ventral tegmental area, and striatum. J Neurophysiol 77:863–873 Jones SR, Mickelson GE, Collins LB, Kawagoe KT, Wightman RM (1994) Interference by pH and Ca2+ ions during measurements of catecholamine release in slices of rat amygdala with fast-scan cyclic voltammetry. J Neurosci Methods 52:1–10 Wu Q, Reith ME, Wightman RM, Kawagoe KT, Garris PA (2001) Determination of release and uptake parameters from electrically evoked dopamine dynamics measured by real-time voltammetry. J Neurosci Methods 112:119–133 Paxinos G, Watson C (1998) The Rat Brain in Stereotaxic Coordinates, 4th edn. Academic, Inc

Chapter 16 Real-Time Chemical Measurements of Dopamine Release in the Brain James G. Roberts*, Leyda Z. Lugo-Morales*, Philip L. Loziuk, and Leslie A. Sombers Abstract Rapid changes in extracellular dopamine concentrations in freely moving or anesthetized rats can be detected using fast-scan cyclic voltammetry (FSCV). Background-subtracted FSCV is a real-time electrochemical technique that can monitor neurochemical transmission in the brain on a subsecond timescale, while providing chemical information on the analyte. Also, this voltammetric approach allows for the investigation of the kinetics of release and uptake of molecules in the brain. This chapter describes, completely, how to make these measurements and the properties of FSCV that make it uniquely suitable for performing chemical measurements of dopaminergic neurotransmission in vivo. Key words: Fast scan cyclic voltammetry, In vivo, Electrochemistry, Carbon fiber microelectrode

1. Introduction

Dopamine is a neurotransmitter of particular interest due to its involvement in motivated behavior and reward-driven learning (1, 2), as well as various neurological disorders such as Parkinson’s disease (3), schizophrenia (4), and drug addiction (5, 6). Extracellular dopamine concentrations in the brain vary on a subsecond timescale due to phasic firing of dopamine neurons (7). These naturally occurring dopamine transients are enhanced upon administration of drugs of abuse (8, 9), and become time-locked to cues that predict reward availability (1, 9–12). The ability to characterize and provide real-time measurements of rapidly

* These two authors contributed equally to this work. Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_16, © Springer Science+Business Media, LLC 2013

275

276

J.G. Roberts et al.

fluctuating dopamine in vivo requires an analytical method with rapid temporal resolution. To date, the most widely used technique to monitor in vivo neurotransmitter release is microdialysis (13). This method provides excellent chemical selectivity and is well suited for measuring dopamine levels averaged over the course of minutes to hours. However, microdialysis lacks the temporal resolution to detect phasic dopamine fluctuations that occur on the subsecond to second timescale (14). In contrast, electrochemical techniques are especially useful for monitoring rapid chemical changes resulting from discrete neurochemical events due to rapid sampling rates (micro-to-millisecond timescale). Broadly speaking, the techniques that involve current flow at an electrode under potential control can be divided into two groups: voltammetric and amperometric methods (15). Of these techniques, fast-scan cyclic voltammetry (FSCV) provides the best combination of temporal resolution, sensitivity, and chemical selectivity—features that are essential for the detection of rapid neurotransmitter fluctuations in vivo (16). Additionally, FSCV has been combined with other techniques including microinjection (7), iontophoresis (12, 17), and electrophysiology (12, 18, 19) to provide a great deal of new information regarding dopamine and its role in the brain. 1.1. Use of FSCV in Dopamine Detection

In FSCV, a dynamic potential is applied to a carbon fiber microelectrode (20). As the voltage is cycled through a triangular potential pattern, current is generated and recorded as a function of potential. The current generated over a single scan is plotted versus the applied potential. This resulting voltammogram is characteristic of the analyte, allowing it to be distinguished from many other electroactive species. This principle by which FSCV operates allows the identification of dopamine by the location of the potentials at which it oxidizes/reduces and the characteristic peak shape (Fig. 1). This also allows dopamine to be distinguished from many other interferents in the brain, such as ascorbic acid and changes in pH (16). Although FSCV generates characteristic voltammograms that can serve as qualitative identifiers for a molecule of interest, selectivity is always a concern. Several criteria need to be followed in order to positively confirm the identity of a voltammetric signal (21). These steps include electrochemical, anatomical, physiological, and pharmacological verification. 1. The in vivo signal should match the standard voltammogram collected in vitro. 2. The signal must be collected in an anatomical region known to be analyte rich. 3. Stimulation of cell bodies should illicit analyte release. 4. Pharmacological manipulation should be used to reduce or/and increase analyte release.

16

Measurements of Dopamine Release in the Brain

277

Fig. 1. Principles of background subtracted FSCV. (a) First, a potential waveform is applied to a working electrode. (b) This generates a stable non-faradaic background current. (c) The redox reaction will exchange electrons at the electrode surface, generating faradaic current. (d) At low analyte concentrations, the faradaic response (dashed red line) is small compared to the background current. (e) The stable non-faradaic background current can be subtracted from the faradaic current arising from the redox reaction. The resulting cyclic voltammogram is an electrochemical fingerprint of the analyte.

These steps must be taken in order to perform reliable electrochemical measurements and authenticate the identity of an analyte. The microelectrodes typically used with FSCV are cylindricallyshaped carbon-fiber microelectrodes. With this approach, a 5–7 μm diameter carbon fiber electrode is sealed in a glass capillary with a portion of the fiber (75–125 μm) extending from the tip. This carbon fiber electrode is approximately 40-fold shorter and 50-fold smaller in diameter than a typical microdialysis probe, and thus it is particularly well suited to probe brain regions that have gradations in the density of neuronal terminals over these dimensions (22). This small size results in minimal tissue damage during in vivo experiments, allows for characterization of specific brain regions, and its cylindrical shape enables detection from all sides of the electrode by way of hemispherical diffusion to the recording surface, which enhances sensitivity.

278

J.G. Roberts et al.

2. Materials 2.1. Electrode Fabrication

1. Carbon fiber: T-650 (GoodFellow, Huntingdon, England). 2. Borosilicate capillary glass: 0.6 mm O.D., 0.4 mm I.D. for freely-moving experiments and 1.0 mm O.D., 0.5 mm I.D. for anesthetized experiments (A-M Systems, Sequim, WA). 3. Vertical electrode puller: PE-21 (Narishige, Tokyo, Japan). 4. Vacuum pump. 5. Optical microscope. 6. Surgical scalpel to cut the carbon fiber. 7. Micromanipulator: for freely-moving experiments (custom made, UNC-CH Chemistry, Machine Shop). 8. Silver paint: Silver Print II (GC Electronics, Rockford, IL). 9. Heat shrink tubing: EPS-200-1/8″ and FP-301-3/32″ (3 M Electronics, Austin, TX). 10. NORIT A® activated carbon is used for isopropyl alcohol purification (MP Biomedicals, LLC, Solon, OH). 11. Silver wire: 0.5 mm diameter for Ag/AgCl reference electrode (Sigma-Aldrich, St. Louis, MO). 12. Gold connector: PCB socket (Newark Electronics, Chicago, IL). 13. Insulated leads: 30 gauge (Squires Electronics, Inc., Cornelius, OR).

2.2. Surgery

1. Anesthetic: xylazine and ketamine for freely-moving experiments, urethane for anesthetized experiments, and 0.25% bupivacaine is used as a local anesthetic. 2. Heating pads. 3. Stereotaxic frame: such as, Model 900 Small Animal Stereotaxic (David Kopf Instruments, Tujunga, CA). 4. Guide cannula (Bioanalytical Systems, Inc., West Lafayette, IN). 5. Anchor screws (Gexpro, Indianapolis, IN). 6. Cranioplastic cement: Grip Cement (Dentsply International, Inc., Milford, DE). 7. Stimulating electrode: 20 mm long bipolar stainless steel (Plastics One, Roanoke, VA).

2.3. Electrochemistry

1. Multifunction input/output cards: PCI-6251 and PCI-6711 (16 bit, 333 kHz) (National Instruments, Austin, TX). 2. Software for data collection and analysis: TH-1 (ESA, Chelmsford, MA), or custom written in house using LabVIEW (National Instruments, Austin, TX).

16

Measurements of Dopamine Release in the Brain

279

Fig. 2. Headstage (UNC-CH Electronics Design Facility). A miniaturized current-to-voltage convertor that consists of (a) operational amplifier, (b) threaded connection to stimulating electrode, (c) lead for reference electrode, (d) lead for working electrode, (e) DB-25 connector.

3. Potentiostat. One of the following is appropriate: EI-400 biopotentiostat (Cypress Systems, Lawrence, KS), Universal Electrochemistry Instrument (UEI, UNC-Chapel Hill, Electronics Design Facility), or Universal Headstage Controller (United World Domination, Mebane, NC). 4. Headstage: miniaturized current-to-voltage converter (UNC-CH Electronics Design Facility; or United World Domination, Mebane, NC) (Fig. 2). A larger version can be used for anesthetized experiments and postexperiment calibration. 5. Commutator: 25 channel (Crist Instruments, Hagerstown, MD). 6. Screened behavioral chamber: custom made for in vivo experiments (Med Associates Inc., St. Albans, VT). 7. Optional equipment: TV, DVD-R, and video character generator (for monitoring, recording, and time-stamping animal behavior). 2.4. Stimulation

1. Multifunction input/output Instruments, Austin, TX).

card:

PCI-6711

(National

2. Bi-phasic stimulus isolator: DS4 (Digitimer, Ltd, Hertfordshire, England). 2.5. Electrode Postcalibration

1. Dopamine HCl: 1 mM dopamine in 0.1 N HClO4 for stock solutions, and dilutions are made in buffer (Sigma-Aldrich, St. Louis, MO).

280

J.G. Roberts et al.

2. Flow injection apparatus: six-port, two-position high-performance liquid chromatography (HPLC) valve, with air actuator, and digital valve interface (VICI, Houston, TX). 3. Grounded Faraday cage: custom built in house. 4. Tris buffer: 3.25 mM KCl, 1.2 mM CaCl2, 1.2 mM MgCl2, 2.0 mM Na2SO4, 1.25 mM NaH2PO4, 145 mM NaCl, and 15 mM Trizma ® HCl at pH 7.4 (Sigma-Aldrich, St. Louis, MO).

3. Methods 3.1. Electrochemistry, Instrumentation, and Software

Dopamine release in brain tissue can be monitored in real-time with high spatial and temporal resolution when micron-scale electrodes and low-noise instrumentation are implemented. Dopamine is electrochemically detected at carbon-fiber microelectrodes by applying a potential sufficient to liberate two electrons from dopamine to form dopamine ortho-quinone. This provides a current that can be converted to a voltage and measured using a current transducer. Instrumentation includes the Universal Electrochemical Instrument, the Universal Headstage Controller, or a EI-400 biopotentiostat. These instruments are generally used with computercontrolled interface boards and locally written software (LabVIEW, National Instruments, Austin, TX). Software is commercially available from ESA. The instrument provides all inputs and supplies power to the headstage, and usually consists of two main components: a low-pass filter and a headamp module. The computergenerated waveform contains digitization noise that must be smoothed by a low-pass filter before the signal reaches the working electrode. The output voltage from the current transducer is further amplified and conditioned by the headstage amplifier. The interface boards are responsible for the digital-to-analog and analog-to-digital conversions that are transmitted to and from the headstage, respectively. The use of FSCV for the electrochemical detection of dopamine at carbon-fiber microelectrodes requires a waveform that optimizes peak currents, response time, and chemical selectivity. The most commonly used waveform holds the working electrode at −0.4 V vs. Ag/AgCl with periodic ramping to +1.3 V and back at a rate of 400 V/s and a frequency of 10 Hz (Fig. 1). The time between scans when the working electrode is held at a negative potential allows positively-charged dopamine to preconcentrate at the electrode surface (23). Due to the fast scan rate, scanning generates a large capacitive charging current at the electrode surface (24), which is significantly larger than faradaic currents resulting from redox processes at the microelectrode surface.

16

Measurements of Dopamine Release in the Brain

281

These background currents are stable over tens of seconds. This allows for subtraction, revealing the interesting faradaic responses. The resulting background-subtracted cyclic voltammograms provide information on the analyte identity, redox potentials, reversibility, and electron transfer kinetics. The shape of the peaks allows for the discrimination of multiple species (however all catecholamines produce similar voltammograms) and can be used to assess the role of mass transfer. The amplitude of the peaks can be correlated to the concentration of the analyte at the electrode surface. Under the conditions described, the cyclic voltammograms for dopamine should have a peak for the oxidative current at around +0.6 V. 3.2. Bipolar Electrical Stimulation

3.3. Electrode Fabrication 3.3.1. Carbon Fiber Microelectrode

Electrical stimulation of dopaminergic cell bodies evokes dopamine release from the terminals in a time-locked manner, enabling the experimenter to monitor the kinetics of dopamine release and uptake with FSCV (25). The computer controlled stimulation is delivered with a biphasic stimulus isolator to the stimulating electrode. The device must be calibrated before use to ensure proper function. The waveform applied to the stimulating electrode is a biphasic square wave that is applied with a frequency, amplitude, pulse width, and number of pulses consistent with the experimental goals. Typical stimulation parameters for dopamine neuron cell bodies are 125 biphasic pulses, 60 Hz, ±125–150 μA, and 2 ms/ phase. This stimulation must be applied between the ramps of the electrochemical waveform, such that the electrochemical data is not disturbed by the current stimulation (Fig. 3). 1. A single carbon fiber is placed on a flat and clean surface that is well illuminated. The fiber is then aspirated into a borosilicate glass capillary, so that it extends from both ends. 2. The filled capillary is tapered in an electrode puller. This forms two electrodes from a single filled capillary. Each is inspected

Fig. 3. Electrical stimulation. The bipolar electrical stimulation (gray), must not overlap with the applied electrochemical waveform (black ).

282

J.G. Roberts et al.

Fig. 4. Micromanipulator (UNC-CH Machine shop). An illustration of a loaded micromanipulator, ready for an experiment.

under the microscope to ensure a tight glass seal around the carbon fiber. 3. The exposed carbon fiber is then cut to length (~100 mm) with a sharp scalpel under a microscope using a magnification of 10×. The electrode should also be inspected under the microscope using a higher magnification of at least 40× for visible cracks or abnormalities in the fiber or glass seal, and discarded if any are present (see Note 1). 4. In freely-moving experiments: (a) An inspected 100 mm carbon fiber microelectrode is loaded into a custom micromanipulator and secured with heat shrink tubing (Fig. 4). (b) A small diameter insulated wire is painted with silver paint, and fed into the back of the capillary to make an electrical connection with the carbon fiber. A slight rotation of the wire ensures connectivity with the carbon fiber. The wire is secured to the micromanipulator with additional heat shrink tubing. (c) All loaded manipulators are stored with the exposed carbon in purified and filtered isopropyl alcohol.

16

Measurements of Dopamine Release in the Brain

283

5. In anesthetized experiments: (a) Larger diameter glass capillaries can be used. (b) The carbon fiber microelectrode is backfilled using a saturated solution of 150 mM potassium chloride and 4 M potassium acetate. A small diameter insulated wire is fed into the back of the capillary to make electrical connection. 3.3.2. Ag/AgCl Reference Electrode

1. A piece of silver wire is cut to approximately 10 mm, inserted into the socket of a gold connector, and soldered in place. 2. The solder is then covered with quick dry epoxy to avoid the contact of the soldering material with tissue. 3. On the day of surgery, the reference is chlorinated by connecting the positive terminal of a 2.5 V power supply to the gold pin on the silver wire and the negative terminal to a wire, with both leads immersed in 0.1 M hydrochloric acid. Chlorination is performed for about 1 min until the surface of the silver wire turns slightly white.

3.4. Surgery 3.4.1. Anesthetized Preparation

1. The rat is anesthetized with urethane (3 g/kg i.p.), the top of the head is shaved, and the animal is placed in a stereotaxic frame. 2. The scalp is locally anesthetized with a subcutaneous injection of 0.25% bupivacaine. An incision is made in the scalp, and the skin retracted to expose a 15–20 mm longitudinal and 10–15 mm lateral area of cranium. 3. Holes are drilled through the skull for stereotaxic placement of electrodes (stimulating, working, reference) (Fig. 5). The stimulating electrode can be positioned either in regions

Fig. 5. A top view illustration of a rat skull, highlighting the general placement of holes (dotted lines) for electrode and surgical screw placement.

284

J.G. Roberts et al.

containing dopaminergic cell bodies (substantia nigra/ventral tegmental area) or at the ascending fibers of the medial forebrain bundle. The hole for the working electrode is drilled above the target terminal region (e.g., 1.3 mm lateral and 1.3 mm rostral from bregma for the caudate-putamen and the core of the nucleus accumbens, and +1.7 mm anterior and +0.8 mm lateral for the nucleus accumbens shell). The hole for the reference electrode is drilled in the contralateral hemisphere, opposite the working electrode. 4. Electrodes are lowered and secured in select areas of the brain using micromanipulators mounted on the stereotaxic frame. The reference electrode is secured with cranioplastic cement. Mild electrical stimulations are applied to evoke neurotransmitter release that is monitored at the microelectrode using FSCV. The rat is maintained on a heated pad for the duration of the experiment. 3.4.2. Freely-Moving Preparation

1. The rat is anesthetized with intramuscular or intraperitoneal ketamine (100 mg/kg) and intramuscular xylazine (10 mg/kg), the top of the head is shaved, and the animal is placed in a stereotaxic frame. 2. The scalp is locally anesthetized with a subcutaneous injection of 0.25% bupivacaine. An incision is made in the scalp, and the skin retracted to expose a 15–20 mm longitudinal and 10–15 mm lateral area of cranium. 3. Holes are drilled for the working electrode guide cannula, stimulating and reference electrodes (Fig. 5). In addition, four holes are drilled at a 45° angle into which anchor screws are secured. 4. Reference electrode and guide cannula are lowered using micromanipulators mounted on the stereotaxic frame. 5. Once the components are in place, they are secured with cranioplastic cement, leaving the stimulating electrode hole exposed. 6. The stimulating electrode is modified in order to provide adequate space between the plastic hub of the stimulating electrode and the guide cannula (for the working electrode) on the animal’s head cap (Fig. 6). The stimulating electrode wires are bent at a 90° angle from the plastic hub and then bent back down at another 90° angle, to give a horizontal distance of ~5 mm between the hub and the main axis of the wires. Next, the tips are separated by 0.8–1.0 mm and carefully cut to a uniform length without disturbing the insulation. Dura mater is thoroughly cleared and the electrode is stereotaxically lowered into the tissue, oriented so that the tips of the electrode are splayed on the coronal plane. The electrode is lowered to 1 mm above the target brain region.

16

Measurements of Dopamine Release in the Brain

285

Fig. 6. Stimulating electrode.

7. The stimulating electrode is connected to the stimulator and a mild electrical stimulation is applied through the stimulating electrode. The animal’s tail should respond to this stimulation by rapidly rising and then slowly falling back to the resting position. The stimulating electrode is lowered in 0.2 mm increments until this response is diminished. It is then lowered further in 0.1 mm increments until this tail response is almost non-detectable. 8. Finally, cranioplastic cement is applied to the exposed cranium, carefully covering the stimulating electrode and lower half of its plastic hub. 9. Immediately following surgery, the animal is placed on a warm heating pad until fully recovered. Once fully awake, soft food and fresh water are offered with a fruit-flavored analgesic, such as acetaminophen (0.1–0.3 g/kg) that the rat will readily lick. 10. The animal is monitored daily and gently handled to facilitate experimental procedures. While handling, the stylet should be gently removed from the guide cannula, cleaned with an alcohol wipe, and reinserted. Experiments can be conducted within 2–5 days of surgery.

286

J.G. Roberts et al.

3.5. Freely-Moving Rat Experiment 3.5.1. Making the Connections

3.5.2. Lowering the Carbon Fiber Microelectrode

Two to five days after surgery, depending on the rat’s postsurgical recovery, the animal is prepared for the experiment. The animal is placed in the behavioral chamber, tethered using the stimulator cable on which the headstage is mounted (Fig. 2) and allowed to acclimate for about 10 min. Before the loaded micromanipulator is placed into the guide cannula, the electrode is inspected once again under a microscope to double check the condition of the seal. The electrode is retracted inside the micromanipulator as the tip of the electrode is monitored. Once the electrode tip disappears, each turn is counted until the electrode is fully retracted. This protects the electrode integrity as it is loaded into the cannula and allows the experimenter to index the tip location inside the manipulator. All connections are cleaned, and the guide cannula stylet is removed and replaced with the micromanipulator containing the retracted microelectrode. The manipulator is locked in place and the working and reference electrodes are connected to the headstage. The electrode is slowly lowered into tissue as its output is monitored on an oscilloscope. To do this, the waveform is applied. As soon as the carbon fiber electrode comes in contact with tissue, the non-faradaic background current appears, and is monitored for stability as the electrode is lowered through the tissue (Fig. 7).

Fig. 7. Oscilloscope output. (a) Diagram of applied waveform. (b) Electrode response when circuit is completed in tissue or buffer.

16

Measurements of Dopamine Release in the Brain

287

A break in the electrode is evident by a sudden change in the shape of the background current to a more resistive profile (approximating a triangular wave), and it should be removed and replaced with a fresh carbon fiber electrode. Once in place at the target region, the electrode is conditioned for about 20 min to stabilize the signal. Electrochemical conditioning consists of applying the triangular waveform mentioned above for at least 10 min at a frequency of 60 Hz and then changing it to 10 Hz for 10 additional minutes of potential cycling. A mild electrical stimulation is applied to the stimulating bipolar electrode while the current output is monitored at the carbon fiber microelectrode. If a dopamine signal is not obtained, the working electrode is lowered in small increments and stimulation repeated until electrically evoked dopamine release is observed. The electrode is then secured in position by a locking device on the micromanipulator and the experiment is initiated (see Note 2). 3.6. Anesthetized Rat Experiment

1. Immediately following surgery, the stereotaxic frame is placed into the grounded Faraday cage. 2. The electrodes (carbon fiber, stimulating, and Ag/AgCl reference) are lowered into the appropriate holes using the stereotaxic frame. There is no need to use screws and cranioplastic cement to secure the electrodes in an anesthetized experiment; however, the reference can be secured in place for stability. 3. The stimulating electrode is connected to the biphasic stimulus isolator and the working and reference electrodes are connected to the headstage. 4. As described above, the microelectrode is lowered in small increments (0.1 mm) into a brain region rich in dopamine terminals. 5. Dopamine neurons are electrically stimulated to illicit dopamine release at the terminals in a time-locked fashion (see Note 3).

3.7. After the Experiment

Upon completion of the experiment(s), there are two options depending on the objective of the experiment and the investigator’s primary interest. These two options are described below.

3.7.1. Verification of Electrode Placement

The electrode tip is too small to leave a visible mark in tissue, thus an electrical lesion is made at the carbon fiber tip by applying a high current to the microelectrode. This unequivocally shows the location of the electrode in the tissue; however, this renders the electrode useless and it cannot be calibrated. The rat is transcardially perfused with 0.9% saline and 10% formalin solution to fix brain tissue. Finally, the animal is decapitated and the brain is removed from the skull and stored in formalin solution at 4°C, until it is sliced for histology.

288

J.G. Roberts et al.

Fig. 8. Flow-injection analysis system. A syringe pump supplies a constant buffer flow across the working and reference electrodes. An HPLC valve controls the introduction of an analyte to the working electrode surface.

3.7.2. Electrode Postcalibration

Alternatively, the microelectrode is carefully removed from the brain, replaced with a sacrificial carbon fiber microelectrode, and an electrical lesion is made as described above. The electrode for calibration is rinsed in water and calibrated in vitro on a flowinjection apparatus using known physiological concentrations of dopamine (usually between 200 and 1,000 nM). This system consists of a custom-made electrochemical cell and a sample loop by which small volumes of analyte are rapidly injected into the cell using a six-port HPLC valve and a computer-controlled pneumatic actuator. A syringe pump is used to continuously supply physiological buffer at a constant flow rate through the electrochemical cell (Fig. 8.). The working electrode is lowered with a micromanipulator into the stream of buffer flowing at 1–3 mL/min. The Ag/ AgCl reference electrode is submerged in the buffer as well and both are connected to the headstage. The same waveform used for the in vivo experiment is applied for the calibration of the electrode. Concentrations of the analyte of interest are loaded into the sample loop and introduced into the electrochemical cell with the digitally-controlled pneumatic actuator. The injection is software controlled. Each concentration of dopamine is sampled at least in triplicate and the averaged peak oxidative current is plotted against concentration. The resulting calibration plot is used to relate the current collected in vivo to corresponding dopamine concentrations.

3.8. Data Analysis

TH-1 (ESA, Chelmsford, MA) software is commercially available and can be used for data analysis. Additionally, custom software

16

Measurements of Dopamine Release in the Brain

289

written with Matlab (MathWorks, Inc., Natick, MA) can mathematically extract information from chemical data for quantitative analysis. The current method of multivariate statistical analysis involves the use of principle component regression (PCR) (16, 28). PCR has the ability to separate intensity based data into relevant components and noise, so that noise can be discarded. A training (calibration) set is used that includes individual cyclic voltammograms for the major species (typically dopamine and pH shifts, depending on the local microcircuitry) at various concentrations. Principle components that best describe the data are chosen. A principle component can be described as a vector that passes through the data that includes the most information. These principle components are then used to predict unknown concentrations from individual cyclic voltammograms collected in vivo, as long as the unknown concentrations fit within the training set. 3.9. FSCV Combined with Electrophysiology

FSCV can be combined with more traditional neuroscience tools such as electrophysiology, a technique that uses an electrode to measure action potentials (12, 18, 29). With this combined approach, the microelectrode employed for electrochemical detection is also used to monitor local synaptic activity. Between scans the holding potential is abbreviated and the electrode is allowed to float, thereby adopting the potential of its local environment, which is digitally recorded. The use of this method has allowed dopamine release to be correlated with changes in the firing of specific neurons in the vicinity of the electrode, shedding light on dopamine function in discrete brain microcircuits.

3.10. FSCV Combined with Intracranial Self-Stimulation

The behavioral paradigm of intracranial self-stimulation (ICSS) is an intensely rewarding experimental model (30). It has also been combined with FSCV (10, 12, 31). In ICSS a stimulating electrode is implanted into a specific brain nucleus and the animal is taught to use a lever to deliver a mild electrical stimulation to the chosen region. This serves as a powerful operant reinforcer and is often used in studies of motivated behavior. The use of ICSS in combination with FSCV has led to the association of rapid dopamine signaling with learned cues (such as audio or visual cues) that precede an electrical stimulation or reward availability (10, 12). This technique reveals information on the chemical mechanisms underlying reward based learning.

3.11. FSCV and Methods of Localized Pharmacological Manipulation

Microinjection and iontophoresis are two methods that have been implemented for administering small quantities of a compound into a specific region of the brain. While systemic application of drugs affects global brain circuitry, localized drug delivery techniques allow the experimenter to pharmacologically manipulate one discrete brain region. Microinjection involves the placement of a small needle into the desired brain location, and the

290

J.G. Roberts et al.

Fig. 9. Iontophoresis probe. Scanning electron micrograph of a five-barrel probe coupling FSCV with iontophoresis, using a carbon fiber microelectrode. Reprinted with permission from ref. (36). Copyright 2008 American Chemical Society.

subsequent pressure-driven infusion of a compound. Intracranial self-administration using microinjection has been used to elucidate the reinforcing action of specific agents in precise brain nuclei (32–34). Additionally, microinjection combined with FSCV has established that dopamine transients recorded in the nucleus accumbens shell require phasic neuronal activity in the ventral tegmental area (7), linking the activity of these two regions. Iontophoresis can be used to locally apply compounds in an ionic solution using an applied current (35). When combined with FSCV (17, 36), capillary barrels are attached to a working electrode to deliver small quantities of a compound into tissue (Fig. 9). This approach enables drug administration to the same site as the working electrode. While it has its advantages, iontophoresis is a largely nonquantitative technique. However, recent advances have allowed researchers to accurately quantify the amount of drug delivered during an iontophoretic ejection by the use of an electroosmotic flow marker (17, 36). This marker allows quantitative analysis by taking into account the variability due to inconsistent barrel dimensions that affect electroosmosis, which in turn affects the observed iontophoretic ejection. 3.12. Recent Advances

One drawback to the use of FSCV in freely-moving experiments has been the cable which tethers the animal. The introduction of wireless integrated circuits has created new opportunities for studying dopamine function in freely-moving animals (37). Advantages of this technology include the ability to perform measurements during multiple animal social interactions, investigation of more natural behaviors and more complex environments, and fewer artifacts introduced during movement of electrical connections. Another

16

Measurements of Dopamine Release in the Brain

291

Fig. 10. Microelectrode array. An array of four carbon fiber microelectrodes, with fused silica insulation, secured with a fixed separation of 250 μm. Reprinted with permission from ref. 41 Copyright 2010 Elsevier.

recent development has been the incorporation of analog background subtraction to enable recordings over 30 min time intervals before distortion of dopamine signals occurs due to background drift (38). Other developments, such as microelectrode arrays (Fig. 10), allow multiple electrodes to be used in a single experiment (39–41). This has opened up the opportunity for researchers to simultaneously measure dopamine release at spatially discrete brain locations (39, 41). This approach also allows for the simultaneous detection of multiple signaling agents at various locations (40) and allows for more representative data to be obtained because of the increased number of recordings that can be acquired in a given experiment. Another recent advance has enabled chronic implantation of microelectrodes, enabling recording at the same electrode over months, rather than hours (42, 43). Finally, as an alternative to electrical stimulation, optogenetics can be used to stimulate specific neuronal populations using light-activated ion channels (44).

4. Notes 1. Optimal electrode length is determined by instrument limitations and experimental goals. 2. Many experiments monitor naturally-occurring dopamine fluctuations or transients. These dopamine release events are

292

J.G. Roberts et al.

evident at some but not all sites that support electrically-evoked dopamine release (26). Also, after several days of implantation, the reference electrode may drift by about 0.2 V, requiring the applied potential to be offset by 0.2 V. 3. Naturally-occurring transient dopamine release events are not generally detected in anesthetized animals, unless pharmacologically evoked (27).

Acknowledgments This work was funded in part by grants from the National Institutes of Health, the National Science Foundation, and NCSU Department of Chemistry. In addition, we gratefully acknowledge our coworkers, past and present, for the studies cited in this review. References 1. Day JJ, Roitman MF, Wightman RM, Carelli RM (2007) Associative learning mediates dynamic shifts in dopamine signaling in the nucleus accumbens. Nat Neurosci 10:1020–1028 2. Schultz W (2007) Behavioral dopamine signals. Trends Neurosci 30:203–210 3. Obeso JA, Rodriguez-Oroz MC, Goetz CG, Marin C, Kordower JH, Rodriguez M, Hirsch EC, Farrer M, Schapira AH, Halliday G (2010) Missing pieces in the Parkinson’s disease puzzle. Nat Med 16:653–661 4. Grace AA (1991) Phasic versus tonic dopamine release and the modulation of dopamine system responsivity: a hypothesis for the etiology of schizophrenia. Neuroscience 41:1–24 5. Carelli RM, Wightman RM (2004) Functional microcircuitry in the accumbens underlying drug addiction: insights from real-time signaling during behavior. Curr Opin Neurobiol 14:763–768 6. Di Chiara G, Bassareo V (2007) Reward system and addiction: what dopamine does and doesn’t do. Curr Opin Pharmacol 7:69–76 7. Sombers LA, Beyene M, Carelli RM, Wightman RM (2009) Synaptic overflow of dopamine in the nucleus accumbens arises from neuronal activity in the ventral tegmental area. J Neurosci 29:1735–1742 8. Cheer JF, Wassum KM, Sombers LA, Heien ML, Ariansen JL, Aragona BJ, Phillips PE, Wightman RM (2007) Phasic dopamine release evoked by abused substances requires

9.

10.

11.

12.

13.

14.

15.

16.

cannabinoid receptor activation. J Neurosci 27:791–795 Phillips PE, Stuber GD, Heien ML, Wightman RM, Carelli RM (2003) Subsecond dopamine release promotes cocaine seeking. Nature 422:614–618 Owesson-White CA, Cheer JF, Beyene M, Carelli RM, Wightman RM (2008) Dynamic changes in accumbens dopamine correlate with learning during intracranial self-stimulation. Proc Natl Acad Sci U S A 105:11957–11962 Roitman MF, Stuber GD, Phillips PE, Wightman RM, Carelli RM (2004) Dopamine operates as a subsecond modulator of food seeking. J Neurosci 24:1265–1271 Cheer JF, Aragona BJ, Heien ML, Seipel AT, Carelli RM, Wightman RM (2007) Coordinated accumbal dopamine release and neural activity drive goal-directed behavior. Neuron 54:237–244 Justice JB Jr (1993) Quantitative microdialysis of neurotransmitters. J Neurosci Methods 48:263–276 Lu Y, Peters JL, Michael AC (1998) Direct comparison of the response of voltammetry and microdialysis to electrically evoked release of striatal dopamine. J Neurochem 70:584–593 Michael AC, Borland LM (eds) (2007) Electrochemical methods for neuroscience, vol 1, 1st edn. CRC Press, Boca Raton Heien ML, Johnson MA, Wightman RM (2004) Resolving neurotransmitters detected

16

17.

18.

19.

20.

21.

22. 23.

24.

25.

26.

27.

28.

Measurements of Dopamine Release in the Brain

by fast-scan cyclic voltammetry. Anal Chem 76:5697–5704 Herr NR, Belle AM, Daniel KB, Carelli RM, Wightman RM (2010) Probing presynaptic regulation of extracellular dopamine with iontophoresis. ACS Chem Neurosci 1:627–638 Kuhr WG, Wightman RM, Rebec GV (1987) Dopaminergic neurons: simultaneous measurements of dopamine release and single-unit activity during stimulation of the medial forebrain bundle. Brain Res 418:122–128 Millar J, Stamford JA, Kruk ZL, Wightman RM (1985) Electrochemical, pharmacological and electrophysiological evidence of rapid dopamine release and removal in the rat caudate nucleus following electrical stimulation of the median forebrain bundle. Eur J Pharmacol 109:341–348 Robinson DL, Hermans A, Seipel AT, Wightman RM (2008) Monitoring rapid chemical communication in the brain. Chem Rev 108:2554–2584 Phillips PEM, Wightman RM (2003) Critical guidelines for validation of the selectivity of invivo chemical microsensors. Trac-Trend Anal Chem 22:509–514 Arbuthnott GW, Wickens J (2007) Space, time and dopamine. Trends Neurosci 30:62–69 Bath BD, Michael DJ, Trafton BJ, Joseph JD, Runnels PL, Wightman RM (2000) Subsecond adsorption and desorption of dopamine at carbon-fiber microelectrodes. Anal Chem 72:5994–6002 Bard AJ, Faulkner LR (2001) Electrochemical methods: fundamentals and applications, 2nd edn. John Wiley, New York Wightman RM, Amatore C, Engstrom RC, Hale PD, Kristensen EW, Kuhr WG, May LJ (1988) Real-time characterization of dopamine overflow and uptake in the rat striatum. Neuroscience 25:513–523 Wightman RM, Heien ML, Wassum KM, Sombers LA, Aragona BJ, Khan AS, Ariansen JL, Cheer JF, Phillips PE, Carelli RM (2007) Dopamine release is heterogeneous within microenvironments of the rat nucleus accumbens. Eur J Neurosci 26:2046–2054 Venton BJ, Wightman RM (2007) Pharmacologically induced, subsecond dopamine transients in the caudate-putamen of the anesthetized rat. Synapse 61:37–39 Keithley RB, Heien ML, Wightman RM (2009) Multivariate concentration determination using principal component regression with residual analysis. Trends Anal Chem 28:1127–1136

293

29. Williams GV, Millar J (1990) Concentrationdependent actions of stimulated dopamine release on neuronal activity in rat striatum. Neuroscience 39:1–16 30. Olds J, Milner P (1954) Positive reinforcement produced by electrical stimulation of septal area and other regions of rat brain. J Comp Physiol Psychol 47:419–427 31. Garris PA, Kilpatrick M, Bunin MA, Michael D, Walker QD, Wightman RM (1999) Dissociation of dopamine release in the nucleus accumbens from intracranial self-stimulation. Nature 398:67–69 32. Ikemoto S, Qin M, Liu ZH (2005) The functional divide for primary reinforcement of D-amphetamine lies between the medial and lateral ventral striatum: is the division of the accumbens core, shell, and olfactory tubercle valid? J Neurosci 25:5061–5065 33. Ikemoto S, Qin M, Liu ZH (2006) Primary reinforcing effects of nicotine are triggered from multiple regions both inside and outside the ventral tegmental area. J Neurosci 26:723–730 34. Ikemoto S, Sharpe LG (2001) A head-attachable device for injecting nanoliter volumes of drug solutions into brain sites of freely moving rats. J Neurosci Methods 110:135–140 35. Rebec GV, Bashore TR (1984) Critical issues in assessing the behavioral effects of amphetamine. Neurosci Biobehav Rev 8:153–159 36. Herr NR, Kile BM, Carelli RM, Wightman RM (2008) Electroosmotic flow and its contribution to iontophoretic delivery. Anal Chem 80:8635–8641 37. Garris PA, Ensman R, Poehlman J, Alexander A, Langley PE, Sandberg SG, Greco PG, Wightman RM, Rebec GV (2004) Wireless transmission of fast-scan cyclic voltammetry at a carbon-fiber microelectrode: proof of principle. J Neurosci Methods 140:103–115 38. Hermans A, Keithley RB, Kita JM, Sombers LA, Wightman RM (2008) Dopamine detection with fast-scan cyclic voltammetry used with analog background subtraction. Anal Chem 80:4040–4048 39. Zachek MK, Park J, Takmakov P, Wightman RM, McCarty GS (2010) Microfabricated FSCV-compatible microelectrode array for real-time monitoring of heterogeneous dopamine release. Analyst 135:1556–1563 40. Zachek MK, Takmakov P, Moody B, Wightman RM, McCarty GS (2009) Simultaneous decoupled detection of dopamine and oxygen using pyrolyzed carbon microarrays and fast-scan cyclic voltammetry. Anal Chem 81:6258–6265 41. Zachek MK, Takmakov P, Park J, Wightman RM, McCarty GS (2010) Simultaneous moni-

294

J.G. Roberts et al.

toring of dopamine concentration at spatially different brain locations in vivo. Biosens Bioelectron 25:1179–1185 42. Clark JJ, Sandberg SG, Wanat MJ, Gan JO, Horne EA, Hart AS, Akers CA, Parker JG, Willuhn I, Martinez V, Evans SB, Stella N, Phillips PE (2010) Chronic microsensors for longitudinal, subsecond dopamine detection

in behaving animals. Nat Methods 7:126–129 43. Garris PA (2010) Advancing neurochemical monitoring. Nat Methods 7:106–108 44. Stuber GD, Hnasko TS, Britt JP, Edwards RH, Bonci A (2010) Dopaminergic terminals in the nucleus accumbens but not the dorsal striatum corelease glutamate. J Neurosci 30:8229–8233

Chapter 17 The MPTP/Probenecid Model of Progressive Parkinson’s Disease Anna R. Carta, Ezio Carboni, and Saturnino Spiga Abstract Parkinson’s disease (PD) is characterized by a progressive degeneration of dopamine (DA) neurons and a chronic loss of motor functions. The investigation of progressive degenerative mechanisms and possible neuroprotective approaches for PD depends upon the development of an experimental animal model that reproduces the neuropathology observed in humans. This chapter describes the generation of the 1-methyl4-phenyl-1,2,3,6-tetrahydropyridine/probenecid (MPTPp) chronic mouse model of PD. This model displays key features of PD, including impairment of motor and olfactory functions associated with partial loss of tyrosine hydroxylase-positive neurons and DA levels in the brain. The MPTPp mouse model provides an important tool for the study of mechanisms contributing to the pathological dysfunction of PD at the cellular and whole animal level. Key words: Neuropathology, Parkinson’s disease, Neuroprotection, Animal model, Preclinical studies

1. Introduction Parkinson’s disease (PD) is a neurodegenerative disorder characterized by a progressive death of nigrostriatal dopamine (DA) neurons and loss of striatal DA content. Symptoms of the disease also have a gradual onset and progression (1). Specific nonmotor deficits, such as olfactory dysfunction and gastrointestinal constipation, are generally considered early symptoms that often precede motor deficits. Appearance of motor symptoms such as bradykinesia, muscle rigidity and tremors, reflect the progressive nigrostriatal degeneration, generally becoming manifest when striatal DA drops to 20–30% of physiological levels, and nigral neurons are largely lost (2). The time-lapse between initial neuronal degeneration and occurrence of motor symptoms is likely attributable Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3_17, © Springer Science+Business Media, LLC 2013

295

296

A.R. Carta et al.

to the development of compensatory mechanism at the striatal level (3). Moreover, motor deficits are associated with postsynaptic functional changes in striatal medium-sized neurons (4, 5). Classically, a reduced expression of mRNA for dynorphin peptide in striatonigral neurons and an overexpression of mRNA for enkephalin in striatopallidal neurons are thought to represent measurable markers of these changes. On a neuropathological basis, diseased dopaminergic neurons display an unbalanced neuronal network, due to a complex scenario of malfunctioning cellular components, including oxidative stress, impaired protein disposal systems, and chronic neuroinflammation (6–10). Moreover, intracellular formations named Lewy bodies and dystrophic neuritis are found in nigral neurons as well as in other brain areas such as the olfactory tubercle and locus ceruleous, and represent a cellular hallmark of PD (11). Given its progressive nature, any neuroprotective intervention aimed at modifying the prognosis of the disease should optimally be commenced at the early stages of PD. For this reason, preclinical investigation of PD neuropathology and testing of neuroprotective strategies should rely on experimental models that reproduce the progressive nature of PD pathology. More generally, animal models of PD should possess the highest number of features of human PD (face validity), the underlying neuropathology should evolve as much as possible as PD and should respond to treatments in a manner comparable to human PD (predictive validity). Lastly, they should also reproduce the complex scenario of multiple interactions between neuronal elements and surrounding cells (construct validity) (12, 13). A number of neurotoxin-based or genetic models that imply the degeneration of dopaminergic nigrostriatal neurons have been developed, although most of them present limitations (14–18). One important limit is the rapid and transient neurodegeneration, which occlude the development of chronic pathogenic mechanisms and/or motor disabilities. Models based on the delivery of neurotoxin 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP), that offer the advantage of selectively targeting dopaminergic neurons, have been largely characterized in the past decade (19–23). MPTP treatment in mice, being easy to handle, affordable and highly reproducible, remains to date the most widely used in vivo model of PD in preclinical studies. MPTP enters the brain after systemic administration. Here, MPTP undergoes a two-step biotransformation process to form the toxic metabolite MPP + (24). First, MPTP is converted to MPDP + by MAO-B in astrocytes. In a second step MPDP + spontaneously oxidizes to MPP+. Released MPP + is specifically taken up by dopaminegic cells via the dopamine transporter (DAT). Importantly, both dopaminergic terminals in the striatum and

17

MPTPp Model of PD

297

dopaminergic dendrites in the SN express DAT, therefore MPP + toxicity is possibly directed at both sites. Inside the cell, MPP + can be sequestered into synaptosomal vesicles via the vesicular monoamine transporters (VMATs), or it can accumulate into mitochondria, which are the subcellular target of toxicity. Hence MPP+, by binding to complex I of the respiratory chain, blocks electron transport leading to ATP depletion, increased oxidative stress, and finally disruption of cellular energy metabolism and cell death (24). Rodents are less sensitive to MPTP toxicity when compared to primates. However, the C57black/6 strain is sensitive to appropriate MPTP protocols of administration, also offering the advantage of being selective in terms of targeting the nigrostriatal neurons (25, 26). MPTP toxicity in mouse is strictly dependent on the dose and paradigm of administration. Thus, the acute and subacute administration models often fail to induce motor disabilities, loss of TH-immunopositive cells in the SNc or persistency of the neurodegeneration after the neurotoxin treatment has been completed (24, 27). In the last decade, several research groups have devoted efforts to the characterization of a chronic protocol of MPTP administration in mice, delivered in association with the clearance inhibitor probenecid (MPTPp). As enumerated below, this protocol overcomes most of the limitations presented by shorter MPTP administration models, leading to validation of a preclinical model of progressive PD (23, 28–30). 1. Administration of MPTPp, in a 5 week chronic protocol, induces the gradual appearance of non-motor and motor symptoms, underlined by a progressive depletion of nigral dopaminergic cells, as shown by TH-immunoreactivity (23, 28). The olfactory dysfunction is detectable as early as after 1 MPTPp administration, whereas motor impairment can be assessed after the third administration, becoming overtly evident only after 7 MPTPp administrations (i.e., in the fourth week of the treatment) (23) (Fig. 1). 2. Dopaminergic cell loss in the SNc is observed after 3 MPTPp administrations and gradually increases at the end of treatment (23) (Fig. 2). Unlike shorter MPTP protocols, the MPTPp chronic administration induced an enduring depletion of dopaminergic neurons, as assessed up to 6 months after the end of treatment (23, 28). 3. After chronic MPTPp typical PD hallmarks such as α-synucleinimmunopositive deposits are present in the SNc (29), (Fig. 2), whereas progressively increasing CD11b-immunoreactivity reflecting microgliosis has been described, starting before neurodegeneration can be measured (23). In addition signs of neuronal apoptosis are detected in the SNc at the end of treatment (30).

298

A.R. Carta et al. vehicle

100 pellet retrieval time (sec)

Olfactory test

*

MPTPp

*

75

*

* 50

25

0 1

3

7

10

administrations

vehicle

5 number of errors per step

Beam traversal test

MPTPp

3

*

4

3

3

3

60 days after treatment

*

3

10

60

*

2 1 0 1

3

7

Pole test

3

time to descend (sec)

100

vehicle

3

3

MPTPp

*

*

7

10

*

50

0 1

3

60

Fig. 1. Progressive impairment of olfactory and motor functions induced by chronic MPTPp administration. Retrieval of a buried smelly pellet is used as olfactory test; the beam traversal test and the pole test are used to detect motor impairments. *p < 0.05 as compared with respective vehicle-treated mice, 3p < 0.05 versus 1 and 3 MPTPp administrations; by Newman-Keuls post hoc test.

4. In order to estimate cell loss in the entire SNc volume, stereological counting is usually required. Alternatively, density of dopaminergic neurons in the SNc can be evaluated by confocal laser scanning microscopy (CLSM) analysis of this area, which provides 3-dimensional measures with accuracy

17

MPTPp Model of PD

299

Fig. 2. Upper panels: confocal 3D reconstraction (40×) of dopaminergic neurons in the substantia nigra compacta, double stained for TH (green) and α-synuclein (orange). Lower panels: confocal 3D reconstraction (100×) of TH-positive fibers (green) in the striatum, colocalized with synapsin-I (yellow ). Images are from the substantia nigra and striatum of mice administered with vehicle or with chronic MPTPp.

higher than conventional light microscopy, allowing the assessment of density as well as morphological changes of cells (31, 32). 5. In the striatum, DA and DA metabolites content are dramatically reduced since the first MPTPp administration, as assessed by HPLC measurement (23) (Fig. 3). In addition, at the postsynaptic level, gradual changes in neuropeptides mRNA expression in medium-sized striatal neurons occur, indicating an abnormal postsynaptic neuronal activity (23). Interestingly, these changes show a delayed development if compared to the drop of presynaptic DA content, possibly reflecting the temporarily development of compensatory mechanisms (23). Based on these observations, the chronic MPTPp protocol of administration represents a valid model of progressive PD to investigate the chronic mechanisms and to test neuroprotective strategies for the disease. Behavioral tests, including non-motor and motor tests, are advised as crucial tools to confer face validity to the preclinical model of a motor disorder.

300

A.R. Carta et al.

DOPAC/Dopamine

5

striatal DA content (pg/mg tissue)

6000

4000

4 3 2 1 0

2000 *

*

* *

0 0

1

3

7

10

MPTP administrations

striatal DOPAC content (pg/mg tissue)

3200

2400 * *

*

1600 ^ * 800

0

0

1

3

7

10

MPTP administrations

Fig. 3. Loss of striatal dopamine and DOPAC content from mice chronically treated with MPTPp. *p < 0.05 versus vehicle-treated mice.

2. Materials 2.1. Animals and Drug Treatment

1. Male C57Bl/6 J mice, 3 months old (25–30 g). 2. MPTP-HCl (25 mg/kg i.p.) (Sigma-Aldrich, St. Louis, MO) dissolved in water. 3. Probenecid (100 mg/kg i.p.) dissolved in 5% NaHCO3.

17

2.2. Olfactory Test

MPTPp Model of PD

301

1. Clean plastic cage (24 width, 42 length, 15 height in centimeters). 2. Cheese-smelly pellet.

2.3. Beam Traversal Test

1. Plexiglas Beam: the beam consists of four sections (25 cm each, 1 m total length) of different width, starting at a width of 3.5 cm and gradually narrowing to 1 cm. 2. Mesh grid (1 cm square) of corresponding width to be placed over the beam, leaving a 1 cm space between the grid and the beam surface. 3. Video recording device.

2.4. Pole Test 2.5. Fluorescence Immunohistochemistry and Confocal Microscopy

Vertical pole (diameter 8 mm, height 55). 1. Surgical tools for the transcardial perfusion. 2. Brain removing and slices processing (Fine Science Tools): 22-gauge needles, scissors, scalpel. 3. Vibratome or Cryostat. 4. Multiwell plates and pipettes. 5. Microscope slides (superfrost plus for cryostat) and coverslips (VWR, Radnor, PA). 6. 2× Phosphate-buffer saline (pH 7.4) (PBS). 7. 4% Paraformaldehyde (PFA) in 1× PBS. Heat H2O to 50°C and add 8% PFA under a hood. Filter and store at 4°C. Add 8% PFA to 2× PBS 1:1 ratio. 8. 0.1% Sodium azide in 1× PBS (Sigma-Aldrich). 9. Antifreezing solution: 30% glycerol, 30% ethylene glycol in 1× PBS, 30% sucrose (for cryostat sections).

2.6. High Pressure Liquid Chromatography (HPLC) 2.6.1. Extraction

The process of measuring neurotransmitters and metabolites from biological samples can be divided into two steps, extraction and assessment. 1. Ultrasonic homogenizer. 2. Centrifuge: to separate the analytes from the cell material a centrifuge is required. The samples can be kept in ice at dark and analyzed immediately after centrifugation. Alternatively, brain tissue can be stored in −80°C in a deep freezer until use.

2.6.2. Assessment

1. HPLC pump (e.g., Jasco PU 1580, Great Dunmow, Essex UK). 2. Chromatographic column (e.g., LC −18 DB, 15 cm, 5 μm particle size, Supelco, Milano, Italy, or Simmetry C-8 Waters, Milford, MA).

302

A.R. Carta et al.

3. Injector (e.g., Valco Instruments Co., Inc., Huston, TX, USA) or an autosampler (e.g., ANTEC AS 110 autosampler, Leyden NL). 4. Electrochemical detector (e.g., ESA Coluchem II Chelmsford MA) with its electrode (e.g., ESA 50514 B). 5. Integrator (e.g., Agilent 3396, Santa Clara CA) or a detection software (e.g., Kromatek Dunmow, Essex UK).

3. Methods 3.1. Animals and Drug Treatment

MPTP and probenecid are administered twice a week for 5 weeks, for a total of 10 administrations. Probenecid is given 30 min before MPTP.

3.2. Behavioral Tests

Olfactory function and motor performance are evaluated after 1, 3, 7, and 10 MPTPp injections, and 2 months after treatment discontinuation. Behavioral tests should not detect acute pharmacological actions of MPTP/MPP+, unrelated to neuronal damage, while should reflect the neurodegenerative effect. For this reason, in our studies we performed all behavioral tests on the third day after each time point (23).

3.2.1. Olfactory Test

Mice are food-deprived for 20 h before test. The test is conducted in a clean plastic cage (24 w, 42 l, 15 h cm). A cheese-smelly pellet is buried 1 cm under the bedding in a cage corner, and the mouse is positioned in the center of the cage. Time spent to retrieve the pellet and bite it is measured (23) (Fig. 1).

3.2.2. Beam Traversal Test

The beam traversal test used to measure motor performance has been adapted from the traditional beam-walking tests (33–35) (Fig. 1). The beam is made of Plexiglas and consists of four sections (25 cm each, 1 m total length) of different width, starting at a width of 3.5 cm and gradually narrowing to 1 cm. Mice are trained for 2 consecutive days, and for five trials each day, to traverse the beam from the widest to the narrowest side. On the test day, a mesh grid (1 cm square) of corresponding width is placed over the beam, leaving a 1 cm space between the grid and the beam surface. Mice are videotaped while traversing the grid-surfaced beam for a total of five trials. Videotapes are viewed and rated in slow motion to detect errors. An error consists of a limb (forelimb or hindlimb) slipping through the grid during a forward movement that is visible between the grid and the beam surface. The severity of the error is measured by scoring each limb slip individually (23, 35).

17

MPTPp Model of PD

303

3.2.3. Pole Test

The pole test is used to assess basal ganglia related movement disorders in mice (36–39) (Fig. 1). Mouse is placed head-up on top of a vertical pole (diameter 8 mm, height 55) placed in the cage. When placed on the pole, mouse orients himself downward and descends the length of the pole back into the cage. This test requires motor coordination to turn downward and to climb to the ground. Mice are trained for 2 consecutive days, for five trials each day. On the test day, animals receive five trials, and total time to orient downward and descend the pole is measured with a maximum duration of 120 s.

3.3. Fluorescence Immunohistochemistry and Confocal Microscopy

The procedure can be subdivided into three phases: Phase 1: Fluorescence immunocytochemistry, 1–3 days; Phase 2: CLSM, time required variable; Phase 3: Computer analysis, time required variable.

3.3.1. Immunofluorescence

Mice are anesthetized at selected time points with chloral hydrate (400 mg/kg i.p.) and transcardially perfused with ice-cold 4% paraformaldehyde in 1× PBS. Brains, carefully removed from the skull, are postfixed in 4% PFA/PBS for 2 h, and stored in SA at 4°C until cut. For cryostat sectioning, brains are PBS rinsed (5 × 2 h), and cryoprotected in 30% sucrose for 2–3 days. Sections (50 μm thick) are vibratome or cryostat-cut, and collected in multiwell plates (in PBS) for free-floating procedure. For the SNc, consecutive sections are collected starting at −2.85 mm anterior from bregma to the posterior ending of the area (18 total sections) (40). Every third other section is processed and analyzed for TH immunohistochemistry. Adjacent SNc sections are Nissl-stained, in order to evaluate real cell loss, or processed to visualize different proteins (see below). 1. Rinse in PBS (3 × 15 min). 2. Incubate for 30–60 min in a solution containing 5% normal goat serum (NGS), 0.5% Triton X-100, and 5% Bovine serum albumin in PBS. 3. Incubate with primary antibodies, overnight at 4°C. Process sections for double labeling in various combinations (TH and PSD-95/synapsin I/alpha-synuclein). 4. Incubate in a cocktail-containing rabbit anti-TH antibody (1:400, Sigma-Aldrich Mo, USA) and mouse anti-PSD-95 (1:400, Santa Cruz Biotec. CA, USA), or rabbit anti-synapsin I (1:400, Santa Cruz Biotec. CA, USA), or mouse anti-alphasynuclein (1:400), BD Transduction Laboratories, USA) in PBS (pH 7.4). 5. Rinse in PBS (3 × 15 min). 6. Incubate with secondary antibodies.

304

A.R. Carta et al.

7. To enhance the signal (for small and scattered structures like PSD-95 antigen), incubate in biotinylated anti-mouse (rabbit) IgG (1:400) for 2 h at RT. PBS rinse (3 × 15 min) and incubate in a cocktail containing streptavidin fluorescein conjugate (1:400, Vector, CA, USA) and goat anti-rabbit (mouse) IgG Alexa Fluor 594 (1:400, Invitrogen, CA, USA) for 4 h at RT. 8. For normal signal, incubate in a cocktail containing anti-mouse IgG TRITC conjugate (1:400 Sigma, Mo USA) and anti-rabbit IgG FITC conjugate (1:400 Vector , CA, USA) for 4 h at RT. 9. Rinse in PBS (3 × 15 min). 10. Glass mount and coverslip with vectashield (Vector, CA, USA) mounting medium. 11. For control procedure follow step-by-step the previous procedure without incubation with primary antibodies (step 3, replace the primary antibody with PBS). 3.3.2. Microscopy Analysis

A main problem with conventional microscopy is the out-of-focus light that leads to reduction in image contrast and a decrease in resolution. The resulting blurring effect degrades the image by shadowing important structures of interest, particularly in thick specimens. In CLSM, out-of-focus structures do not contribute to image shaping. By visualizing fluorescence associated with multiple markers, CLSM is extremely valuable for co-localization studies. CLSM scans can be collected in different datasets for computer combination, volume and surface rendering (Fig. 3).

3.3.3. Computer Analysis Via Rendering

The rendering technique is a computer algorithm used to transform serially acquired images into 2D images containing 3D information. Maximum Intensity Projection (MIP), Extended (depth-of) Focus (EdF), Simulated Fluorescence Process (SFP), Surface rendering and Colocalization algorithms are frequently used to display, analyze, and counting. In particular, the MIP technique considers the brightest point (the pixel with maximum intensity value) along the viewing direction. This rendering process is fast and provides an estimation of the amount of fluorescence without 3-D information. On the contrary, the EdF synthesizes the three-dimensional information of the specimen in a projection. The SFP renders a “realistic” view by surface shading, transparency, and various lighting effects. This algorithm simulates how the material would appear as if it was excited and how the emitted light would travel. Surface rendering algorithm, based on triangulated surfaces, is best used for volumetric counts, required in small structures as the substantia nigra, to create shaded solid 3-D objects. This method utilizes information from sequenced image slices, in a topologically consistent way. The rendered surfaces are interactively displayed and analyzed for global structure

17

MPTPp Model of PD

305

properties. Accordingly, automatic measurements of geometrical characteristics, as area, volume, length, etc., can be directly obtained by dedicated softwares. Appropriate spatial resolution (X, Y, and Z) and high image quality are required for consistency and accuracy of the results. In addition, high quality contrast images enable segmentation simply by thresholding, while low contrast images, due to uneven staining or heterogeneous acquisition, require more advanced techniques of image analysis and processing. 3.4. High Pressure Liquid Chromatography (HPLC)

The possibility of detecting biogenic amines such as catecholamines (dopamine (DA) and noradrenaline (NA)) or indolamines (serotonin (5-HT)) and relative metabolites with HPLC coupled with electrochemical detection (ED) has provided a very powerful tool for investigating peripheral and central nervous system (40–42). The essential feature that allows the detection of these molecules is their readiness to be oxidized, in fact oxidation of DA, NE, and 5-HT occurs spontaneously in solution. Nevertheless, if spontaneous oxidation of DA, NA, 5-HT, and metabolites is prevented the quantitative detection of these molecules in a biologic fluid or in a biological extract could be successfully managed by oxidizing them after separation on a chromatographic column. Briefly, the application of an electrical oxidation potential to a carbon-based electrode on which the substance to be oxidized flows, will produce a chemical derivative with the giving up of electrons that will alter the basal current detected, generating a signal. The electronic elaboration of this signal through an electrochemical detector will allow the quantitative detection of oxidable substances because the current alteration is proportional to the concentration of the substance in the sample. 1. Mice are sacrificed by CO2 inhalation one at a time. 2. The brain is rapidly removed and the striatum is dissected on an iced surface prepared by freezing saline in an expanded polystyrene container. 3. The striatum is quickly introduced in microcentrifuge tube previously weighted. Tubes are stored in dry ice until supplementary storage at −80°C or immediate processing (see Note 1). 4. In the latter case the number of sample to be processed depends on the capability of the assaying procedure through the HPLC (see Note 2). 5. Keeping all the test tubes in ice, 250 μl of previously cooled 0.2 M perchloric acid is added in each test tube. 6. Striatal tissue is sonicated (see Note 3) and then centrifuged at 9,000 × g for 15 min at 4°C. 7. Supernatant is filtered (0.6 μm) and diluted 1: 200 (see Note 4).

306

A.R. Carta et al.

8. Twenty microliters are injected into an HPLC apparatus, equipped with a reverse-phase column (LC −18 DB, 15 cm, 5 μm particle size Supelco, Milano, Italy) and coulometric detector (Esa Coulochem II and 5014B electrode) to quantitate DA and DOPAC. Electrodes are set at +150 mV (oxidation) and −200 mV (reduction). 9. The mobile phase (in mM: CH3COONa, 0.23 M; Citric acid, 0.15 M; Na2EDTA, 100 mg/mL; pH 5.5) is pumped by a Jasco PU 1580 pump at 1 mL/min flow rate. The assay sensitivity for DA and DOPAC is 5 and 10 fmol/sample, respectively (see Note 4).

4. Notes 1. Extreme care must be taken to prevent neurotransmitter degradation by heat or light. Degradation of neurotransmitters and metabolites during extraction are the most likely cause for variability in substance values (i.e., pg/mg of fresh tissue) among samples or found by different laboratories in the same animal species. During the entire procedure test tubes must be kept on ice and in dark conditions. 2. A typical HPLC run lasts from 10 to 12 min or 15–20 min. Therefore, the number of samples to be processed must be carefully scheduled, considering that up to 3–4 samples can be processed per hour. 3. The selection of sonication parameters is crucial. We usually use a probe with 2 mm diameter, sonicating each sample 4 × 5 s, with 3 s intervals, at 30% power (130 W). 4. The sample dilution depends on the HPLC system sensibility for DA and metabolites and may vary from 1:50 to 1:200. Considering that an HPLC apparatus can detect as little as 1 pg of DA in a 20 μL sample and the average content of striatal tissue is about 5,000 pg/mg of fresh tissue, it is possible to assess DA and metabolites in each striatum in order to have an internal control for the extraction and detection procedure. 5. The absolute amount of neurotransmitter and metabolite detected can be obtained by using a standard curve reference. However, standards may undergo partial degradation, resulting in the detection of higher amounts of neurotransmitters and metabolites. Keeping them on ice and in the dark will minimize oxidative degradation. Additionally sodium metabisulfite can be added (41). Each standard must be injected in duplicate.

17

MPTPp Model of PD

307

References 1. Hawkes CH, Del Tredici K, Braak H (2007) Parkinson’s disease: a dual-hit hypothesis. Neuropathol Appl Neurobiol 33:599–614 2. Hornykiewicz O, Kish SJ (1987) Biochemical pathophysiology of Parkinson’s disease. Adv Neurol 45:19–34 3. Hirsch EC (2000) Nigrostriatal system plasticity in Parkinson’s disease: effect of dopaminergic denervation and treatment. Ann Neurol 47:S115–S120 4. Nisbet AP, Foster OJ, Kingsbury A, Eve DJ, Daniel SE, Marsden CD, Lees AJ (1995) Preproenkephalin and preprotachykinin messenger RNA expression in normal human basal ganglia and in Parkinson’s disease. Neuroscience 66:361–376 5. Goto S, Hirano A, Matsumoto S (1990) Metenkephalin immunoreactivity in the basal ganglia in Parkinson’s disease and striatonigral degeneration. Neurology 40:1051–1056 6. Barcia C, Fernández Barreiro A, Poza M, Herrero MT (2003) Parkinson’s disease and inflammatory changes. Neurotox Res 5:411–418 7. Hirsch EC, Hunot S (2009) Neuroinflammation in Parkinson’s disease: a target for neuroprotection? Lancet Neurol 8:382–397 8. Imamura K, Hishikawa N, Sawada M, Nagatsu T, Yoshida M, Hashizume Y (2003) Distribution of major histocompatibility complex class II-positive microglia and cytokine profile of Parkinson’s disease brains. Acta Neuropathol 106:518–526 9. Olanow CW (2007) The pathogenesis of cell death in Parkinson’s disease (2007). Mov Disord 22(17):S335–S342 10. Zhou C, Huang Y, Przedborski S (2008) Oxidative stress in Parkinson’s disease: a mechanism of pathogenic and therapeutic significance. Ann N Y Acad Sci 1147:93–104 11. Wakabayashi K, Tanji K, Mori F, Takahashi H (2007) The Lewy body in Parkinson’s disease: molecules implicated in the formation and degradation of alpha-synuclein aggregates. Neuropathology 27:494–506 12. Meredith GE, Sonsalla PK, Chesselet MF (2008) Animal models of Parkinson’s disease progression. Acta Neuropathol 115: 385–398 13. Olanow CW, Kieburtz K, Schapira AH (2008) Why have we failed to achieve neuroprotection in Parkinson’s disease? Ann Neurol 64(Suppl 2):S101–S110 14. Manning-Bog AB, Langston JW (2007) Model fusion, the next phase in developing animal

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

models for Parkinson’s disease. Neurotox Res 11:219–240 Blandini F, Armentero MT, Martignoni E (2008) The 6-hydroxydopamine model: news from the past. Parkinsonism Relat Disord 14(Suppl 2):S124–S129 Chesselet MF (2008) In vivo alpha-synuclein overexpression in rodents: a useful model of Parkinson’s disease? Exp Neurol 209:22–27 Jenner P (2008) Functional models of Parkinson’s disease: a valuable tool in the development of novel therapies. Ann Neurol 64(Suppl 2):S16–S29 Schneider B, Zufferey R, Aebischer P (2008) Viral vectors, animal models and new therapies for Parkinson’s disease. Parkinsonism Relat Disord 14(Suppl 2):S169–S171 Bezard E, Dovero S, Bioulac B, Gross C (1997) Effects of different schedules of MPTP administration on dopaminergic neurodegeneration in mice. Exp Neurol 148:288–292 Fornai F, Schlüter OM, Lenzi P, Gesi M, Ruffoli R, Ferrucci M, Lazzeri G, Busceti CL, Pontarelli F, Battaglia G, Pellegrini A, Nicoletti F, Ruggieri S, Paparelli A, Südhof TC (2005) Parkinson-like syndrome induced by continuous MPTP infusion: convergent roles of the ubiquitin-proteasome system and alpha-synuclein. Proc Natl Acad Sci U S A 102:3413–3418 Smeyne RJ, Jackson-Lewis V (2005) The MPTP model of Parkinson’s disease. Brain Res Mol Brain Res 134:57–66 Yazdani U, German DC, Liang CL, Manzino L, Sonsalla PK, Zeevalk GD (2006) Rat model of Parkinson’s disease: chronic central delivery of 1-methyl-4-phenylpyridinium (MPP+). Exp Neurol 200:172–183 Schintu N, Frau L, Ibba M, Garau A, Carboni E, Carta AR (2009) Progressive dopaminergic degeneration in the chronic MPTPp mouse model of Parkinson’s disease. Neurotox Res 16:127–139 Schmidt N, Ferger B (2001) Neurochemical findings in the MPTP model of Parkinson’s disease. J Neural Transm 108:1263–1282 Heikkila RE, Hess A, Duvoisin RC (1984) Dopaminergic neurotoxicity of 1-methyl-4phenyl-1,2,5,6-tetrahydropyridine in mice. Science 224(4656):1451–1453 Zuddas A, Fascetti F, Corsini GU, Piccardi MP (1994) In brown Norway rats, MPP + is accumulated in the nigrostriatal dopaminergic terminals but it is not neurotoxic: a model of natural resistance to MPTP toxicity. Exp Neurol 127:54–61

308

A.R. Carta et al.

27. Ricaurte GA, Langston JW, Delanney LE, Irwin I, Peroutka SJ, Forno LS (1986) Fate of nigrostriatal neurons in young mature mice given 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine: a neurochemical and morphological reassessment. Brain Res 376:117–124 28. Petroske E, Meredith GE, Callen S, Totterdell S, Lau YS (2001) Mouse model of Parkinsonism: a comparison between subacute MPTP and chronic MPTP/probenecid treatment. Neuroscience 106:589–601 29. Meredith GE, Totterdell S, Petroske E, Santa Cruz K, Callison RC Jr, Lau YS (2002) Lysosomal malfunction accompanies alphasynuclein aggregation in a progressive mouse model of Parkinson’s disease. Brain Res 956:156–165 30. Novikova L, Garris BL, Garris DR, Lau YS (2006) Early signs of neuronal apoptosis in the substantia nigra pars compacta of the progressive neurodegenerative mouse 1-methyl-4phenyl-1,2,3,6-tetrahydropyridine/ probenecid model of Parkinson’s disease. Neuroscience 140:67–76 31. Spiga S, Serra GP, Puddu MC, Foddai M, Diana M (2006) Morphine withdrawalinduced abnormalities in the VTA: confocal laser scanning microscopy. Eur J Neurosci 17:605–612 32. Spiga S, Lintas A, Migliore M, Diana M (2010) Altered architecture and functional consequences of the mesolimbic dopamine system in cannabis dependence. Addict Biol 15:266–276 33. Drucker-Colín R, García-Hernández F (1991) A new motor test sensitive to aging and dopaminergic function. J Neurosci Methods 39:153–161 34. Carter RJ, Lione LA, Humby T, Mangiarini L, Mahal A, Bates GP, Dunnett SB, Morton AJ (1999) Characterization of progressive motor

35.

36.

37.

38.

39.

40.

41.

42.

deficits in mice transgenic for the human Huntington’s disease mutation. J Neurosci 19:3248–3257 Fleming SM, Salcedo J, Fernagut PO, Rockenstein E, Masliah E, Levine MS, Chesselet MF (2004) Early and progressive sensorimotor anomalies in mice overexpressing wild-type human alpha-synuclein. J Neurosci 24:9434–9440 Ogawa N, Hirose Y, Ohara S, Ono T, Watanabe Y (1985) A simple quantitative bradykinesia test in MPTP-treated mice. Res Commun Chem Pathol Pharmacol 50:435–441 Matsuura K, Kabuto H, Makino H, Ogawa N (1997) Pole test is a useful method for evaluating the mouse movement disorder caused by striatal dopamine depletion. J Neurosci Methods 73:45–48 Sedelis M, Hofele K, Auburger GW, Morgan S, Huston JP, Schwarting RK (2000) MPTP susceptibility in the mouse: behavioral, neurochemical, and histological analysis of gender and strain differences. Behav Genet 30:171–182 Fernagut PO, Chalon S, Diguet E, Guilloteau D, Tison F, Jaber M (2003) Motor behaviour deficits and their histopathological and functional correlates in the nigrostriatal system of dopamine transporter knockout mice. Neuroscience 116:1123–1130 Paxinos G, Franklin KBJ (2001) The Mouse Brain in Stereotaxic Coordinates, 2nd edn. Academic, San Diego Marsden CA, Joseph MH (1986) Biogenic amines. In: Kim CK (ed) HPLC of small molecules, a practical approach. IRL Press, Oxford Carboni E (2003) Microdialysis coupled with electrochemical detection: a way to investigate brain monoamine role in freely moving animals. Methods Mol Med 79:415–432

INDEX

A

C

Action potential ............................... 124, 125, 130–132, 235, 261, 264, 265, 270, 289 Acute slice preparation .................................................... 128 Adenosine A2A ................................................................... 96 ADHD. See Attention deficit hyperactivity disorder (ADHD) Animal model .................................................................. 296 Parkinson’s disease (PD) ............................................ 296 β-Arrestin2 ...............................................108–113, 116–120 Attention deficit hyperactivity disorder (ADHD)............................................ 3, 107 Axonal dopamine release ......................................... 243–271

Calcium imaging advantages and disadvantages .................................... 133 dendritic calcium ............................................... 123–135 voltage calcium imaging ............................ 127, 132–133 Calmodulin kinase ............................................................. 44 CaMKII activity .......................................................... 44 Cannabinoids CB1 ............................................................ 96 Carbon-fiber microelectrode .... 245–258, 262, 263, 270, 280 Caveolae ................................................................ 16, 19, 20 Cell culture ................................ 8, 17, 19, 26–27, 29–32, 45, 80–84, 99, 100, 109–110, 147–148, 175, 176 Chimeric mice breeding ............................................................. 182, 195 production ......................................................... 192–195 Chromosomal mapping ........................................... 209–212 zebrafish dopamine receptor genes .................... 209–212 Cloning....................... 99, 150, 158, 181–183, 187, 203–204 dopamine receptor genes ........................................... 182 signaling pathways ....................................................... 62 Confocal microscopy .....................................90–92, 99, 130, 133, 301, 303 Contigs ............................................................................ 202 Crosslinking .................................................... 46–47, 55–59 cell permeable crosslinking .................................... 55–57 protein crosslinking ............................................... 46, 56 reverse crosslinking .................................... 46–47, 57–59 Cyclic AMP (cAMP) .......................... 25, 26, 37–38, 41, 44, 66, 99, 108, 109, 111, 114–118, 120, 142, 149–151, 160, 166, 169–173, 176, 178 whole-cell cAMP assay..............................149–150, 160, 166, 169, 173, 176, 178

B Backpropagation ...................................................... 124, 125 Beam traversal test ............................................298, 301, 302 Binding assays ................................................... 4, 28, 35–37 Bioluminescence resonance energy transfer (BRET) .................................... 95–104, 108 Biotinylation biotinylation of cell surface receptors ..................................................................4 sulfo-NHS-biotin ............................................ 4–6, 8, 10 Bipolar electrical stimulation, 281 Blastocysts harvesting .......................................................... 184, 192 injection ..............................................184–185, 192–195 Brain ...........................................4, 39, 44, 45, 49, 53, 55, 66, 107, 108, 124–126, 128, 129, 133–136, 181, 182, 201, 216, 217, 220, 223, 224, 233, 240, 243–271, 275–292 protein preparation ...................................................... 45 Brain slice forebrain slices ....................................244, 245, 258–261 midbrain slices .................................................. 244, 245, 259, 261 Breeding ...........................................................182, 192, 195 BRET. See Bioluminescence resonance energy transfer (BRET) BS3..................................................................................... 57

D DAPI ..............................................................6, 9, 11, 29, 32 DAT. See Dopamine transporter (DAT) Dendritic excitability.................................................................. 268 spike........................................................................... 124 Diffusion constant ................................................. 64–66, 73 DNA ligation .................................................. 150, 155–156

Nadine Kabbani (ed.), Dopamine: Methods and Protocols, Methods in Molecular Biology, vol. 964, DOI 10.1007/978-1-62703-251-3, © Springer Science+Business Media, LLC 2013

309

DOPAMINE 310 Index Dopamine (DA) binding ....................................................................... 61 binding assays ........................................................35–37 dose-response curve ...................................166–171, 177 extraction .........................................................29, 38–39 [3H]-SCH 23390 labeling .............................. 81, 84, 85, 164, 167 oxidation peak....................................................251, 270 quantification .........................................................38–39 release .............................26, 69, 229, 243–271, 275–292 signaling ................................ 61–74, 202, 215–225, 289 uptake assay ....................................... 231–232, 235, 241 Dopamine (DA) receptor D1-like ..............................15, 25, 95, 108, 142, 151, 181 D2-like .......................................... 15, 95, 108, 181, 205 D2long .........................................................................110 D2Rshort ......................................................................142 interacting protein, DRIP................................44, 52–55 oligomerization ......................................................79–93 Dopamine release axonal.................................................................243–271 somatodendritic .................................................243–271 Dopamine transporter (DAT) .............. 25, 30–32, 229–241, 262, 269, 296, 297 Drosophila ....................................... 208, 209, 213, 215–225 Drosophila melanogaster ......................................208, 216 Dye injection ...........................................................128–129 Dynabeads ......................................45, 50–52, 216, 220, 221

E Electrochemistry .....................................................278–281 ELISA. See Enzyme-linked immunoabsorbant assay (ELISA) Embryonic stem (ES) cells cloning .......................................................182, 190–191 electroporation ...........................................................190 isolation ...............................................................87–188 plating........................................................................189 screening ............................................................182, 191 transfection .......................................................183–184, 187–192 Enriched neuronal cultures ............................................... 29 Enzyme-linked immunoabsorbant assay (ELISA) ....................................................4, 6, 8–11 Exchange protein activated by cAMP (EPAC) sensor ...........................................................114, 120 Expressed sequence tag (EST) ........................ 202, 203, 205 EST database............................................. 202, 203, 205

F Fast-scan cyclic voltammetry (FSCV)....................243–271, 276–278, 280, 281, 284, 289–290

Fluorescence resonance energy transfer (FRET)...................80–82, 86–92, 95–104, 108, 110 Freely moving rat .....................................................286–287 FRET. See Fluorescence resonance energy transfer (FRET) FSCV. See Fast-scan cyclic voltammetry (FSCV) Fusion protein ...............................80, 82–84, 86, 87, 91, 92, 96–100, 103, 104, 220

G GAL4/UAS ...................................................... 216, 217, 220 Genotyping ............................................. 182, 185, 195–197 Green fluorescent protein (GFP)............... 97, 217, 219–223 dopamine cell labeling ...............................................219

H High pressure liquid chromatography (HPLC) ................. 7 dopamine measurements .....................................38, 299 dopamine metabolite measurement ...................305, 306 Homologous recombination ........................... 182, 185, 186, 191, 197, 198

I ICSS. See Intracranial self-stimulation (ICSS) Immunocytochemistry..................... 26, 30–32, 66, 303–305 immunofluorescence ..........................................303–304 Immunoprecipitation ........................... 45–46, 51, 56, 57, 59 co-immunoprecipitation ........................................50–52 In gel digestion ................................................ 46–47, 52, 58 In situ hybridization.........................................................202 Intracranial self-stimulation (ICSS) ................................289 Ion exchange chromatography ...............................37–38, 41

K Kidney cells .................................................................15–23 Knockout (KO) mouse ............................................182, 185 D2LR ................................................................182, 186

L Ligand binding assay competition assay ...................................................84–86 saturation assay ........................................................... 84 Lipid raft immunoblotting to analyze ......................................... 21 preparation with detergent method .......................20–21 preparation with non-detergent method................19–20

M Macromolecular complex.................................................. 96 Magnetic bead cell sorting ................................. 216, 218–220, 222, 223 MAP kinase.....................................................................239

DOPAMINE 311 Index Mass spectrometry liquid chromatography electrospray ionization (LC-ESI) ........................................44, 52 matrix assisted laser desorption ionization.................. 44 maximum intensity projection ...................................304 tandem mass spectrometry.......................................... 54 Maximum parsimony...............................................207–209 MEM. See Minimal essential media (MEM) Membrane preparation crude membrane from HEK293 cells ................162, 166 preparation from cultured striatal cells........................ 47 Mesencephalic primary cultures .............. 230–231, 233–234 3-Methoxytyramine (3-MT) ..................................108, 110, 111, 114, 115 Methyl-β-cyclodextrin (βCD) ..............................17, 19, 22 1-Methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP).......................................................295–306 Microdomain ....................................... 15, 17, 19–21, 23, 89 Minimal essential media (MEM) ................... 109, 113, 147, 148, 159, 160, 166, 169, 173, 175 MPTP. See 1-Methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) 3-MT. See 3-Methoxytyramine (3-MT) Müller glia ....................................................... 26, 29–30, 37 Mutagenesis.......................................................82, 141–179 mutant receptor .................................................153–155 Mutant cassettes ..............................................................155

N Neuronal-glial cultures ..................................................... 29 Neuroprotection ............................................................... 26 Neurospheres ...............................................................26, 30 retinal culture .............................................................. 30 Northern blot ..................................................188, 196–197

O Olfactory test ................................................... 298, 301, 302 Oligonucleotides ......................... 27, 28, 35, 40, 80, 82, 143, 145, 146, 175, 196, 199, 202 Optical imaging .......................................................126–127 Ortholog dopamine receptor genes in zebrafish .......... 82, 201, 209

P Parkinson’s disease (PD) ..........................3, 39, 61, 107, 201, 202, 245, 275, 295–306 Patch electrode recording ........................................128–129 PCR. See Polymerase chain reaction (PCR) Phasic dopamine firing ..........................................................................275 signal .........................................................................130 β-Phenylethylamine (β-PEA) ........................ 108, 110, 111, 114, 115, 118

Phylogenetic analysis ...............................................206–209 Plate reader .................................................. 6, 114, 116–118 BRET settings ...................................................116–118 Pole test ........................................................... 298, 301, 303 Polymerase chain reaction (PCR) ................... 27, 28, 34–35, 40, 82, 92, 99, 143–146, 150, 152–154, 175, 185, 188, 196–197, 204, 205, 209, 213, 231, 233, 234, 237–239, 241, 289 Preclinical studies ............................................................296 Probenecid ...............................................................295–306 Proteomics protein-protein interaction ......................................... 57 sample preparations .................................................... 45 trypsin digestion ......................................................... 54

R Radioligand binding Bmax ............................................................................. 84 saturation studies .......................................................160 Rate constant .......................................................63–64, 262 Receptor dopamine receptor oligomerization .......................79–93 heteromer ............................................................95–104 Renilla Luciferase (Rluc) .................................................111 Restriction enzyme ..........................154, 157, 159, 173, 183, 186, 191, 196, 1447 Retina retinal cultures .................................................29–30, 37 RNA extraction .......................................................... 39 Reverse transcriptase-PCR (RT-PCR) D1A and D1B receptors .............................................33–35 dopamine receptor .................................................33–35 dopamine transporter (DAT) ....................................237 quantitative RT-PCR ................................ 233, 234, 237 RNA isolation ......................................... 218–220, 222–223

S Schizophrenia (SZ) ................3, 85, 107, 201, 202, 245, 275 Scintillation ........................... 36, 38, 41, 81, 84, 86, 93, 149, 162, 166, 169, 170, 178, 232, 235 Sequential BRET-FRET (SRET) saturation curves ........................................ 101, 103, 104 SRET detection .................................................101–103 Sequential chromatography .............................151, 170–172 Site-directed mutagenesis ......................... 82, 143, 145, 147, 149, 151–155, 157, 159, 161, 163, 165, 167, 169, 171, 173, 175, 177, 179 Striatum..................... 41, 45, 49, 61, 79, 188, 196, 243–245, 249, 259–261, 263–268, 296, 299, 305, 306 Substantia nigra .......... 61, 202, 243, 244, 260, 284, 299, 304 substantia nigra pars compacta .................... 61, 243, 244 Sucrose gradient ..........................................................16–23 sucrose gradient centrifugation ........................17–19, 22

DOPAMINE 312 Index T

V

TAAR1....................................................................107–121 Targeting vector...................................... 182–183, 185–188, 190, 196 Third intracellular loop ........................... 143, 182, 186, 206 Toxicity ....................................................................120, 297 MPTP toxicity ..........................................................297 Trace amines .................................................... 108, 110, 111 Transfection embryonic stem cell ..........................................183–184, 187–192 transient transfection ..................... 83, 91, 100–101, 103 Transformation .......................................... 92, 156, 174, 296 Transgene ................................................ 195, 217, 219, 220 Transmembrane (TM) domain transmembrane region ...............................................142 Transporter ........................................... 25, 39, 44, 229–241, 262, 296, 297 Tyrosine hydroxylase (TH) ............................. 25, 26, 30–33, 217, 232, 238–240, 262, 278, 288, 297, 299, 303

Ventral tegmental area (VTA) ........................ 108, 243–245, 262, 264, 267, 268, 270, 284, 290 Voltage calcium imaging..................................127, 132–133 Voltammetry ....................................................243–271, 276 VTA. See Ventral tegmental area (VTA)

W Western blot dopamine markers .................................................30–31 dopamine transporter ................................................238 lipid raft proteins ........................................................ 18

Z Zebrafish chromosomal mapping ......................................209–212 phylogenetic analysis .................................................208