Synaptic Vesicles: Methods and Protocols (Methods in Molecular Biology, 2417) 1071619152, 9781071619155

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Methods in Molecular Biology 2417

Jana Dahlmanns Marc Dahlmanns Editors

Synaptic Vesicles Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Synaptic Vesicles Methods and Protocols

Edited by

Jana Dahlmanns Department of Psychiatry and Psychotherapy, Universit€atsklinikum Erlangen, Erlangen, Bayern, Germany

Marc Dahlmanns Institute for Physiology and Pathophysiology, University of Erlangen-Nuremberg, Erlangen, Germany

Editors Jana Dahlmanns Department of Psychiatry and Psychotherapy Universit€atsklinikum Erlangen Erlangen, Bayern, Germany

Marc Dahlmanns Institute for Physiology and Pathophysiology University of Erlangen-Nuremberg Erlangen, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1915-5 ISBN 978-1-0716-1916-2 (eBook) https://doi.org/10.1007/978-1-0716-1916-2 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022 This work is subject to copyright. All rights are solely and exclusively licensed 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. 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. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Neuroscience has become an ever-growing and fascinating discipline. The entire field of neuroscience centers around neurons—a cell type mainly characterized by its ability to form chemical synapses. An integral part of the presynaptic terminal is synaptic vesicles: small (around 40 nm in diameter) acidic vesicles that are filled with neurotransmitter molecules, fuse to the presynaptic membrane, release their transmitter content, and are reformed from the membrane to be used in a recycling cycle. These vesicles are of interest to a plethora of scientists, with a variety of research questions, usually focusing on one specific step in their function. The diverse life cycle of synaptic vesicles, though, offers a rich working surface for different neuroscientific research questions, and an equally wide range of methodological approaches is used to investigate synaptic vesicles. This book is intended to provide an overview of techniques to investigate synaptic vesicles at many different stages: From different techniques to detect RNA, over the isolation of single synaptosomes and single synaptic vesicles for protein content investigation, over imaging and quantifying the transport and loading of neurotransmitter and monitoring vesicular pH, over imaging and quantifying the membrane fusion process and release of neurotransmitter, to the dynamics of endocytosis and membrane reuptake. With this collection, we hope to enable scientists to broaden their view of and technical approach to synaptic vesicles to gain a more comprehensive view of the tightly regulated and fascinating chemical synapse dynamics. This whole book series specializes in providing hands-on protocols that scientists can easily establish in their home labs. We hope that with this issue, too, we provide you with simple—yet state-of-the-art—methods that are packed with all the little details and tricks you need to follow, and we wish you all best of luck with your experiments. Erlangen, Bayern, Germany

Jana Dahlmanns Marc Dahlmanns

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

1 Real-Time Quantitative PCR and Fluorescence In Situ Hybridization for Subcellular Localization of miRNAs in Neurons . . . . . . . . . . . . . . . . . . . . . . . . . Antonis Tatarakis and Danesh Moazed 2 Live-Imaging of Axonal Cargoes in Drosophila Brain Explants Using Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caroline Medioni, Anne Ephrussi, and Florence Besse 3 Whole Endosome Recording of Vesicular Neurotransmitter Transporter Currents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roger Chang and Robert H. Edwards 4 Quantitative Analysis of Presynaptic Vesicle Luminal pH in Cultured Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yoshihiro Egashira, Shutaro Katsurabayashi, and Shigeo Takamori 5 Investigation of Synaptic Vesicle Proteins in Rat Brain Tissue Using Real-Time qPCR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Betina Elfving 6 Mass Synaptometry: Applying Mass Cytometry to Single Synapse Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chandresh R. Gajera, Rosemary Fernandez, Nadia Postupna, Kathleen S. Montine, C. Dirk Keene, Sean C. Bendall, and Thomas J. Montine 7 A Guide to Analysis of Relative Synaptic Protein Abundance by Quantitative Fluorescent Western Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ ller Heidi K. Mu 8 Synaptosomes and Metamodulation of Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . Anna Pittaluga and Mario Marchi 9 A Novel Method to Monitor GABA Loading into Synaptic Vesicles by Combining Patch Pipette Perfusion and Intracellular, Caged-GABA Photolysis in Brain Slice Preparations . . . . . . . . . . . . . . . . . . . . . . . . . Manami Yamashita and Tetsuya Hori 10 Rapid Isolation of Functional Synaptic Vesicles from Tissues Through Cryogrinding, Ultracentrifugation, and Size Exclusion Chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Huinan Li 11 Isolation of Synaptic Vesicles from Mammalian Brain. . . . . . . . . . . . . . . . . . . . . . . . Marcelo Ganzella, Momchil Ninov, Dietmar Riedel, and Reinhard Jahn 12 Building and Using a Two-Photon Fluorescence Cross-Correlation Spectroscopy Setup Including Fluorescence Lifetime Analysis . . . . . . . . . . . . . . . . Tobias Grothe and Peter J. Walla

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Fluorescence Lifetime and Cross-correlation Spectroscopy for Observing Membrane Fusion of Liposome Models Containing Synaptic Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tobias Grothe and Peter J. Walla Synaptic Vesicle Pool Monitoring with Synapto-pHluorin. . . . . . . . . . . . . . . . . . . . Marc Dahlmanns and Jana Katharina Dahlmanns Imaging Neuropeptide Release at Drosophila Neuromuscular Junction with a Genetically Engineered Neuropeptide Release Reporter . . . . . . . . . . . . . . . Yifu Han and Keke Ding Imaging Synaptic Glutamate Release with Two-Photon Microscopy in Organotypic Slice Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ rst and Thomas G. Oertner Ce´line D. Du Dynole 34-2 and Acrylo-Dyn 2-30, Novel Dynamin GTPase Chemical Biology Probes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jennifer R. Baker, Nicholas S. O’Brien, Kate L. Prichard, Phillip J. Robinson, Adam McCluskey, and Cecilia C. Russell Synthesis of Phthaladyn-29 and Naphthalimide-10, GTP Site Directed Dynamin GTPase Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cecilia C. Russell, Kate L. Prichard, Nicholas S. O’Brien, Adam McCluskey, Phillip J. Robinson, and Jennifer R. Baker

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors JENNIFER R. BAKER • Chemistry, School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW, Australia SEAN C. BENDALL • Department of Pathology, Stanford University Medical Center, Stanford, CA, USA FLORENCE BESSE • Universite´ Coˆte d’Azur, CNRS, Inserm, iBV, Nice, France ROGER CHANG • Department of Physiology, UCSF School of Medicine, San Francisco, CA, USA; Department of Neurology, UCSF School of Medicine, San Francisco, CA, USA; Graduate Program in Biomedical Sciences, UCSF School of Medicine, San Francisco, CA, USA JANA KATHARINA DAHLMANNS • Department of Psychiatry and Psychotherapy, University Hospital Erlangen, Erlangen, Germany MARC DAHLMANNS • Institute for Physiology and Pathophysiology, University of Erlangen-Nuremberg, Erlangen, Germany KEKE DING • Division of Biology and Biological Engineering, California Institute of Technology, Pasadena, CA, USA CE´LINE D. DU¨RST • Institute for Synaptic Physiology, Center for Molecular Neurobiology Hamburg, Hamburg, Germany; Department of Basic Neurosciences, University Medical Center, Geneva, Switzerland ROBERT H. EDWARDS • Department of Physiology, UCSF School of Medicine, San Francisco, CA, USA; Department of Neurology, UCSF School of Medicine, San Francisco, CA, USA; Graduate Program in Biomedical Sciences, UCSF School of Medicine, San Francisco, CA, USA; Kavli Institute for Fundamental Neuroscience, UCSF School of Medicine, San Francisco, CA, USA; Weill Institute for Neurosciences, UCSF School of Medicine, San Francisco, CA, USA YOSHIHIRO EGASHIRA • Laboratory of Neural Membrane Biology, Graduate School of Brain Science, Doshisha University, Kyoto, Japan; Department of Physiology, Faculty of Medicine, Osaka Medical and Pharmaceutical University, Osaka, Japan BETINA ELFVING • Translational Neuropsychiatry Unit, Department of Clinical Medicine, Aarhus University, Aarhus C, Denmark ANNE EPHRUSSI • European Molecular Biology Laboratory (EMBL), Heidelberg, Germany ROSEMARY FERNANDEZ • Department of Pathology, Stanford University Medical Center, Stanford, CA, USA CHANDRESH R. GAJERA • Department of Pathology, Stanford University Medical Center, Stanford, CA, USA MARCELO GANZELLA • Laboratory of Neurobiology, Max-Planck-Institute for Biophysical Chemistry, Go¨ttingen, Germany TOBIAS GROTHE • Laboratory of Neurobiology, Max-Planck-Institute for Biophysical Chemistry, Go¨ttingen, Germany; Department of Biophysical Chemistry, Institute for Physical and Theoretical Chemistry, TU Braunschweig, Braunschweig, Germany YIFU HAN • Department of Neurobiology, University of Southern California, Los Angeles, CA, USA; Neuroscience Graduate Program, University of Southern California, Los Angeles, CA, USA

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TETSUYA HORI • Cellular and Molecular Synaptic Function Unit, Okinawa Institute of Science and Technology Graduate University, Okinawa, Japan REINHARD JAHN • Laboratory of Neurobiology, Max-Planck-Institute for Biophysical Chemistry, Go¨ttingen, Germany; Georg-August University, Go¨ttingen, Germany SHUTARO KATSURABAYASHI • Department of Neuropharmacology, Faculty of Pharmaceutical Sciences, Fukuoka University, Fukuoka, Japan C. DIRK KEENE • Department of Pathology, University of Washington, Seattle, WA, USA HUINAN LI • Department of Biochemistry and Biophysics, University of California San Francisco, San Francisco, CA, USA; Laboratory for Genomics Research, University of California San Francisco, San Francisco, CA, USA MARIO MARCHI • Department of Pharmacy, School of medical and Pharmaceutical Sciences, and CEBR, University of Genova, Genova, Italy ADAM MCCLUSKEY • Chemistry, School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW, Australia CAROLINE MEDIONI • Universite´ Coˆte d’Azur, CNRS, Inserm, iBV, Nice, France DANESH MOAZED • Department of Cell Biology, and Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA KATHLEEN S. MONTINE • Department of Pathology, Stanford University Medical Center, Stanford, CA, USA THOMAS J. MONTINE • Department of Pathology, Stanford University Medical Center, Stanford, CA, USA HEIDI K. MU¨LLER • Translational Neuropsychiatry Unit, Department of Clinical Medicine, Aarhus University, Aarhus C, Denmark MOMCHIL NINOV • Laboratory of Neurobiology, Max-Planck-Institute for Biophysical Chemistry, Go¨ttingen, Germany; Bioanalytical Mass Spectrometry, Max-Planck-Institute for Biophysical Chemistry, Go¨ttingen, Germany NICHOLAS S. O’BRIEN • Chemistry, School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW, Australia THOMAS G. OERTNER • Institute for Synaptic Physiology, Center for Molecular Neurobiology Hamburg, Hamburg, Germany ANNA PITTALUGA • Department of Pharmacy, School of medical and Pharmaceutical Sciences, and CEBR, University of Genova, Genova, Italy; IRCCS San Martino Hospital, Genova, Italy NADIA POSTUPNA • Department of Pathology, University of Washington, Seattle, WA, USA KATE L. PRICHARD • Chemistry, School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW, Australia DIETMAR RIEDEL • Facility for Transmission Electron Microscopy, Max Planck Institute for Biophysical Chemistry, Go¨ttingen, Germany PHILLIP J. ROBINSON • Cell Signaling Unit, Children’s Medical Research Institute, The University of Sydney, Sydney, NSW, Australia CECILIA C. RUSSELL • Chemistry, School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW, Australia SHIGEO TAKAMORI • Laboratory of Neural Membrane Biology, Graduate School of Brain Science, Doshisha University, Kyoto, Japan ANTONIS TATARAKIS • Department of Cell Biology, and Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA

Contributors

PETER J. WALLA • Laboratory of Neurobiology, Max-Planck-Institute for Biophysical Chemistry, Go¨ttingen, Germany; Department of Biophysical Chemistry, Institute for Physical and Theoretical Chemistry, TU Braunschweig, Braunschweig, Germany MANAMI YAMASHITA • Department of Physiology, Faculty of Medicine, Osaka Medical and Pharmaceutical University, Osaka, Japan

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Chapter 1 Real-Time Quantitative PCR and Fluorescence In Situ Hybridization for Subcellular Localization of miRNAs in Neurons Antonis Tatarakis and Danesh Moazed Abstract Neuronal miRNAs play major roles in regulation of synaptic development and plasticity. The small size of miRNAs and, in some cases, their low level of expression make their quantification and detection challenging. Here, we outline methods to quantify steady state levels of miRNAs in neurons and the brain by using real-time quantitative PCR (RT-qPCR) and to determine miRNA subcellular localization in primary neurons by a sensitive fluorescence in situ hybridization (FISH) method. Key words miRNAs, Neurons, Brain, Expression profile, Isolation, miRNA RT-qPCR assays, miRNA fluorescence in situ hybridization

1

Introduction MicroRNAs (miRNAs) have emerged as a major class of regulators that act at the post-transcriptional level to control the expression of numerous target genes [1]. Neuronal miRNAs play major roles in regulation of synaptic development and plasticity and have been also identified as components of regulatory pathways that modulate memory formation [2, 3]. An attractive model for regulation of activity-related synaptic plasticity describes miRNAs as modulators of gene expression that are capable of both positively and negatively influencing synaptic growth and connections. For example, a group of experience dependent miRNAs were recently identified that target the vesicle exocytosis pathway and are upregulated in the hippocampus of adult rats following contextual fear conditioning and in primary neurons following neuronal activation [4]. Neurons exhibit a high degree of spatial compartmentalization and a rapid and dynamic signaling by processing information in a precise and spatially restricted manner [5]. The signaling that occurs in axons and dendrites necessitates tight control of their

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Antonis Tatarakis and Danesh Moazed

local proteomes. Local translation of mRNAs into protein is one mechanism that neurons use to respond rapidly to intrinsic or extrinsic cues. There is evidence suggesting that miRNAs contribute in the local regulation of mRNAs. For example, miR-134 modulates synapse morphology by inhibiting the expression of LimK1 protein kinase in hippocampal neurons [6]. Thus, understanding the biological roles of miRNAs and miRNA-associated gene regulatory networks requires quantitative and spatio-temporal analyses of miRNA expression in neurons and the brain at the whole cell and subcellular levels. MiRNAs small size and in some cases their low level of expression make their quantification and detection challenging. Here, we outline methods to quantify miRNA expression alterations in neurons and the brain and to determine miRNA subcellular localization in primary neurons by a sensitive fluorescence in situ hybridization (FISH) protocol. The procedures herein include isolation of total RNA, enrichment for the small RNA fraction from primary neurons or brain tissue, reverse transcription of miRNAs, real-time quantitative PCR (RT-qPCR), analysis of miRNA RT-qPCR, and miRNA fluorescence in situ hybridization (FISH).

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Materials

2.1 Isolation of Total RNA from Cultured Neurons or Brain Tissue

1. Trizol, store at 4
C. Trizol should be opened in a fume hood. 2. Chloroform. Chloroform is harmful and should be opened in a fume hood. 3. Isopropanol. 4. Glycogen, aqueous solution (20 mg/mL). Store at 20
C. 5. 75% Ethanol: 75 mL of 100% ethanol to 25 mL of RNase-free water. 6. RNase-free water. 7. Kontes glass dounce tissue grinders.

2.2 Isolation of microRNA Fraction from Cultured Neurons or Brain Tissue

1. miRVana miRNA isolation Kit (Ambion).

2.3 MicroRNA Reverse Transcription

1. TaqMan miRNA reverse transcription kit: 100 mM dNTPs, MultiScribe reverse transcriptase 50 U/μL, 10 RT buffer and RNase inhibitor 20 U/μL (Applied Biosystems), store at 20
C.

2. RNase-free water. 3. Glass dounce tissue grinders (Kontes).

2. TaqMan RT primers, specific for individual miRNAs, from TaqMan miRNA assays (Applied Biosystems), store at 20
C.

miRNA RT-qPCR and FISH in Neurons

3

3. Thermo Scientific™ ABgene™ Thermo-Fast™ 96-Well SemiSkirted PCR Plates. 4. Thermocycler. 2.4 Quantitative Real-Time PCR (qRT-PCR) Profiling of microRNAs

1. TaqMan Universal PCR master mix, no amperase UNG (Applied Biosystems), store at 4
C. 2. Nuclease-free water. 3. TaqMan miRNA assays available for each specific miRNA to be analyzed (Applied Biosystems), light sensitive, store at 20
C. 4. MicroAmp fast optical 96-well reaction plate with barcode 0.2 mL (Applied Biosystems). 5. MicroAmp optical adhesive film (Applied Biosystems). 6. MicroAmp Adhesive Film Applicator (Applied Biosystems). 7. Real-time PCR instrument.

2.5 MiRNA Fluorescence In Situ Hybridization

1. 4% (wt/vol) paraformaldehyde (PFA) in PBS, pH 7.6 (see Note 1). 2. DEPC-PBS. 3. 20x SSC solution: 3 M NaCl, 300 mM tri-Sodium citrate dihydrate, pH 7.0 (see Note 2). 4. Hybridization Mix: 50% (vol/vol) formamide, 5 SSC, 500 mg/mL yeast tRNA (Invitrogen), 1 Denhardt’s solution (Sigma), DEPC-treated water (see Note 3). 5. Acetylation solution: Add 500 mL of 6 N HCl and 670 mL of triethanolamine to 48.5 mL DEPC-treated water, mix and, just before use, add 300 mL acetic anhydride. 6. TN buffer: 0.1 M Tris–HCl, pH 7.5, 0.15 M NaCl. Autoclave and store at room temperature (RT). 7. TNT buffer: 0.1 M Tris–HCl, pH 7.5, 0.15 M NaCl, 0.3% (vol/vol) TritonX-100. 8. 3% (vol/vol) H2O2 (see Note 4). 9. Blocking buffer: 0.1 M Tris–HCl, pH 7.5, 0.15 M NaCl, 0.5% (wt/vol) blocking reagent, 0.5% (wt/vol) BSA. Aliquot and store aliquots at –20
C. 10. Blocking reagent (Roche Applied Sciences). 11. miRCURY LNA™ Detection probe, 250 pmol, 5‘-DIG and 3‘-DIG labeled LNA-oligonucleotide probes (Exiqon-Qiagen) (see Note 5). 12. Anti-Digoxigenin-POD (poly), Fab fragments, sheep (Roche) (see Note 6). 13. TSA PLUS FITC-fluorescein isothiocyanate (Perkin Elmer). Tyramide amplification solution: Dilute FITC or other

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Antonis Tatarakis and Danesh Moazed

relevant fluorochrome-conjugated tyramide 1:100 in the amplification buffer supplied with the kit and use immediately (see Note 7). 14. 40 ,6-diamidino-2-phenylindole (DAPI). 15. Glass coverslips no. 1.5 12 mm (Electron Microscopy Sciences). 16. Parafilm. 17. Humidified incubation chamber (see Note 8). 18. Pair of tweezers. 19. Hybridization oven.

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Methods

3.1 MicroRNA Expression Analysis

3.1.1 Isolation of Total RNA from Cultured Neurons or Brain Tissue

The protocol described below has been used successfully to quantify miRNA levels in both primary cortical and hippocampal neurons as well as in brain tissue (cortex, hippocampus, cerebellum) [4, 6–8]. Cultures of dissociated primary neurons from embryonic day 17 (E17) Swiss Webster wild-type mice (Charles River laboratories) are prepared as described [9]. They are maintained in Neurobasal medium supplemented with B27 and N2 supplements (Invitrogen), penicillin-streptomycin (50 μg/mL penicillin and 50 U/mL streptomycin, Invitrogen), and Glutamine (1 mM, Invitrogen). Neurons are plated at a density of up to 100,000–150,000 cells/cm2 on poly-L-lysine coated multi-well dishes. Three days after plating (DIV3) 5 μM final concentration of cytosine-b-Darabinofuranoside is added into the cultures to inhibit glial cell proliferation and the cultures are fed every 3 days from there on with one-third of the medium to remain in each well. 1. Add 0.5 mL of Trizol or equivalent reagent to 2.5  105 neurons that have been previously pelleted and snap frozen. Pipette up and down about 10–15 times to lyse neurons and allow to stand at room temperature (RT) for 5 min. For brain tissue, use 10x volumes of Trizol to the volume of the brain tissue (weight the tissue and use 1 mL of Trizol for 50–100 mg of tissue). Homogenize the tissue by using a 2 mL glass douncer until no tissue remains can be seen, transfer in a tube of the appropriate volume (1.5 mL Eppendorf tubes for cells) and allow to stand at RT for 5 min (see Note 9). 2. Add 0.2 mL of chloroform per 1 mL of Trizol (0.1 mL for the volume of Trizol used in step 1). Shake vigorously for 15 s and allow to stand at RT for 3 min. 3. Centrifuge at 12,000  g for 15 min at 4
C.

miRNA RT-qPCR and FISH in Neurons

5

4. Transfer the upper aqueous phase containing the RNA to a new Eppendorf tube and add 0.5 mL of ice-cold isopropanol per mL of Trizol used in step 1 (see Note 10). 5. Add 20 μg of glycogen (1 μL of 20 mg/mL stock) (see Note 11). 6. Mix samples by inverting and incubate overnight at 20
C. 7. Centrifuge at 12,000  g for 30 min at 4
C to pellet precipitated RNA. 8. Remove supernatant without disturbing the pellet. 9. Wash pellet with 1 mL of 75% ethanol. Vortex sample briefly and centrifuge at 8000  g for 5 min at 4
C. 10. Remove supernatant and repeat washing as in step 8. 11. Remove supernatant and allow RNA pellet to air-dry for 5–10 min (see Note 12). 12. Add an appropriate volume of RNase-free water, mix by repeat pipetting, allow to stand at room temperature for 5–10 min for the RNA pellet to dissolve (see Note 13). 13. Assess RNA quality and quantity using NanoDrop and/or Qubit (see Note 14). 14. Store at 80
C until required (see Fig. 1 and Note 15). 3.1.2 Isolation of microRNA Fraction from Cultured Neurons or Brain Tissue

For less abundant miRNAs, enrichment of the small RNA fraction of neurons or brain tissue will give better results with regard to the quantification of miRNA levels. Below we outline a protocol to enrich for small RNA ( 1 Gigaohm and access resistance Ra < 100 Megaohm. Capacitance is determined by the endosome size itself. 14. Perfuse with the first recording solution for 30 s before recording by opening the first recording chamber solution and turning on the vacuum (Fig. 7b). 15. For each subsequent (cytoplasmic/bath) recording solution, cycle the next recording solution for 30–60 s before recording again (Fig. 7b).

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Notes 1. For both the bath (cytosolic) and pipette (luminal) solution, filter sterile. While osmolarity can be checked, this is not absolutely necessary. To evaluate the role of different substrates, substitute concentrations of NMDG gluconate/Cl with equimolar concentration of desired substrate to maintain osmolarity. To maintain pH, use either 20 mM HEPES (for pH 7.2) or 20 mM MES (for pH 5.0) as buffers. 2. We describe a schedule that produces a substantial number of transfected cells distributed over several days to allow time for recording. 3. For continued high expression of transport protein, discard after ~10 passages and thaw a new vial of HEK293T cells. 4. Smaller coverslips generally do not have enough space for a sufficient number of ideal, transfected cells with wellpositioned endosomes.

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7. Schenck S, Wojcik SM, Brose N et al (2009) A chloride conductance in VGLUT1 underlies maximal glutamate loading into synaptic vesicles. Nat Neurosci 12(2):156–162. https:// doi.org/10.1038/nn.2248 8. Kirichok Y, Krapivinsky G, Clapham DE (2004) The mitochondrial calcium uniporter is a highly selective ion channel. Nature 427(6972):360–364. https://doi.org/10. 1038/nature02246 9. Garg V, Kirichok YY (2019) Patch-clamp analysis of the mitochondrial calcium uniporter. In: Raffaello A, Vecellio Reane D (eds) Calcium signalling: methods and protocols. Springer New York, New York, NY, pp 75–86. https:// doi.org/10.1007/978-1-4939-9018-4_7 10. Saito M, Hanson PI, Schlesinger P (2007) Luminal chloride-dependent activation of endosome calcium channels: patch clamp study of enlarged endosomes. J Biol Chem 282(37):27327–27333. https://doi.org/10. 1074/jbc.M702557200 11. Chen CC, Cang C, Fenske S et al (2017) Patch-clamp technique to characterize ion channels in enlarged individual endolysosomes. Nat Protoc 12(8):1639–1658. https://doi. org/10.1038/nprot.2017.036 12. Samie M, Wang X, Zhang X et al (2013) A TRP channel in the lysosome regulates large particle phagocytosis via focal exocytosis. Dev Cell 26(5):511–524. https://doi.org/10.1016/j. devcel.2013.08.003 13. Wang X, Zhang X, Dong XP et al (2012) TPC proteins are phosphoinositide- activated sodium-selective ion channels in endosomes

and lysosomes. Cell 151(2):372–383. https:// doi.org/10.1016/j.cell.2012.08.036 14. Miao Y, Li G, Zhang X et al (2015) A TRP Channel senses lysosome neutralization by pathogens to trigger their expulsion. Cell 161(6):1306–1319. https://doi.org/10. 1016/j.cell.2015.05.009 15. Cang C, Bekele B, Ren D (2014) The voltagegated sodium channel TPC1 confers endolysosomal excitability. Nat Chem Biol 10(6):463–469. https://doi.org/10.1038/ nchembio.1522 16. Cang C, Zhou Y, Navarro B et al (2013) mTOR regulates lysosomal ATP-sensitive two-pore Na(+) channels to adapt to metabolic state. Cell 152(4):778–790. https://doi.org/ 10.1016/j.cell.2013.01.023 17. Barbieri MA, Li G, Mayorga LS et al (1996) Characterization of Rab5:Q79L-stimulated endosome fusion. Arch Biochem Biophys 326(1):64–72. https://doi.org/10.1006/ abbi.1996.0047 18. Stenmark H, Parton RG, Steele-Mortimer O et al (1994) Inhibition of rab5 GTPase activity stimulates membrane fusion in endocytosis. EMBO J 13(6):1287–1296 19. Lu Y, Dong S, Hao B et al (2014) Vacuolin-1 potently and reversibly inhibits autophagosome-lysosome fusion by activating RAB5A. Autophagy 10(11):1895–1905. https://doi.org/10.4161/auto.32200 20. Chang R, Eriksen J, Edwards RH (2018) The dual role of chloride in synaptic vesicle glutamate transport. eLife 7:e34896. https://doi. org/10.7554/eLife.34896

Chapter 4 Quantitative Analysis of Presynaptic Vesicle Luminal pH in Cultured Neurons Yoshihiro Egashira, Shutaro Katsurabayashi, and Shigeo Takamori Abstract Newly generated synaptic vesicles (SVs) are re-acidified by the activity of the vacuolar-type H+-ATPases. Since H+ gradient across SV membrane drives neurotransmitter uptake into SVs, precise measurements of steady-state vesicular pH and dynamics of re-acidification process will provide important information concerning the H+-driven neurotransmitter uptake. Indeed, we recently demonstrated distinct features of steady state and dynamics of vesicular pH between glutamatergic vesicles and GABAergic vesicles in cultured hippocampal neurons. In this article, we focus on an experimental protocol and setup required to determine steady-state luminal pH of SVs in living neurons. This protocol is composed of efficient expression of a pH-sensitive fluorescent protein in the lumen of SVs in cultured neurons, and recordings of its fluorescence changes under a conventional fluorescent microscope during local applications of acidic buffer and ionophores-containing solution at a given pH. The method described here can be easily applied for measuring luminal pH of different types of secretory organelles and other acidic organelles such as lysosomes and endosomes in cultured cell preparations. Key words Vesicular pH, pHluorin, mOrange2, Synaptic vesicle, Intracellular organelle

1

Introduction Intracellular organelles have characteristic luminal pH suited to their biochemical functions [1]. Most of the organelles, such as secretory vesicles and lysosomes, maintain acidic interiors by the action of an electrogenic proton pump, the V-ATPase. Dysregulation of luminal pH, e.g., lysosomal pH, is thought to cause severe disorders generally referred to as lysosomal storage diseases [2]. In case of synaptic vesicles (SVs), which utilize the proton electrochemical gradient to concentrate neurotransmitter molecules [3], vesicular pH has been estimated to be ~5.7 [4, 5] in cultured hippocampal neurons. However, the excitatory glutamatergic neurons predominated in most of the preparations used in this kind of

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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assays, and it remains largely unexplored whether the pH of SVs storing other neurotransmitters such as GABA, acetylcholine, and biogenic monoamines is also ~5.7. Green fluorescent protein (GFP) and its derivatives have been proven to be useful for measuring local pH, since the fluorescence intensities of most of them are sensitive to pH with distinct pKa [6, 7]. Spectroscopic data, in combination with structural analysis of GFP and its derivatives, have revealed that protonation of the key residues around the fluorophore (Tyr66) critically affects their fluorescent states (reviewed in [8]). Among such pH-sensitive GFP derivatives, a mutant named “pHluorin” has been frequently used to monitor organellar pH changes and also to monitor synaptic vesicle exo-endocytotic recycling [4, 5]. However, it is evident that, due to its apparent pKa of ~7.1, pHluorin is not the most optimal pH sensor for organelles whose luminal pH is less than 6 [9]. Therefore, it is important to select the optimal pH sensor according to the organelle of interest. Recently, we have compared luminal pH and post-endocytic re-acidification dynamics of SV pH in inhibitory GABAergic neurons using cultured neurons derived from vesicular GABA transporter (VGAT)—Venus transgenic mice [10] and mOrange2 (pKa ~ 6.5) [11] as a luminal pH sensor for SVs [12]. The pH of GABAcontaining vesicles was found to be much higher (pH 6.4 on average) than that of glutamate-containing vesicles, and that re-acidification during GABA reuptake exhibited a unique biphasic behavior that indicated distinct proton coupling upon GABA transport [12]. In this article, we described experimental details and made some important notes that may help researchers to apply this method.

2

Materials The method included multiple steps, including production of lentiviral vectors to express the pH probe, primary neuronal cultures derived from postnatal mouse brain, and live fluorescent imaging upon the local application of distinct solutions to the cells. The preparation of cDNA constructs expressing pH-sensitive fluorescent proteins requires conventional molecular cloning laboratory equipment. Neuronal cell cultures and lentiviral vector production require conventional cell culture equipment ideally with two CO2 incubators. Live imaging can be performed on an inverted fluorescent microscope equipped with >60
objective lens [e.g., UPlanSApo 60
/1.35 Oil (Olympus, Japan)]. To apply solutions with defined composition in regions of interests (ROIs), a microperfusion system with a custom-made microcapillary was constructed (see Note 1).

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2.1 Molecular Biology

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A combination of plasmids was used to produce lentiviral vectors that introduce the pH probe to the luminal side of synaptic vesicles in cultured neurons. The pH probe was generated by fusing mOrange2 to a luminal side of synaptophysin and referred to as syp-mOr [9, 12] (see Notes 2 and 3). 1. Lentiviral transfer plasmid (pLenti6PW) which encodes either tTAad under the synapsin 1 promoter (syn-tTA) or the pH probe syp-mOr under the TRE promoter (TRE-syp-mOr) [13] (see Fig. 2). 2. Helper plasmids (pGAG-kGP1, pCAG-RTR2, pCAG-VSVG) are used for packaging lentivectors in HEK293T cells.

2.2 Production of Lentiviruses

1. HEK293T cell line. 2. Culture medium: DMEM containing 10% fetal bovine serum (FBS), 2 mM L-glutamine, 1 mM sodium pyruvate, 100 units/ mL penicillin, and 100 units/mL streptomycin. 3. 2.5 M CaCl2. 4. BES-buffered saline (2
BBS): 50 mM BES sodium salt, 280 mM NaCl, and 1.5 mM Na2HPO4 at pH 6.95. 5. Filter units (0.45 μm).

2.3 Neuronal Cultures

1. Glass bottom dishes (ϕ35 mm) with glass microwells (ϕ27 mm; see Note 4). 2. Poly-D-lysine (PDL; M.W. 30,000–70,000). 3. Papain suspension. 4. Culture medium: Neurobasal-A medium containing 2% B27 supplement and 0.5 mM L-glutamine. 5. Plating medium: culture medium supplemented with 5% FBS. Serum can be obtained from any commercial sources, but it is recommended to test several lots to find suitable ones for neuronal cultures. 6. Fluorodeoxyuridine. 7. Uridine. 8. Hemocytometer. 9. CO2 (5%) humidified incubator (ideally 3% CO2 humidified incubator in addition).

2.4 Fluorescence Microscope Setup (Fig. 1)

1. Inverse fluorescence microscope with appropriate excitation/ emission filter set and wheel changer [e.g., Lambda (Sutter Instruments, Novato, USA)]. 2. Appropriate digital camera [e.g., a scientific cMOS camera (Andor, Belfast, UK)].

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Fig. 1 Schematic drawing of our imaging setup with the fast-flow perfusion system. Neurons cultured on a glass bottom dish were continuously perfused with standard extracellular solution using gravity-driven flow. Solutions used during image acquisition were directly applied onto the area of interest with custom-made microperfusion capillaries (see Fig. 5), whose tips were positioned closely above the target area. The bundle of microperfusion capillaries mounted on a fast stepper was connected to soft tubes that open and close by pinch valves. Fast stepper and pinch valves were simultaneously controlled by Clampex software, allowing efficient perfusion with high spatial and temporal precision. A puff pipette was used for application of the mixture of ionophores, whose timing and duration were also controlled by Clampex

3. Software for acquisition and analysis of images [e.g., MetaMorph (Molecular Devices, San Jose, USA)]. 4. Perfusion system with gravity-driven flow. 5. Microperfusion system with a combination of fast-flow exchange microperfusion device [e.g., SF-77B (Warner Instruments, Hamden, USA)] and a valve controller [e.g., PS-8H (Bioscience tools, San Diego, USA)]. For efficient perfusion with high spatial and temporal precision, use a custom-made microperfusion capillary (see Note 1). 6. Suction device with an air pressure sensor [e.g., PC-21 (Narishige, Tokyo, Japan)]. 7. Software to control the microperfusion system and the valve controller [e.g., Clampex 10 (Axon Instruments, San Jose, USA)]. 8. Puff pipette connected to a pulse pressure device [e.g., picopump (WPI, Sarasota, USA)] to apply a cocktail of ionophores onto the imaging region in a spatially limited manner (see Note 5).

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9. Standard extracellular solution: 140 mM NaCl, 2.4 mM KCl, 10 mM HEPES, 10 mM glucose, 2 mM CaCl2, 1 mM MgCl2, 0.02 mM CNQX, and 0.025 mM D-APV at pH 7.4. 10. K+-rich calibration solutions adjusted at a given pH: 122.4 mM KCl, 20 mM NaCl, 10 mM HEPES or MES, 10 mM glucose, 2 mM CaCl2, and 1 mM MgCl2. Use MES buffer for solutions with pH set to below 6.2 and HEPES buffer for solutions with pH set above 6.8.

3

Methods

3.1 Production of Lentiviral Vector Encoding a pH Sensor for Synaptic Vesicle Lumen

1. Add 5 mL of 0.01% PDL to a 75 cm2 flask and swirl to spread over the whole surface. 2. Incubate at room temperature (RT; 24–26  C) for at least 30 min. 3. Remove PDL (or collect into another flask) and wash twice with sterilized distilled water (DW). 4. Add PBS and incubate at RT or in a 5% CO2 incubator until use (PDL solution can be reused twice). 5. Seed HEK293T cells at a concentration of 50,000 cells/ 10 mL/ϕ10 cm dish in the morning and incubate in the 5% CO2 incubator to reach 50–70% confluency by the evening. 6. On the next day, split one-third of HEK293T cells from the ϕ10 cm dish (~90% confluent) onto a PDL-coated flask. 7. Two hours after passage, perform transfection. For a single flask, mix the following amount of plasmids and 50 μL of 2.5 M CaCl2 and adjust the volume to 500 μL with sterilized DW. Lentiviral transfer plasmid (pLenti6PW): pCAG-kGP1: pCAG4-RTR2:pCAG-VSVG ¼ 17:10:5:5 μg/75 cm2 flask. 8. Add 500 μL of 2
BBS and incubate at RT for 20 min. A very slight turbidity of the mixture will become apparent within seconds. Note that transfection efficiency is critically dependent on the pH of BBS [14]. 9. Add transfection mixture onto the cells dropwise and transfer to a 3% CO2 incubator. Note that using 5% CO2 incubator results in slightly lower transfection efficiency (see Note 6). 10. Sixteen hours after transfection, remove medium, wash once with sterilized PBS, and add fresh neuron culture medium (10 mL/75 cm2 flask). 11. After 48 h, collect culture medium into a 50 mL tube through a 0.45 μm filter unit.

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12. Aliquot into 1.5 mL screw capped tubes and freeze them in liquid nitrogen. Store at 80  C until use (see Note 7). 3.2 Preparation of Cultured Neurons Expressing Vesicular pH Probe

1. Coat glass bottom dishes with 0.5 mg/mL PDL dissolved in 0.1 M sodium tetraborate buffer (pH 8.4) overnight. After washing with sterile DW twice, add sterile PBS and incubate at RT until use (see Note 8). 2. Dissect hippocampi from 0- to 1-day-old ICR or C57BL/ 6 mice. 3. Transfer hippocampi into a 1.5 mL tube containing HBSS supplemented with 1/500 volume of papain suspension and incubate at 37  C with mild shaking using thermomixer. After 20 min of incubation, add 1/100 volume of 0.8% DNase solution and further incubate at 37  C for 20 min. 4. After digestion, collect cells by centrifugation and resuspend with plating medium. Count the cell density using a hemocytometer and adjust the density to 2.8–3.4
105 cells/mL with plating medium. Plate 0.5 mL of cell suspension onto the PDL-coated glass microwell (ϕ27 mm) of the glass bottom dish, resulting in a cell density of 2.5–3
104 cells/cm2. Incubate in 5% CO2. 5. Two to four hours after plating, add 1.5 mL of normal culture medium and incubate again in 5% CO2. 6. At 3 days in vitro (DIV), add 40 μM fluorodeoxyuridine and 100 μM uridine to inhibit the growth of glial cells. Replace one-third of the culture medium with fresh medium every 3–4 days. 7. Transfect neuronal cultures with 5–50 μL each of syn-tTAlentiviral solution and pH-probe-lentiviral solution at 6–7 DIV and subject to experiments at 13–21 DIV (Fig. 2) (see Note 9).

syn-tTA

5’LTR RRE pA

TRE-Syp-mOr

3’LTR tTAad

co-transfection

SYN WPRE

cPPT pA Syp-mOr

TRE

WPRE

Fig. 2 Schematic drawing of a pair of the lentiviral vectors that depend on the Tet-Off system to drive syp-mOr expression in a neuron specific manner. Transgene sequences flanked by long terminal repeat (LTR) sequences, which facilitate the integration into the host genome, are shown. The regulator vector (syn-tTA) expresses an advanced tetracycline transactivator (tTAad) (1) under the control of synapsin 1 promoter and the response vector (TRE-syp-mOr) expresses syp-mOr (3) in the presence of tTAad (2)

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3.3 Estimation of the pKa and Hill Coefficient of pH Probes In Situ

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For precise determination of the luminal pH of interest, it is recommended to determine the pKa and Hill coefficient of the respective probes in situ, since the conjugation of the pH-sensitive fluorescent protein to the organelle-specific cargo protein may alter their biophysical properties. In the example given below, we determined the pKa and Hill coefficient for pHluorin and mOrange2 fused to the synaptic vesicle marker, synaptophysin. 1. Treat neurons for 1 min with a cocktail of ionophores composed of 20 μM FCCP, 10 μM valinomycin, 10 μM nigericin, 2 μM bafilomycin A1, and 0.02% Triton-X 100 in a standard extracellular solution to equilibrate the vesicular pH to extracellular pH. 2. Place a glass bottom dish on which neurons transduced with either sypHy or syp-mOr are cultured in a custom-made chamber. Perfuse the neurons continuously with extracellular solution (see Note 10). 3. Expose the neurons for 14 s to K+-rich calibration solutions, each adjusted at a given pH. 4. During application of the solutions described above, acquire fluorescence images in a time-lapse mode at 0.5 Hz with appropriate exposure time (typically 100–200 ms). 5. For calculation, subtract the background signals from signals; normalize the fluorescent intensities at each pH to that at pH 8.6 (Fig. 3) and fit with the following equation:

Fig. 3 Titrations of sypHy (green) and syp-mOr (red) expressed in cultured hippocampal neurons. (a) pHluorin and mOrange2 were positioned at the luminal part of synaptophysin. (b) Representative fluorescence changes of sypHy and syp-mOr during sequential application of distinct pH solutions. Fluorescence intensities at each pH were normalized to those at pH 8.6. Fluorescent intensities during the last 6 s of each pH step were averaged and used for an analysis. (c) Average relative fluorescence was expressed as a function of pH. Both data were well fitted by a single site titration model. The pKa and Hill coefficient were determined as 6.54  0.05 and 0.99  0.02 for syp-mOr, and 7.09  0.02 and 1.35  0.02 for sypHy (n ¼ 6 experiments with 38 boutons for sypHy and n ¼ 5 experiments with 64 boutons for syp-mOr)

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F F pH 8:6

B  ¼Aþ 1 þ 10nH
ðpK apHÞ

where pKa is the pH at which 50% of the probe molecules are protonated and nH is the Hill coefficient which is proportional to slope of the fitting curve at pKa. Parameters A and B are signal offset and gain, respectively. pKa and nH values (as well as A and B) were obtained by the least squares method using Microsoft Excel software. In case of sypHy and syp-mOr, the pKa were measured as 7.1 and 6.5, whereas the nH value was measured as 1.35 and 1.0, respectively [9]. 3.4 Estimation of Vesicular pH

A method to estimate vesicular pH of SV lumen has been developed by Mitchell and Ryan [15], by which the vesicular pH as well as the fractional expression on the cell surface can be simultaneously determined. In the original study, 50 mM NH4Cl was used to clamp cytoplasmic as well as vesicular pH at pH 7.4 [16]. However, we noticed that vesicular pH during NH4Cl application can be higher or lower than 7.4, depending on the buffering capacity of the lumen of interest [12]. Therefore, it is recommended to use a mixture of ionophores whose pH is set at pH 7.4. We also noted that sypHy is not suitable to accurately measure SV pH due to its relatively high pKa and large nH. The results obtained by syp-mOr imaging are given below. 1. Prepare a glass bottom dish on which transduced neurons are cultured as above. 2. Expose neurons to an acidic solution (pH 5.5) for 3 s, and then to a mixture of ionophores (pH 7.4) for 6 s. Measure the fluorescence of the probes (syp-mOr) during acid quenching (FQ) and ionophore application (FpH7.4) as averages during application. Example traces recorded from glutamatergic boutons and GABAergic boutons separately are shown in Fig. 4. Venus fluorescence was obtained by a snapshot and then mOrange2 fluorescence was imaged in a streaming mode at 5 Hz under control of MetaMorph (see Note 11). The boutons were categorized into glutamatergic and GABAergic according to the absence and the presence of Venus fluorescence, respectively. 3. Calculate vesicular pH using the following equations. Essentially, the observed fluorescent signals in a given synaptic terminal are thought to be the sum of fluorescence derived from the cell surface fraction (S) that experiences extracellular pH and from the vesicular fraction (1 – S) that experiences SV pH (pHv). Therefore, the fluorescence during the baseline period (F0) and the fluorescence during FQ can be expressed as follows:

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Fig. 4 SV pH measurement with syp-mOr in cultured neurons prepared from VGAT-Venus Tg mice. (a) Syp-mOr (red) was expressed in VGAT-Venus Tg neuronal culture, where GABAergic neurons specifically express yellow fluorescent protein, Venus (green). Filter sets that effectively separate the two fluorescence spectra were used (see Note 10). Live fluorescence imaging of syp-mOr (average of five consecutive images) at rest and on addition of a mixture of ionophores at pH 7.4 are also shown. Scale bar indicates 5 μm. (b) Representative fluorescent changes of syp-mOr during acid quenching and upon puff application of a mixture of ionophores. Venus-negative glutamatergic boutons (Glu) and Venus-positive GABAergic boutons (GABA) were separately analyzed. (c) Average SV pH and surface fraction were calculated according to the method described here. SV pH of GABAergic boutons (pH 6.44  0.03, n ¼ 14 experiments with 89 boutons) was significantly higher than that of glutamatergic boutons (pH 5.80  0.04, n ¼ 16 experiments with 160 boutons). ***P < 0.001, unpaired t test

F 0 ¼ S
F pH 7:4 þ ð1  S Þ
F pHv

ð1Þ

F Q ¼ S
F pH 5:5 þ ð1  S Þ
F pHv

ð2Þ

and where FpH5.5 and FpHv are the total fluorescence values predicted when all probe molecules in the terminals are exposed to pH 5.5 and pHv, respectively. By solving eqs. (1) and (2), S and FpHv/FpH7.4 were calculated as follows: S¼

F0 F pH 7:4

F

 F pHQ7:4 F

5:5 1  F pH pH 7:4

and F0 F pHv F pH 7:4  S ¼ 1S F pH 7:4

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F0/FpH7.4, FQ/FpH7.4 can easily be obtained from the above measurements, whereas FpH5.5/FpH7.4 was given according to the Henderson–Hasselbalch equation as follows: 1

F pH 5:5 1þ10nH
ðpK a5:5Þ ¼ 1 F pH 7:4 nH
ðpK a7:4Þ 1þ10

Using pKa and nH measured in the previous section, SV pH (pHv) is then calculated as follows:   nH
ðpka7:4Þ Log 1þ10  1 F pHv=F pH 7:4 pHV ¼ pK a  nH

4

Notes 1. This protocol includes perfusion of several different pH solutions and a buffer containing a cocktail of ionophores. To ensure local perfusion of the solution within the area of imaging, we conjugated a hand-made capillary bundle to the fastflow microperfusion device, and the stepwise horizontal movement and opening of the valve controller were synchronized under the control of Clampex 10, so that the perfused solutions were always flowed into the area of imaging. We have briefly described the construction of the capillary bundle below (Fig. 5). Since the size of the single cell culture preparation (autaptic culture), routinely used for our experiments, is about 300 μmsquares [17, 18], the inner diameter of the capillary needs to be 300 μm or more to ensure perfusion of the extracellular solution within the whole area of a single neuron. Therefore, a fused silica capillary having an inner diameter (ID) closest to 300 μm was adopted (OD: 430 μm, ID: 320 μm, Trajan Scientific Japan, Kanagawa, Japan). The step width of the capillary was set at 500 μm because Fast Step Perfusion System (SF-77B, Warner Instruments, Hamden, USA) can arbitrarily set the step width by a factor of 100 μm units. Since the outer diameter (OD) of the capillary is 430 μm, it is necessary to arrange the capillaries at 70 μm intervals for the 500 μm-steps. In order to evenly arrange the capillaries at 70 μm intervals, a groove of 200 μm-width was hollowed out with 500 μm depth at 300 μm intervals in an acrylic block (Fig. 5a). The design of the acrylic block is shown in Fig. 5b. Three blocks were mounted on the acrylic board (Fig. 5c). As shown in Fig. 5d, the capillaries were set on the grooves in a way aligning their tips. Because there are eight

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Fig. 5 Procedure to construct a custom-made microcapillary bundle. (a) An image of an acrylic block with grooves to arrange the capillaries. The scale is shown in b. (b) A drawing design of the acrylic block. (c) An image of the acrylic blocks mounted on an acrylic board. (d) Configuration of the arrangement of the capillaries. Note that the applied epoxy glue should not attach to the acrylic blocks. (e) An image of a hand-made capillary bundle. The left side is the tip of the capillary bundle

grooves in the acrylic block, up to eight capillaries can be loaded, if necessary. The capillary bundle was then fixed by epoxy glue (Fig. 5d). An image of a hand-made capillary bundle is presented in Fig. 5e. The horizontal stepping of the capillary bundle sideways allows quick exchange of multiple solutions with different compositions. 2. DNA encoding syp-mOr was generated by replacing pHluorinencoding sequence of sypHy [19] with mOrange2-encoding sequence. pHluorin and mOrange2 are located in the second luminal loop of synaptophysin in sypHy and syp-mOr, respectively. 3. In case of a pH probe for the synaptic vesicle lumen, a number of synaptic vesicle proteins have been used for specific targeting to synaptic vesicles. In addition to synaptophysin used in this study, synaptobrevin 2, synaptotagmin 1, SV2, and vesicular neurotransmitter transporters (VGLUT1 and 2, VGAT) have been used to probe synaptic vesicles [20, 21]. It has been demonstrated that pHluorin-fused cargo proteins are differentially distributed between synaptic vesicle membrane and the presynaptic plasma membrane [20]. Furthermore, although it

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has not been rigorously assessed in a quantitative manner, some of the pHluorin-fused cargo proteins, such as VGLUT1pHluorin, can be missorted to non-SV compartments, which do not undergo activity-dependent exocytosis (our unpublished observation). Therefore, one needs to ensure that pHluorin-fused cargo proteins are correctly sorted into synaptic vesicles, e.g., by immunofluorescence or by monitoring exocytic responses during repetitive stimulation. 4. In our experience, the use of 27 mm microwell dishes generally results in healthier cultures compared to smaller ones. Although we have used 15 mm microwell dishes several times to save the amount of cell suspension required per dish, the cultured neurons were not suitable to be subjected to the live imaging experiments described here. 5. The cocktail of ionophores, which also contains low concentration of Triton X-100, easily affects the condition of cultured cells. Therefore, their effects should be minimized as much as possible before the imaging experiment begins. The custommade microperfusion capillary was not suitable for ionophore cocktail application since a slight amount of diffusion out of the capillary may be present even when the flow is shut off. Thus, we adopted puff application for ionophores. The puffer pipette had a tip diameter of 3–5 μm and was generally positioned ~1 mm in the horizontal plane and 500–700 μm above from the center of imaging region. The air pressure should be optimized pipette by pipette to obtain stable fluorescence signal upon application of ionophore mixture. 6. Transfection efficiency also depends on cell density. It appears that HEK293T cells closely attached to neighboring cells show low transfection efficiency. Therefore, transfection should be performed before the cells start dividing after the last passage. 7. Viral titer was estimated using the Lenti-X qRT-PCR Titration Kit (Takara, Kusatsu, Japan) according to the manufacturer’s instructions. Routinely, 0.1–10
105 copies/μL viral solutions were obtained and cultured neurons were transduced with 5–50 μL viral solution per dish (see also Note 9). 8. Cleaning the glass surface prior to PDL coating seems important for achieving good cell culture quality. Glass microwells were washed with 1 M KOH for 2–4 h with mild shaking. KOH should be completely removed by vigorous rinsing with water in 1 L beaker. Dishes were dried up, sterilized with UV irradiation and then coated with PDL solution. 9. For a good quality imaging, a proportion of probe-expressing neurons is preferably less than ~50% to avoid overlap of fluorescent signals. Accordingly, we recommend not to use excess amount of viral solution. Normally, the transduction efficacy of

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individual viral lots should be tested to find the appropriate volume of a pair of viral solutions to transduce 20–50% of cultured neurons. 10. Treatment of cultured neurons with a cocktail of ionophores that also contains Triton X-100 notably kills neurons and leads to cell degeneration. Therefore, the imaging experiment after this treatment should be completed as quick as possible. 11. Venus fluorescence was imaged with 470/22 nm excitation and 514/30 nm emission filters. Syp-mOr fluorescence was imaged with 556/20 nm excitation and 600/50 nm emission filters. Although the fluorescent spectrum of Venus and mOrange2 are relatively close, the above filter sets effectively separate each other at the expense of signal intensities and no bleed-through was observed.

Acknowledgments This work was supported in part by grants from JSPS KAKENHI (16H04675), the JSPS Core-to-Core Program, A. Advanced Research Networks grant, and a research grant from Takeda Foundation to S.T., from JSPS KAKENHI (16K18397) to Y.E and from JSPS KAKENHI (25350988 and 17K08328) to S.K. Finally, we would like to thank Editage (www.editage.jp) for English language editing. References 1. Grabe M, Oster G (2001) Regulation of organelle acidity. J Gen Physiol 117(4):329–344. https://doi.org/10.1085/jgp.117.4.329 2. Schultz ML, Tecedor L, Chang M et al (2011) Clarifying lysosomal storage diseases. Trends Neurosci 34(8):401–410. https://doi.org/ 10.1016/j.tins.2011.05.006 3. Edwards RH (2007) The neurotransmitter cycle and quantal size. Neuron 55(6): 835–858. https://doi.org/10.1016/j.neu ron.2007.09.001 4. Miesenbock G, De Angelis DA, Rothman JE (1998) Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature 394(6689):192–195. https://doi.org/10.1038/28190 5. Sankaranarayanan S, De Angelis D, Rothman JE et al (2000) The use of pHluorins for optical measurements of presynaptic activity. Biophys J 79(4):2199–2208. https://doi.org/10.1016/ S0006-3495(00)76468-X 6. http://www.fpvis.org/FP.html

7. Shaner NC, Steinbach PA, Tsien RY (2005) A guide to choosing fluorescent proteins. Nat Methods 2(12):905–909. https://doi.org/ 10.1038/nmeth819 8. Choy E, Philips M (2001) Green fluorescent protein-tagged Ras proteins for intracellular localization. Methods Enzymol 332:50–64. https://doi.org/10.1016/s0076-6879(01) 32191-2 9. Egashira Y, Takase M, Takamori S (2015) Monitoring of vacuolar-type H+ ATPasemediated proton influx into synaptic vesicles. J Neurosci 35(8):3701–3710. https://doi. org/10.1523/JNEUROSCI.4160-14.2015 10. Wang Y, Kakizaki T, Sakagami H et al (2009) Fluorescent labeling of both GABAergic and glycinergic neurons in vesicular GABA transporter (VGAT)-venus transgenic mouse. Neuroscience 164(3):1031–1043. https://doi. org/10.1016/j.neuroscience.2009.09.010 11. Shaner NC, Lin MZ, McKeown MR et al (2008) Improving the photostability of bright

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monomeric orange and red fluorescent proteins. Nat Methods 5(6):545–551. https:// doi.org/10.1038/nmeth.1209 12. Egashira Y, Takase M, Watanabe S et al (2016) Unique pH dynamics in GABAergic synaptic vesicles illuminates the mechanism and kinetics of GABA loading. Proc Natl Acad Sci U S A 113(38):10702–10707. https://doi.org/10. 1073/pnas.1604527113 13. Hioki H, Kuramoto E, Konno M et al (2009) High-level transgene expression in neurons by lentivirus with Tet-off system. Neurosci Res 63(2):149–154. https://doi.org/10.1016/j. neures.2008.10.010 14. Chen C, Okayama H (1987) High-efficiency transformation of mammalian cells by plasmid DNA. Mol Cell Biol 7(8):2745–2752. https:// doi.org/10.1128/mcb.7.8.2745 15. Mitchell SJ, Ryan TA (2004) Syntaxin-1A is excluded from recycling synaptic vesicles at nerve terminals. J Neurosci 24(20): 4884–4888. https://doi.org/10.1523/ JNEUROSCI.0174-04.2004 16. Fernandez-Alfonso T, Ryan TA (2008) A heterogeneous “resting” pool of synaptic vesicles that is dynamically interchanged across boutons in mammalian CNS synapses. Brain Cell Biol 36(1–4):87–100. https://doi.org/10. 1007/s11068-008-9030-y

17. Katsurabayashi S, Kawano H, Ii M et al (2016) Overexpression of Swedish mutant APP in aged astrocytes attenuates excitatory synaptic transmission. Physiol Rep 4(1):e12665. https://doi.org/10.14814/phy2.12665 18. Wojcik SM, Katsurabayashi S, Guillemin I et al (2006) A shared vesicular carrier allows synaptic corelease of GABA and glycine. Neuron 50(4):575–587. https://doi.org/10.1016/j. neuron.2006.04.016 19. Granseth B, Odermatt B, Royle SJ et al (2006) Clathrin-mediated endocytosis is the dominant mechanism of vesicle retrieval at hippocampal synapses. Neuron 51(6):773–786. https://doi. org/10.1016/j.neuron.2006.08.029 20. Pan PY, Marrs J, Ryan TA (2015) Vesicular glutamate transporter 1 orchestrates recruitment of other synaptic vesicle cargo proteins during synaptic vesicle recycling. J Biol Chem 290(37):22593–22601. https://doi.org/10. 1074/jbc.M115.651711 21. Santos MS, Park CK, Foss SM et al (2013) Sorting of the vesicular GABA transporter to functional vesicle pools by an atypical dileucine-like motif. J Neurosci 33(26): 10634–10646. https://doi.org/10.1523/ JNEUROSCI.0329-13.2013

Chapter 5 Investigation of Synaptic Vesicle Proteins in Rat Brain Tissue Using Real-Time qPCR Betina Elfving Abstract For many years real-time quantitative polymerase chain reaction (qPCR) has been the golden standard to measure gene expression levels in brain tissue. However, today it is generally accepted that many factors may affect the outcome of the study and more consensus is required to perform and interpret real-time qPCR experiments in a comparable way. Here we describe the basic techniques used for more than a decade in our laboratory to extract RNA and protein from the same piece of frozen brain tissue and to quantify relative mRNA levels with real-time qPCR and SYBR Green. Key words RNA extraction, Brain tissue, PARIS kit, Precellys, Mixer Mill, Real-time qPCR, Reference genes, SYBR Green

1

Introduction For more than decades, laboratories worldwide have been using real-time quantitative polymerase chain reaction (qPCR) to investigate the regulation of synaptic vesicle proteins at the mRNA level in brain tissue [1]. Real-time qPCR is a sensitive molecular method characterized by the collection of data during the PCR process and not as in conventional PCR measuring the end product. The two most common chemistries used in real-time qPCR experiments are the Taqman based and the SYBR Green based detection, respectively. Both methods have been widely used and it should be emphasized that the performance and the quality of the results is usually comparable after optimization of the SYBR Green method. However, even though real-time qPCR seems a straightforward method for quantification of mRNA levels and Bustin et al. in 2009 published a set of guidelines to ensure a higher consensus of how to conduct and interpret real-time qPCR measurements [2], it has recently been reported that the majority of the published real-time qPCR mRNA data is of fluctuating quality [3]. Therefore,

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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it should be emphasized that investigating mRNA levels in brain tissue with real-time qPCR require a number steps that may be critical to the outcome of the study. Some of the handling procedures that may be reflected in the reported results and therefore require special attention are tissue homogenization, RNA extraction, cDNA synthesis, primer design, real-time qPCR measurements, and normalization of data. To ensure reproducible realtime qPCR results a number of quality checks are required as well as a proper selection of reference genes for normalization to be used in brain tissue [4]. In my laboratory we have established a method from inception to completion that enables investigation of synaptic vesicle proteins using real-time qPCR and SYBR Green [5]. Furthermore, as correlation analysis in large-scale data sets reports only 50% correspondence between mRNA and protein levels [6, 7] and brain lateralization may have critical impact in many brain disorders, we have established a standardized protocol using the Ambion® PARIS ™ system (ThermoFisher Scientific) for the isolation of RNA and protein from the same piece of frozen brain tissue [5, 8].

2

Materials

2.1 Extraction of RNA and Protein with PARIS™ Kit

Remember always to wear cloves and use RNase free tubes, pipettes, and tips. 1. Ambion® PARIS ™ kit (ThermoFisher Scientific). Follow the protocol provided by the distributor. 2. 0.1% DEPC water: 1 mL DEPC solution, add ultrapure water to 1 L. Leave it overnight at 37
C before autoclavation. Store DEPC water at 4
C. 3. Protease inhibitor: cOmplete™, EDTA-free protease inhibitor cocktail (Merck), dissolve 1 tablet in 2 mL DEPC water. Aliquot and store at 20
C. 4. Phosphatase inhibitors (optional): 100 mM NaF and 100 mM Na3VO4 (see Note 1). 100 mM NaF solution: Add 8 mL ultrapure water to 0.041988 g NaF, stir and fill up to 10 mL with ultrapure water. 100 mM Na3VO4 solution: Add 8 mL ultrapure water to 0.18391 g Na3VO4 and stir. Depending on the pH of the solution, slowly add either 1 M NaOH or 1 M HCl with stirring to adjust pH to 10. Adding HCl will make the solution yellow. Boil solution by heating in a microwave for 5–15 s. After boiling for 5–15 s, the solution will be clear and colorless. Cool on ice until the Na3VO4 solution reaches room temperature. At this point, the pH will be greater than 10. Add a small amount (several drops, with stirring) of 1 M HCl to adjust solution pH to 10. Repeat steps 3–5 in a total of 3–5 times.

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After several cycles of boiling, cooling, and adjusting pH, the solution should reach a point where the pH stabilizes at ~10. At this point, adding HCl should result in little, if any, appearance of yellow color in the solution. Fill up to 10 mL with ultrapure water. Aliquot and store the solutions at 20
C. 5. Homogenization with Precellys Evolution—tissue homogenizer (Bertin-Instruments) or Mixer Mill MM400 (Retsch). 5 mm zirconia ceramic beads or stainless steel beads are used. Beads for homogenization are kept at 4
C. 2.2 RNA Quality and Quantity Check

1. Nanodrop™ (ThermoFisher Scientific), determination of the RNA concentration and the purity (A260/280 and A260/230 ratios). 2. QIAxcel Advanced System, determination of the RNA integrity score (RIS), an objective quality measurement.

2.3

cDNA Synthesis

1. iCycler Thermal Cycler (BIORAD). 2. 10 mM dNTP mix. 3. 3 μg/μL Random primers (ThermoFisher Scientific). Random primers are diluted 12 times in DEPC water to 250 ng/μL. The dilution is stored at 20
C. 4. Superscript® IV RT (ThermoFisher Scientific). Total RNA from tissue is reversely transcribed using random primers (1 μL of 250 ng/μL per 20 μL reaction) and Superscript IV RT according to the manufacturers’ protocol.

2.4

Real-Time qPCR

1. The Mx3005P or AriaMx real-time qPCR instrument. 2. 96-well plates, Sorenson™, Bioscience, Inc. 3. SYBR® Green JumpStart™ Taq ReadyMix (Sigma-Aldrich). 4. Selected primer pairs for reference and target genes, respectively. 5. Normalization strategy (selection of reference genes, Normfinder (Excel add-in)).

3

Methods

3.1 Collection of Brain Tissue

1. Euthanize the rats by decapitation. 2. Isolate the brain and dissect brain areas by hand on an ice-cold tile, i.e., left/right prefrontal cortex, left/right hippocampus, and cerebellum. 3. Immediately freeze the tissue on powdered dry-ice. 4. Collect the tissue into tubes (2 mL Eppendorf) and store at 80
C until further processing.

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5. Collect the weight of each piece of tissue (~50 mg hippocampus, ~25 mg prefrontal cortex). 3.2 Homogenization of Brain Tissue

1. Clean the lab bench, pipettors, and equipment with RNase decontamination solution before initiating the extraction of RNA. 2. Prepare all solutions as described in the Ambion® PARIS™ kit protocol. 3. Add the cOmplete™, EDTA-free protease inhibitor cocktail (16 μL per 400 μL Cell Disruption Buffer) to the Cell Disruption Buffer, and eventually NaF (20 μL of 100 mM solution per 400 μL Cell Disruption Buffer) and Na3VO4 (0.4 μL 100 mM solution per 400 μL Cell Disruption Buffer) (see Note 1). Keep the solution on ice. 4. Place the Eppendorf tubes with tissue on ice. 5. Add Cell Disruption Buffer with protease/phosphatase inhibitors (8 volumes per tissue mass; 50 mg tissue ! 400 μL Buffer) to the tube. 6. Place a cold bead in each tube. 7. Homogenize the tissue using either the Mixer Mill MM 400 (48 samples, 1 min, 30 Hz/s) or the 3-dimensional beadbeating technology of the Precellys Evolution Homogenizer (24 samples, 2  15 s, 5000 rpm). Both have the advantages of homogenizing from 24–48 samples per run. 8. Visually inspect the tubes after homogenization to ensure that tissue is completely homogenized. 9. If the tissue is not completely homogenized, then the procedure is repeated (see Note 2). 10. In case the homogenization procedure leads to foam formation, then leave the samples on ice for 5 min or more before proceeding with RNA extraction.

3.3 Extraction of RNA and Protein from the Brain Samples

1. Centrifuge the samples (3 min, 800  g, 4
C). 2. Move the supernatant to a new 2 mL tube (avoid the white precipitation; waste about 75 μL). 3. 50–150 μI of each sample is transferred to new 1.5 mL tubes for Western blotting analysis. Usually, 50–100 μI for prefrontal cortex and 100–150 μI for hippocampus. Store 5 μL for determination of the protein concentration. 4. Measure the exact volume of the remaining supernatant and mix it with an equal volume of 2 Lysis/Binding Solution. 5. Carry out the rest of the extraction as described in the Ambion® PARIS ™ kit protocol (Section: C. RNA Isolation, page 12).

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6. After elution of RNA, take aliquots (2  3 μL) for quality and quantity check using the Nanodrop™ and the QIAxcel Advanced System. 7. Store the RNA samples at 80
C. 3.4

1. Use the sample with the lowest RNA concentration from the Nanodrop measurement as the input concentration in the cDNA synthesis. Normally the RNA concentrations vary from 80 to 300 ng/μL.

cDNA Synthesis

2. Conduct the cDNA synthesis using random primers and Superscript® IV RT (see Note 3). 3. Store the cDNA samples at 80
C. 3.5

1. Include 2 No Template Controls (NTCs) and a standard curve with duplicates of standard solutions (S1, S2, S3, S4, and S5) with dilution factor 5 on each plate (Table 1). The standard solutions are a mixture of all the included samples in the experiment.

Real-Time qPCR

2. Based on the amount of samples in the specific experiment, the real-time qPCR measurement will either be determined in singlets or duplets. 3. Dilute the samples to 1.5 ng/μL based on the relative cDNA synthesis concentration and S1 to 15 ng/μL (The singlet plate layout is presented in Table 1). 4. Generate a stock plate including cDNA dilutions for the entire experiment (reference genes and target genes). 5. The stock plate may be kept at 4
C for 3–4 weeks. If the experiment cannot be conducted within a month, the diluted cDNA solutions should be dispersed into individually Sorenson Plates and kept at 20
C (see Note 4). Table 1 The plate layout. S1-S5: Standard 1–5; NTC: No template control; P1-P84: Sample 1–84 1

2

3

4

5

6

7

8

9

10

11

12

A

NTC

S1

S2

S3

S4

S5

NTC

S1

S2

S3

S4

S5

B

P1

P2

P3

P4

P5

P6

P7

P8

P9

P10

P11

P12

C

P13

P14

P15

P16

P17

P18

P19

P20

P21

P22

P23

P24

D

P25

P26

P27

P28

P29

P30

P31

P32

P33

P34

P35

P36

E

P37

P38

P39

P40

P41

P42

P43

P44

P45

P46

P47

P48

F

P49

P50

P51

P52

P53

P54

P55

P56

P57

P58

P59

P60

G

P61

P62

P63

P64

P65

P66

P67

P68

P69

P70

P71

P72

H

P73

P74

P75

P76

P77

P78

P79

P80

P81

P81

P83

P84

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6. Transfer 3 μL of the diluted cDNA with a multichannel pipette from the stock plate to the Sorenson Plates. In the NTC wells cDNA is exchanged with DEPC water. 7. Each real-time qPCR reaction (10 μL total volume) contains 5 μL SYBR Green, 0.5 μL of 10 μM primer pair mix, 1.5 μL DEPC water (SYBR Green mastermix), and 3 μL diluted cDNA. Add 7 μL of SYBR Green mastermix to each well. 8. Spin the plate quickly and conduct the real-time qPCR experiment. 9. The thermal conditions for the PCR are 3 min at 95
C to activate the hot-start iTaqDNA polymerase, followed by 40 cycles of 10 s denaturation at 95
C, 30 s annealing at 60
C, and 60 s extension at 72
C. Each run is completed by dissociation curve analysis to confirm the amplification specificity and absence of primer dimers, 1 min 95
C, 30 s 60
C, and 30 s 95
C. 3.6

Primer Design

1. Primers are designed to be intron-spanning, whenever possible and amplicons are selected to be 80–250 bp long. Initially the most optimal primer set is selected based on the rating with NetPrimer (http://www.premierbiosoft.com/netprimer/) (see Note 5). 2. Initially, the primer pairs are tested on standard solutions (S1, S2, S3, S4, and S5) with dilution factor 5. The relative cDNA concentration of S1 being 15 ng/μL. The test is conducted with identical tissue as used in the study. 3. The primer pairs has to fulfill the following requirements to be included in the study: (a) the amplicon size is correct when measured with 1% EtBr agarose gel electrophoresis (see Note 6), (b) only one PCR product is detected in the samples, when they are subjected to a heat dissociation protocol after the final cycle of the PCR, and (c) the real-time qPCR efficiency is 90–110%.

3.7

Normalization

In real-time qPCR experiments, sound data normalization is imperative to correct for sample-to-sample variations in RNA integrity, pipetting errors, etc. In the studies with rat brain tissue we have for many years been using 8 selected reference genes and the excel software add-in, Normfinder [4]. Normfinder calculates a stability value based on the combined estimate of both intra- and intergroup values. 1. Collect the real-time qPCR data for the 8 reference genes. 2. Use the relative mRNA values and Normfinder to select the 2 most stably expressed genes.

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3. Normalize each individual sample value of the target genes with the geometric mean of the 2 selected reference genes (Primer pairs for the 8 reference genes and selected synaptic vesicle proteins are given in Table 2) [5]. Table 2 Characteristics of gene-specific real-time qPCR primers

Gene symbol

Gene name

Accession no.a

Primer sequence

Amplicon sizeb

Reference genes 18 s rRNA

18 s subunit ribosomal RNA M11188

(+) 310 acggaccagagcgaaagcat () tgtcaatcctgtccgtgtcc

Actb

Beta-actin

NM_031144

165 (+) tgtcaccaactgggacgata () ggggtgttgaaggtctcaaa

CycA

Cyclophilin A

XM_345810

248 (+) agcactggggagaaaggatt () agccactcagtcttggcagt

Gapdh

Glyceraldehyde-3NM_017008 phosphate dehydrogenase

(+) tcaccaccatggagaaggc 168 () gctaagcagttggtggtgca

Hmbs

Hydroxy-methylbilane synthase

NM_013168

(+) tcctggctttaccattggag 176 () tgaattccaggtgagggaac

Hprt1

Hypoxanthine guanine phosphoribosyl transferase 1

NM_012583

(+) gcagactttgctttccttgg 81 () cgagaggtccttttcaccag

Rpl13A

Ribosomal protein L13A

NM_173340

167 (+) acaagaaaaagcggatggtg () ttccggtaatggatctttgc

Ywhaz

Tyrosine BC094305 3-monooxygenase/tryptophan 5-monooxygenase activa-tion protein, zeta

(+) 136 ttgagcagaagacggaaggt () gaagcattggggatcaagaa

Target genes Presynaptic markers Scamp2

Secretory carrier membrane NM_023955.1 protein 2

(+) tggctgagttcaatcccttc 196 () agctcagcagctttcctgtc

Snap25

Synaptosomal associated protein 25 kDa

NM_030991

(+) ctggcatcaggactttggtt 200 () attattgccccaggcttttt

Snap29

Synaptosomal associated protein 29 kDa

NM_053810

219 (+) acacggagaagatggtggac () tggcttggtacttgctttcc (continued)

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Betina Elfving

Table 2 (continued)

Gene symbol

a

Gene name

Accession no.a

Primer sequence

Amplicon sizeb

Snapin

NM_001025648 (+) tggatctggacccctatgtt 182 () tttgcttggagaaccaggag

Synapsin1

NM_019133

Synapsin2

NM_001034020 (+) catgggtgtttgctcagatg 127 () accacgacaggaaacgtagg

Synapsin3

NM_017109

182 (+) cacagcaagaatggcagaga () ttagtctgtggaccccaagg

Synaptophysin

NM_012664

(+) cagtgggtctttgccatctt 222 () ttcagccgacgaggagtagt

Synaptotagmin1

NM_001033680 (+) cttctccaagcacgacatca 219 () ccacccacatccatcttctt

Synaptotagmin2

NM_012665

241 (+) aggtgaaagtgcccatgaac () ctcttgccattctgcatcaa

Synaptotagmin3

NM_019122

(+) ggactccaatgggttctcag () agcaggttgtccaaaaccac

Syntaxin 1A

NM_053788

(+) accgcttcatggatgagttc 155 () gagctcctccagttcctcct

(+) 184 caccaggatgaagacaagca () gtcgttgttgagcaggaggt

234

Vamp1

Vesicle-associated membrane protein 1

NM_013090

88 (+) gtgctgccaagctaaaaagg () actaccacgattgatggcaca

Vamp2

Vesicle-associated membrane protein 2

NM_012663

(+) ctgcacctcctccaaatctt 191 () cttggctgcacttgtttcaa

Vamp5

Vesicle-associated membrane protein 5

NM_053555

147 (+) gcagaccaagtgacggaaat () ctgctgggctaaagtcttgg

Vamp7

Vesicle-associated membrane protein 7

NM_053531

(+) gttctggctgcacaactgaa 187 () aggtgacggacgaatctacg

Genbank accession number of cDNA and corresponding gene, available at http://www.ncbi.nlm.nih.gov/ Amplicon length in base pairs

b

SVPs and Real-Time qPCR

4

67

Notes 1. NaF and Na3VO4 are only added if phosphatase inhibitors are required in the Western blotting experiments. NaF is diluted to 5 mM and Na3VO4 to 1 mM in Cell Disruption Buffer. 2. Homogenization of cerebellum may be troublesome and sonication (2–3 s, 20 kHz, 40%, Sonopuls HD 2200, Bandelin) with a sonicator probe on ice has improved the real-time qPCR results in our lab. 3. To reduce costs SuperScript® IV Reverse Transcriptase may be diluted 1:1 with DEPC water. 4. We have demonstrated that repeated freezing/thawing of the diluted cDNA solution affects the real-time qPCR results. Therefore, the diluted cDNA solutions should be placed either at 4
C, or at 20
C in individual plates. We have demonstrated with cDNA from the rat hippocampus and the reference gene Gapdh that standard dilutions (S1-S5) starting at 17 ng/μ L, with dilution factor 5 may be stored at 20
C for 2 months without changes in Ct values or real-time qPCR efficiency. With the samples stored at 4
C we experienced increased Ct values for S5 after 4 weeks causing decreased real-time qPCR efficiency. 5. As SYBR Green indiscriminately binds to double-stranded DNA, other products of the PCR such as primer dimers may be detected along with the target gene. Therefore, primer pairs are tested initially. 6. To be certain about the target specificity, sequencing of the PCR product is required to prove that the intended sequence has been amplified.

References 1. Elfving B, Bonefeld BE, Rosenberg R et al (2008) Differential expression of synaptic vesicle proteins after repeated electroconvulsive seizures in rat frontal cortex and hippocampus. Synapse 62(9):662–670. https://doi.org/10. 1002/syn.20538 2. Bustin SA, Benes V, Garson JA et al (2009) The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin Chem 55(4):611–622. https://doi.org/10.1373/clinchem.2008. 112797 3. Bustin S, Nolan T (2017) Talking the talk, but not walking the walk: RT-qPCR as a paradigm for the lack of reproducibility in molecular

research. Eur J Clin Investig 47(10):756–774. https://doi.org/10.1111/eci.12801 4. Bonefeld BE, Elfving B, Wegener G (2008) Reference genes for normalization: a study of rat brain tissue. Synapse 62(4):302–309. https:// doi.org/10.1002/syn.20496 5. Elfving B, Mu¨ller HK, Oliveras I et al (2019) Differential expression of synaptic markers regulated during neurodevelopment in a rat model of schizophrenia-like behavior. Prog NeuroPsychopharmacol Biol Psychiatry 95:109669. https://doi.org/10.1016/j.pnpbp.2019. 109669 6. Tian Q, Stepaniants SB, Mao M et al (2004) Integrated genomic and proteomic analyses of

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gene expression in mammalian cells. Mol Cell Proteomics 3(10):960–969. https://doi.org/ 10.1074/mcp.M400055-MCP200 7. Pradet-Balade B, Boulme F, Beug H et al (2001) Translation control: bridging the gap between genomics and proteomics? Trends Biochem Sci 26(4):225–229. https://doi.org/10.1016/ s0968-0004(00)01776-x

8. Mu¨ller HK, Wegener G, Popoli M et al (2011) Differential expression of synaptic proteins after chronic restraint stress in rat prefrontal cortex and hippocampus. Brain Res 1385:26–37. https://doi.org/10.1016/j.brainres.2011.02. 048

Chapter 6 Mass Synaptometry: Applying Mass Cytometry to Single Synapse Analysis Chandresh R. Gajera, Rosemary Fernandez, Nadia Postupna, Kathleen S. Montine, C. Dirk Keene, Sean C. Bendall, and Thomas J. Montine Abstract Synaptic degeneration is one of the earliest and phenotypically most significant features associated with numerous neurodegenerative conditions, including Alzheimer’s and Parkinson’s diseases. Synaptic changes are also known to be important in neurocognitive disorders such as schizophrenia and autism spectrum disorders. Several labs, including ours, have demonstrated that conventional (fluorescence-based) flow cytometry of individual synaptosomes is a robust and reproducible method. However, the repertoire of probes needed to assess comprehensively the type of synapse, pathologic proteins (including protein products of risk genes discovered in GWAS), and markers of stress and injury far exceeds what is achievable with conventional flow cytometry. We recently developed a method that applies CyTOF (Cytometry by Time-Of-Flight mass spectrometry) to high-dimensional analysis of individual human synaptosomes, overcoming many of the multiplexing limitations of conventional flow cytometry. We call this new method Mass Synaptometry. Here we describe the preparation of synaptosomes from human and mouse brain, the generation and quality control of the “SynTOF” (Synapse by Time-Of-Flight mass spectrometry) antibody panel, the staining protocol, and CyTOF parameter setup for acquisition, post-acquisition processing, and analysis. Key words Synapse, Synaptosome, SynTOF (Synapse by Time of Flight), Mass Synaptometry, Human, Mass cytometry, CyTOF (Cytometry by Time of Flight)

1

Introduction Alzheimer’s disease (AD), Parkinson’s disease (PD), and Lewy body dementia (LBD) are all proposed to initiate with regional synaptic injury and degeneration [1–6]. Despite animal model support for this [7–11], the study of human synaptic changes has been mostly from tissue homogenates, Golgi stains, or electron microscopy. We and others have adapted conventional flow cytometric analysis of human synaptosomes [12–16] and shown that

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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despite using different instruments, samples, and probes, cytometry of human synaptosomes is a reproducible and robust technique. However, this approach has practical limitations on the number of distinct probes that can be used simultaneously. Like cytometry in cancer biology and immunology, the repertoire of probes needed for comprehensive assessment of the type of synapse, pathologic proteins (such as GWAS-discovered risk genes protein), and injury and stress markers far exceeds the limits of conventional flow cytometry. Cytometry by time-of-flight mass spectrometry (CyTOF) overcomes conventional flow cytometry multiplexing limitations and allows multiplexing of upwards of 20–40 antibodies. CyTOF can give detailed insights into biological complexities that are otherwise difficult, or impossible, to appreciate from the analysis of bulk tissue. Indeed, CyTOF has been used by us and others to great effect in immunology and cancer biology research to reveal cell and tissue diversity [17–26]. We recently published a novel method (“mass synaptometry”) using mass cytometry for high-throughput molecular characterization of individual synapses [27]. Prior to this, the application of mass cytometry for synaptosome analysis had not been reported, likely owing to the absence of neurosciencerelated antibody panels, methods for optimized sample preparation and acquisition protocols, and analytical pipelines for postacquisition processing. Mass synaptometry offers a new opportunity to gain fundamental insights into early synaptic changes in the brain and nervous system; such molecular signatures may serve as the basis for disease bioassays. The chapter provides enhanced experimental details for the workflow shown in our recent publication (Fig. 1; Fig. 2 in [27]). Subheading 3.1 (yellow in Fig. 1) covers proper cryopreservation of tissue from rapid autopsies, or mouse brain tissue, followed by synaptosome preparation, as we described previously [13, 14], but with modifications developed by us [27] to adapt the samples for CyTOF. As noted in Fig. 1, both brain tissue and synaptosomes can be stored at 80  C. Subheading 3.2 (represented in Fig. 1 with an asterisk) covers considerations for antibody panel design, conjugation of each antibody to a specific heavy metal ion, and quality control measures to verify specificity of staining. Subheading 3.3 (green in Fig. 1) describes staining the synaptosomes with a cocktail of antibodies and further processing the samples for mass cytometry. Subheading 3.4 (blue in Fig. 1) covers consideration of acquisition settings, post-acquisition normalization, and gating. This method yields more than 30 parameters for a single synapse multiplied by an average of 100,000 synapses per run. Although the complex data analysis required is beyond the scope of this chapter, we include a brief description (Subheading 3.5) of the analysis tools as a starting point for interested researchers.

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Fig. 1 Overview of workflow diagram of mass synaptometry procedure. Schematic showing the stepwise procedure with optimized experimental parameters, pause points, and critical steps. Yellow shading corresponds to Subheading 3.1. *Antibody cocktail design and preparation is described in Subheading 3.2. Green shading corresponds to Subheading 3.3. Blue shading corresponds to Subheading 3.4. (Adapted from Fig. 2, Gajera et al. 2019, High-dimensional multi parametric assay for single synapses. J Neurosci Methods. 2019 Jan15;312:73–83, with permission from Elsevier). P1 and S1 are the nuclei/cell debris pellet and synaptosome-containing supernatant, respectively. P2 is the crude synaptosome pellet (S2 is discarded)

2

Materials

2.1 Reagents and Disposables (See Note 1)

1. MaxPar® Antibody Labeling Kit (Fluidigm) (see Note 2). 2. Stock Antibody: carrier-free (no BSA, ovalbumin or gelatin) IgG, 100 μg (or multiples of 100) per reaction. 3. 3 kDa Amicon® Ultra-0.5 ml Centrifugal Filters (Millipore). 4. 50 kDa Amicon® Ultra-0.5 ml Centrifugal Filters (Millipore). 5. Bond-Breaker® TCEP solution (Thermo Scientific).

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6. Antibody Stabilizer (Candor Bioscience), supplemented with 0.09% sodium azide and 0.22 μm filtered. 7. 16% PFA (Alfa Aesar), freshly diluted to 1% with Maxpar PBS. 8. Ultrapure (e.g., MilliQ) water for washes and final suspension of cells. 9. 5-ml polystyrene round-bottom 12  75 mm tubes with 35 μm cell-strainer caps (Falcon/Corning) (see Note 3). 10. Maxpar PBS (Fluidigm) or Metal-free, Ca2+ and Mg2+ ion-free PBS (Rockland Immunochemicals): Make 1 final concentration by diluting 10X concentrate with MilliQ water and store in LDPE bottles. 11. Conjugated antibodies (see Table 1). 12. Maxpar Fix I Buffer (Fluidigm). 13. Cell ID™ Intercalator-Ir 125 μM (Fluidigm). 14. Maxpar 10 Barcode Perm Buffer (Fluidigm). 15. EQ™ Four Element Calibration Beads (Fluidigm). 16. Isotope-conjugated antibodies (Fluidigm). 17. MaxPar Cell Staining Buffer (Fluidigm). 18. Homogenization buffer (HB) (see Note 4). 19. Krebs-Ringer phosphate (KRP) buffer: 118 mM NaCl, 5 mM KCl, 4 mM MgSO4, 1 mM CaCl2, 1 mM KH2PO4, 16 mM sodium phosphate buffer (pH 7.4), and 10 mM glucose. 20. Cryopreservation solution (10% DMSO, 10 mM Tris–HCl buffer (pH 7.4), 0.32 M sucrose). 21. 5 μm Ultrafree-CL-centrifugal filter devices (Merck/Millipore/Sigma). 22. 0.1 μm centrifugal filters (Millipore/Sigma). 23. Benzonase® Nuclease HC, Purity >99%, 250 U/μl (EMD Millipore/Sigma). 2.2

Equipment

1. Helios™ mass cytometer with CyTOF® Software version 7.0.5189.0 (Fluidigm, CA, USA) (see Note 5). 2. Particle or cell counter (see Note 6). 3. Refrigerated centrifuge with rotor/adaptors for 1.5-ml, 2-ml, 5-ml tubes. 4. Refrigerated centrifuge with adaptors for 15-ml tubes. 5. Incubator at 37  C. 6. Water bath at 37  C. 7. NanoDrop™ or other equivalent nano- or micro-volume spectrophotometer.

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Table 1 Antibodies used in SynTOF panel. The panel comprises 36 conjugated antibodies, defining cell type, synapse type, protein products of several AD or PD risk genes, and markers of injury/response to injury. Included are clone identifiers, conjugated metal isotype, and the commercial supplier

Antibody

Marker for:

Clone ID

Metal isotype

Supplier

Neuron

NCAM16.2 163 Dy

Fluidigm

LNH-94

173 Yb

Biolegend

Cell type: Neuronal CD56

CD298-Na K ATPase Neuron Cell type: Non-neuronal CD11b (OX42)

Microglia

M1/70

144 Nd

Biolegend

GFAP

Astrocyte

GA5

143 Nd

Fluidigm

MBP

Myelin/oligodendrocyte

SMI 99

150 Nd

Biolegend

SNAP-25

Presynapse

SMI 81

155 Gd

Biolegend

CD47

Presynapse

B6H12

151 Eu

Thermo Scientific

PSD-95

Postsynapse

K28/43

157 Gd

Biolegend

DAT

Dopaminergic

hDAT-NT

154 Sm

Abcam

NET

Noradrenergic

251A9

145 Nd

Synaptic System

SERT

Serotonergic

Poly

175 Lu

Millipore Sigma

TH

Dopaminergic/catecholamine biosynthesis

2/40/15

166 Er

Biolegend

VGAT

GABAergic

117G4

153 Eu

Synaptic System

VGLUT1

Glutamatergic

N28–9

169 Tm

Biolegend

VGLUT2

Glutamatergic

321A8

172 Yb

Synaptic System

Synapse type

Negative markers/nuclear controls Ir-191

Nuclear DNA

Not 191 Ir applicable

Fluidigm

Ir-193

Nuclear DNA

Not 193 Ir applicable

Fluidigm

Discovery: Injury and response 3NT- 3Nitrotyrosine

Oxidative damage

1A6

165 Ho

Millipore Sigma

8-OH-guanosine

Oxidative damage

15A3

152 Sm

Abcam

C1q

Aging, pruning and inflammation

Poly

160 Gd

Dako/Agilent (continued)

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Table 1 (continued)

Antibody

Marker for:

Clone ID

Metal isotype

Supplier

Caspase 3 (cleaved Asp175)

Apoptosis

Poly

146 Nd

GeneTex

K48-ubiquitin

Ubiquitination/ proteasome

Apu2

174 Yb

Millipore Sigma

K63-ubiquitin

Endocytic trafficking and inflammation

HWA4C4

176 Yb

Biolegend

LC3B

Autophagy

Poly

171 Yb

Novus

Discovery: Alzheimer’s disease ApoE

Lipid and Aβ transport, inflammation

D6E10

167 Er

Biolegend

BIN1

Endocytic recycling

99D

147 Sm

Biolegend

CD33

Aβ clearance, microglia activation

WM53

164Dy

Biolegend

CD36

Scavenger receptor

5–271

159 Tb

Biolegend

CLU/Clusterin (aka Apo J)

Lipid and Aβ transport, inflammation

Poly18133

162 Dy

Biolegend

INPP5D (aka SHIP1) Inflammation

P1C1-A5

148 Nd

Biolegend

p-tau (Ser202, Thr205)

Hyperphosphorylated tau

AT8

158 Gd

Thermo Scientific

P-TDP-43 (Ser409/ 410)

Hyperphosphorylated DNA binding 1D3/TDP- 170 Er protein 43

Biolegend

PTK2B (aka PYK2)

Synaptic modulation

P85E2D5

149 Sm

Biolegend

T22 (tau oligomer)

Soluble tau oligomer

Poly

142 Nd

Millipore sigma

β-Amyloid, x-40

Aβ-40

29-6

168 Er

Biolegend

β-Amyloid, x-42

Aβ-42

BA3-9

161 Dy

Biolegend

8. Freezing container such as CoolCell™ FTS30 freezing container (Corning) or Mr. Frosty™ (Thermo Fisher). 9. Glass/Teflon homogenizer: Glass Potter-Elvehjem tissue grinder with smooth polytetrafluoroethylene (PTFE) pestle, clearance 0.1–0.15 mm, (Wheaton). 10. Two microcentrifuges, preferably refrigerated. 11. 37  C water bath with floating tube rack. 12. Eurostar 100 stirrer/homogenizer machine (Ika).

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1. FlowJo® software (FlowJo, Ashland, Oregon). 2. R and RStudio software. 3. SPADE, ViSNE, Citrus packages available from GitHub or accessible through cloud-based machine learning platform such as Cytobank.

2.4 Brain Tissue/Biological Material

3

Follow all biosafety and hazardous waste guidelines as per institutional guidelines (see Note 7). Mouse tissue should be collected following appropriate protocols approved by the Institutional Animal Care and Use Committee (IACUC). Whole mouse brain and/or dissected brain regions can be used as appropriate. Human brain tissue must be collected following appropriate protocols approved by the Institutional Review Board (IRB) at your institution.

Methods Perform all procedures on ice unless otherwise specified. For steps involving centrifugation at room temperature (RT), set the centrifuge to 25  C to avoid inadvertent heating of samples.

3.1 Synaptosome Preparation

1. Obtain brain tissue at a human autopsy or immediately after sacrifice of a mouse. The tissue should be ~0.5 cm3. Collected tissue must be kept on ice and processed rapidly (Fig. 1; see Note 7). 2. Finely mince tissue using a scalpel on a pre-chilled surface, such as an inverted cell culture dish on ice. Fully minced tissue will look like a slurry and no lumps of tissue should be visible. This is necessary to ensure complete and even cryopreservation. If you intend to proceed immediately with synaptosome preparation, skip to step 5 below. 3. Use the blade to transfer minced tissue to 1.5-ml cryo-tube tube containing 1 ml of RT cryopreservation solution. Mix by brief manual shaking and immediately insert the tubes into the freezing container, which should be at RT. Place the freezing container in a 80  C freezer. After overnight storage (~12 h), the samples may be transferred to liquid nitrogen or kept at 80  C (see Notes 8 and 9). 4. When resuming, transfer the stored samples rapidly to a 37  C water bath. Once completely thawed, add ice-cold HB to near the top of the tube. Samples should be spun at 4000  g for 4 min at 4  C. Gently pour off the supernatant and resuspend in 1 ml of HB. Repeat this wash step twice more to remove fully the cryopreservation solution.

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5. Transfer tissue from step 4 or from step 2 into a pre-chilled glass/Teflon size homogenizer. Based on the weight, add 10 volumes of ice-cold HB and homogenize at 800 rpm with 8 up-and-down strokes over ice. 6. Transfer the homogenate to pre-chilled 2-ml centrifuge tubes and spin at 1000  g at 4  C for 10 min to remove nuclei and cell debris. The synaptosome-containing supernatant (S1; see Fig. 1) should then be transferred (by pipetting) into new pre-chilled centrifuge tubes. The pellet (P1) may be discarded. 7. Centrifuge S1 at 10,000  g for 20 min at 4  C to obtain the crude synaptosome pellet (P2). The supernatant (S2) can be discarded. 8. Resuspend P2 in 1 ml of cryopreservation solution. These samples can again be cryopreserved by cooling slowly as described in step 3 above (see Notes 8 and 9) and stored in liquid nitrogen, or at 80  C (see Note 7). Store P2 prepared from a starting tissue size of approximately 0.5 cm3 in 5 tubes of 200 μl each (1 ml is divided in 5 tubes); if starting with more material scale up proportionally. 3.2 Modular Design and Implementation of Synaptic Antibody Panel 3.2.1 Selecting the Antibody Panel

Choose antibodies for the panel to reflect specific research interests (see Note 10). See Table 1 for specifics of our published panel (see [27]), which can be broadly categorized as follows: 1. Cell-type markers—use to identify the cellular origin of events detected by the mass cytometer, as well as interactions between adjacent cellular structures, and to detect homogenate debris. 2. Synapse type—use to subclassify individual particles (also see Note 11). 3. Nuclear controls (negative markers)—use to exclude nuclear fragments that can be distinguished by high signal from DNA intercalators: 4. Discovery markers—use to characterize synaptosomes for neurodegenerative research:

3.2.2 Optimizing Antibodies

1. Perform systematic QC on all conjugated antibodies (see Note 12 and [27]). 2. Use stable, non-radioactive lanthanide series metals and conjugation reagents [17, 28, 29]. 3. Because some metal conjugates are known to provide stronger signal than others in CyTOF, conjugate these with antibodies against targets with low expression. 4. If necessary, perform pilot experiments to guide pairing of antibodies with metals.

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1. Make sure antibodies are carrier protein-free and 100 μg in quantity to be compatible for commercially available conjugation kits (see Note 13). 2. Use 50–100 μg of purified antibody per reaction of conjugation (see Note 14). 3. For antibody conjugation with heavy metals, use the Maxpar Antibody Labeling Kit (Fluidigm) and follow the manufacturer’s protocol (see Note 14).

3.3 Staining Synaptosomes for Mass Synaptometry

See Note 15 before proceeding with this multi-day protocol. 1. At the start of day one, transfer frozen P2 sample(s) quickly to a 37  C water bath. Once thawed, top off each tube with KRP buffer, if necessary. Wash at 25  C and resuspend the pellet in KRP buffer. Wash twice more with KRP buffer and resuspend the pellet in 800 μl of KRP buffer. 2. Pre-wet a 5-μm Ultrafree-CL-centrifuge filter with 200 μl of KRP buffer. Add the resuspended pellet and spin at 1000  g at 25  C for 4 min. Discard filter and gently transfer the ~1 ml of filtrate into a fresh centrifuge tube (see Note 16). 3. Spin the filtrate at 8000  g at 25  C for 4 min. Discard the supernatant. Resuspend (by pipetting) the synaptosome pellet in 190 μl of KRP buffer. 4. Add 10 μl of Benzonase Nuclease HC (250 U/μl), gently mix and incubate for 30 min at 37  C (see Note 17). 5. Wash three times with 1 ml PBS, then spin at 8000  g at 25  C for 4 min. 6. To the pellet, add 1 ml of 1 Fix I Buffer in PBS. Mix well by pipetting gently up and down to resuspend the synaptosomes and incubate for 15 min at RT. Do not decrease the volume of fixative or increase the fixative concentration or fixation time. Also do not incubate this step on ice. 7. Wash the fixed synaptosomes twice with 1 ml PBS by centrifuging at 8000  g, at 25  C for 4 min. Optional Pause point: Store fixed synaptosomes overnight in 200 μl PBS at 4  C. This pause may be advisable if multiple samples are to be staining concurrently (see Notes 6 and 18). 8. At the start of day two, wash synaptosomes twice with 1 ml of 1 Perm-S Buffer prepared in PBS. 9. Resuspend in 1 ml 1 Perm-S Buffer and incubate at RT for 30 min (see Note 19). 10. Spin at 8000  g at RT for 4 min and remove buffer. Wash once with 1 ml of Cell Staining Buffer.

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11. Dissociate the pellet by pipetting up and down in 500 μl of Cell Staining Buffer. Keep synaptosomes on ice while preparing cocktail of antibodies. 12. Make up a cocktail of antibodies from stocks of individually conjugated labeled antibodies stored in separate tubes. Keep antibodies on ice at all times (see Note 20 and Subheading 3.2). 13. Using a 0.1 μm centrifugal filter, pre-wetted with 10 μl of Cell Staining Buffer, filter the antibody cocktail, by centrifuging at 1000  g, for 2 min, at RT. 14. To synaptosomes from step 11, add an additional 500 μl of Cell Staining Buffer. Spin at 8000  g at 4  C for 4 min and discard supernatant (Critical observation: At this point, ensure that the pellet size is approximately equal in all tubes (see Note 18)). 15. Add filtered antibody cocktail from step 13 to the pellet and resuspend by adding Cell Staining Buffer to a final volume of 150 μl (see Note 21). 16. Incubate at RT for 1.5 h, with occasional gentle mixing. 17. Wash twice with Cell Staining Buffer, followed by two washes with 1 ml PBS (by spinning at 8000  g at RT for 4 min). 18. Fix synaptosomes by resuspending in 990 μl of freshly prepared 1% PFA and incubating at 4  C, overnight (see Note 22). 19. At the start of day three, place tubes on ice and add Cell-ID intercalator-Ir-125 μM to final concentration of 125 nM, 10 min on ice (i.e., add 10 μl of 100x in 990 μl, 12 μl 100 stored as aliquots at 20  C). 20. Spin at 8000  g at 4  C for 4 min to remove fixative and wash once with ice-cold PBS. Optional Pause point: freeze samples for future run (see Note 23). 21. Wash three times with ice-cold MilliQ water (8000  g, 6 min, 4  C). Be careful not to lose the synaptosome pellet during this wash. The pellet becomes increasingly transparent or glassy. It is normal for the pellet to become smaller in size at this stage; however, it is important to ensure that the pellet is not lost completely. 22. Resuspend the synaptosome pellet in 20 μl MilliQ and store on ice until ready to run on the mass cytometer. 23. In a separate 50-ml tube on ice, dilute EQ four element calibration Beads (Fluidigm) in MQ water, 1:5 dilution. 24. Take a new 5-ml polystyrene round-bottom tube with cellstrainer cap (Falcon/Corning). Prepare tube for loading on the mass cytometer by diluting approximately 2 μl of synaptosome sample (from step 22) with 3 ml of MQ water containing 2 EQ four element calibration beads (from step 23).

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3.4 Synaptosome Acquisition Parameters and Processing 3.4.1 Acquisition Setting for Synaptosomes

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1. Adjust sample with MQ water containing 2 EQ four element calibration beads (Fluidigm) to achieve an acquisition rate of approximately 100–400 events per second. 2. Perform initiation and tuning of the mass cytometer according to manufacturer’s and your facility’s recommendations. 3. Advanced settings used in the acquisition of synaptosomes are minimum event length (10); maximum event length (100) and lower convolution threshold (200) (Fig. 2). 4. Acquire data and save in both. FCS and. IMD formats (see Note 24).

3.4.2 Post-Acquisition Normalization and Gating

1. Normalize mass synaptometry data to beads using Fluidigm CyTOF software (version 6.7.1014 or later) (see Fig. 3). 2. Bead-normalized data can be used further for analysis. Depending on downstream processing, these multi parametric and high-dimensional data can be saved as. FCS or converted to .txt (which can be opened and saved as .csv or .xls as well). 3. Initial gating and analysis should be done using FlowJo- or CytoBank, a cloud-based computational and machine learning platform with advanced tools and functionality such as SPADE, viSNE, Citrus, and FlowSOM. 4. For synaptosomes, first gate out normalizing beads and DNAhi (nuclear) particles; second gate out non-neuronal events using a CD11b and MBP negative gate; third select for neuronal events using CD56 or CD298; fourth select for presynaptic events using CD47 and SNAP25; fifth gate out “snowmen” using PSD95 (see Notes 11, 25, and 26).

3.5 Advanced Visualization and Analysis Tools for High-Dimensional Mass Synaptometry Datasets

Due to the high-dimensional nature of data generated by CyTOF, it is often necessary to apply computational tools, statistical tools, and algorithms, such as principal component analysis (PCA), SPADE, viSNE, and Citrus (see Note 27). Example analysis steps for DAT+ events (see [27]) are presented here. 1. Carefully consider the number of input events, number of nodes, and the number of markers used for clustering to generate SPADE. 2. Empirically test by changing node size (e.g., from 3 to 500, for proof of principle demonstration for mass synaptometry (see Note 28)). 3. Select ~50 nodes for 30,000 events per sample to obtain optimal visualization of DAT + events in caudate. 4. Export events from the DAT + cluster to perform in-depth analysis and interrogation of further marker changes in this cluster.

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Fig. 2 Acquisition parameters setting for mass synaptometry. (A) Screenshot of CyTOF software indicating specific setting in “acquire” mode (a) under “experimental manager” tab (b) antibody panel and other necessary details are entered. Under “advanced” (c) setting SynTOF-specific parameters are entered (Lower convolution threshold: 200, Min event duration: 10, and Max event duration:100) (d). (B) Under “acquire” mode while recording events ensure to run the sample at optimal dilution so as to acquire 100 to 400 events/second (c). A representative typical screen view of dashboard while running sample showing rain plot, status panel, and plot viewer (c). All other parameters are used as default

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Fig. 3 Processing parameters setting for mass synaptometry. (A) Screenshot of CyTOF software indicating specific setting in “Process” mode (a) with “FCS processing” (b) selection highlighting “Time Interval Normalization” as 250 s (c). (B) Under “Process” (a) mode with “IMD processing” selection (b) selection shows representative parameters that may be modified to generate a new .FCS file from original .IMD files. All other parameters are used as default

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Notes 1. Reagents and disposables: All solutions for mass cytometry should be prepared with ultrapure water. Whenever possible, avoid reagent contact with glass or metal as they are common sources of heavy metal contamination. Please note that standard laboratory dishwashers, autoclaves, and dishwashing detergents are frequently contaminated with heavy metals (particularly barium), which can disrupt experiments or damage the mass cytometer. Use of disposable plasticware is strongly recommended. Use only mass spectroscopy-grade chemicals. Reagents can be tested for heavy metal contamination by running them in “solution mode” on the mass cytometer. In addition, some solutions need to be freshly prepared: dilutions of fixative buffers (Maxpar Fix 1 or 1% paraformaldehyde (PFA)); Cell ID™ Intercalator-Ir and Perm Buffer; and Benzonase. Care should be used in handling Maxpar Fix I Buffer PFA, as they are toxic; appropriate protective measures (e.g., gloves, eye and respiratory protection) should be employed when needed. Follow all waste handling regulations when disposing of waste materials. 2. The Fluidigm MaxPar® conjugation kit provides a choice of 35 different isotopically enriched 50 mM metal salt solutions ranging from 141 praseodymium to 176 ytterbium using DN3 polymers. 3. Samples should be filtered through a 35 μm (or smaller) cell strainer directly before running on the mass cytometer. This minimizes the risk of clump formation and machine clogging. 4. Composition of homogenization buffer (HB): 0.32 M sucrose in 10 mM Tris–HCl buffer (pH 7.4) with EDTA-free, mass spectroscopy-grade protease and phosphatase inhibitor cocktail (Sigma MSSAFE). 1 HB is made by mixing 20 ml of 0.32 M sucrose in 10 mM Tris–HCl buffer (pH 7.4) to 1 vial of MSSAFE. HB must be made fresh on the day of the experiment. We prepare 0.32 M sucrose in 10 mM Tris–HCl buffer (pH 7.4) in bulk, 0.22 μm filter and store as 50-ml aliquots at 20  C. 5. The description in this section is based on a Helios model mass cytometer (Fluidigm) running software version 7.0.5189.0. Older models (e.g., CyTOF1 and CyTOF2) are suitable but have different mass responsiveness and sample injection rates and may have different sensitivity and software steps. 6. During our optimization and search for a practical solution to achieve equal counts across samples, we tested the following: a Sysmex XN-1000 hematology Analyzer (Sysmex), TC20™ automated cell counter (BioRad), Moxi Flow MXF001, a

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flow cytometer and cell count/volume Analyzer (Orflo technologies), qNano-nano and micro particles measurement using Tunable Resistive Pulse Sensing (TRPS) technology (Izon Science), and a Countess II FL Automated Cell Counter (Thermo Invitrogen). We also used protein measurement by BCA protein assay before fixation (Pierce/Thermo scientific). We found the Moxiflow counter to be the most pragmatic solution for standardizing counts, especially when measurements performed at diluted concentrations. Also see Note 18. 7. Whole mouse brain or specific brain regions may be used. For testing the method, we recommend using whole mouse brain. Human brain samples should be obtained ideally with a postmortem interval < 8 h. Human brain regions (cerebral cortex, caudate nucleus, hippocampus, and cerebellum) of approximately 0.5 cm3 in size can be collected. Since specific mouse brain regions are relatively small, tissue can be pooled from multiple mice. Please note that P2 derived from approximately 0.5 cm3 of starting tissue material will typically be aliquoted into 5 tubes for cryopreservation. 8. Brain tissue that has been snap-frozen or fixed will not work. 9. This is applicable for preservation of both collected tissue and prepared synaptosomes, as well as shipping them to other locations. (a) Larger freezing containers like CoolCell™ FTS30 accommodate 30 centrifuge tubes, while freezing containers like Mr. Frosty™ Freezing Container, which require isopropyl alcohol, usually accommodate 12–18 tubes. In either case follow the manufacturer’s instructions. It is critical that the freezing container is equilibrated at RT and then gradually cooled with samples in it ( 1  C/min to 80  C). (b) Do not transfer a sample tube directly in 80  C, liquid nitrogen or dry ice as this will compromise sample integrity. Do not use a pre-chilled freezing container or put tubes in a freezing container which are already at 80  C. (c) If bulk tissues are collected in large tubes (e.g., 5-, 15- or 50-ml tubes that do not fit standard freezing container for cryopreservation), then wrap tubes in paper towels, place in a small Styrofoam box, which should then be taped at all joints to seal completely. Place this container in 80  C overnight. (d) These frozen tissues or synaptosome samples can be shipped on dry ice. 10. We designed and optimized a SynTOF panel of antibodies (see [27]) that should be widely applicable. Our chief intention,

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however, was to develop a protocol that would be most useful for research of neurodegenerative conditions. This modular design approach should serve as a guide for modifying the panel; specifically optimized techniques or procedures which affect panel performance are noted individually below at the appropriate experimental steps. See Table 1, for selection of individual components and epitopes of the SynToF panel. 11. PSD-95 permits the detection of “snowmen,” i.e., the interaction of presynaptic and post-synaptic elements. In our experience, around 10% of the final synaptosome preparation consists of snowmen. 12. Quality control (QC) of conjugated antibody and SynTOF panel: We undertook four QC steps to ensure the sensitivity and specificity of our SynTOF panel. First, only antibodies with Western blot banding pattern unaltered by conjugation on positive samples were included. Second, we used metal minus (MM) controls, in which synaptosomes were incubated without conjugated antibody to determine background signal in the omitted channel; MM controls also were used to set gates. Third, isotype antibodies were obtained from the same supplier as the primary antibody whenever possible, always from the same host species and Ig subclass, and conjugated with the same metal ion and used at the same concentration as the primary antibody conjugate. Any signal overlapping with isotype control was excluded as non-specific. Lastly, if applicable for the antibody, we use regional variation in synaptic innervation between regions of brain or difference in presence of pathological markers in synaptosome/tissue from pathologically defined versus normal brain. 13. Some antibodies were also custom ordered from vendors or processed by us to remove carrier protein. 14. We scale up conjugation by parallel multiple conjugation reactions—never exceeding 100 μg of antibody per reaction. We modified the last step and elute with a smaller volume (10–40 μl) of W-buffer to achieve a higher titer of conjugated antibodies. 15. This is a multi-day protocol. For convenience, optional pause points are given where possible; it is not advisable to stop at other steps in the protocol. The protocol includes multiple washing steps. Each wash step includes adding 1 ml of appropriate buffer or solution, spinning at 8000  g for 4 min at the indicated temperature, and carefully discarding the resulting supernatant. Following each wash, the pellet is resuspended in the appropriate solution by gently pipetting up and down several times, without causing bubbles.

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16. This 5 μm filtration is critical. For reproducibility we use 5 μm Ultrafree-CL-centrifugal filter devices, spun at 1000  g, at 25  C, for 4 min. Do not use disc filter with manual syringes; we have found it difficult to maintain constant pressure across samples and it results in considerable sample loss. If liquid remains above the centrifugal filter after spinning, it is likely that the starting P2 was too large or that there is non-homogenized tissue debris carry-over. Consider using a smaller starting pellet or using multiple filters to avoid clogging. Do not exceed 1000  g speed. 17. Use high purity, protease-free, pan-nuclease (Benzonase® Nuclease HC, Purity >99%). This is important to reduce viscosity, preventing multiple synaptosomes flocculating together. 18. Fixed synaptosomes may be divided into multiple tubes depending on experimental plans. Some examples include: (1) titrating antibodies to determine optimal signal; (2) testing different antibody clones with the same metal mass; (3) testing antibody signal with metal minus and isotype controls; and (4) combining different panel antibodies with negative controls. Note that dividing synaptosomes from a single tube into multiple tubes (by volume) should yield an equivalent quantity of synaptosomes. When working with multiple unique samples, it is important to equalize the quantity in each tube (see Note 6). 19. Incubation of synaptosomes with 1 Perm-S Buffer, at RT, for 30 min incubation is critical for non-surface/intra synaptosomal antibody staining to work. We tried permeabilization with cold methanol, triton, and/or tween. However, we found from our empirical testing that, while these steps improve the staining of some antibodies, others are adversely affected. We have found using 1 Perm-S Buffer, at RT, for 30 min to be optimal for the given panel. We also attempted to use chilled methanol for permeabilization, as this method is frequently used in the preparation of peripheral blood monocytes for CyTOF. However, we found that this method leads to considerable synaptosome loss. 20. One can make a “master mix” antibody cocktail to divide them equally, wherever possible, and add additional antibodies in required tubes. Please note that practical volumes may differ slightly from the theoretical total of antibody volume, due to pipetting loss in handling small volumes from multiple tubes. It is a good idea to make an antibody master mix which can be aliquoted and stored in 80  C/LN2 for ready to use. This approach will save time and avoid variability associated with antibody and cocktail preparation (see [30]).

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21. Most of the literature suggests the antibody cocktail be added to sample and then add a cell staining quantity sufficient to 100 μl, which makes it easy to calculate final antibody concentration. However, we practically find that when using in-house conjugated and titrated antibodies for synaptic or neurobiology panels (such as our SynTOF panel), the optimal cocktail volume reaches around 70–90 μl at times, even after using a high titer antibody. Therefore, we add Cell Staining Buffer to make final volume to 150 μl. Whatever volume you use, keep this volume constant for given set of samples in experiments where comparisons are intended. 22. Make fresh 1% PFA before each use from new ampoule of 16% PFA (Alfa Aesar). Be careful not to lose the synaptosomes following fixation as the pellet can loosen from the wall and wash off. Avoid decanting supernatants by inverting the tube. It is best to use filter pipette tips to remove buffers from each tube for every following step. Mix to suspend stained synaptosomes by pipetting several times. Do not decrease the volume of fixative below 500 μl or increase the concentration of fixative as it may lead to crowding of synaptosomes that would lead to increase in doublets. Immediately transfer tubes at 4  C, do not keep at RT. 23. At this point, it is possible to freeze the samples for a future run on another day. After the wash in Step 20, wash two more times with ice-cold PBS. Mix gently with 1 ml of 10% DMSO in FBS (make multiple aliquots if intend to run multiple times, multiple locations). Immediately place in freezing box and transfer samples to a 80  C freezer, to cool the sample at a rate of 1  C/min, starting at RT (see Note 9). When ready to run, thaw sample quickly at 37  C, add MQ water to make volume 1 ml. Spin 8000  g at 4  C for 4 min to remove supernatant and resume at step 21 (see [31]). 24. Data are collected by default as an .IMD (integrated mass data) file, which generates multiple .FCS (flow cytometry standard) files at the end of the acquisition. It is advisable to save the . IMD file, because it contains all the original data, which can be reprocessed later to generate additional .FCS files after modifying acquisition parameters such as minimum event length, maximum event length, and lower convolution threshold. This post-acquisition processing can be done away from the mass cytometer on computers with CyTOF software. 25. Third and fourth gating filters can be combined to use CD56 and SNAP25 to select neuronal presynaptic events in one step. This may be required if one wants to create space in the panel for antibody of interest. Please note that standard steps that are used for single-cell analysis used in immuno-oncology were

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modified and adapted for synaptosome data processing. The gating strategy was adapted and aligned with the panel design for analysis of identification of bonafide synapses using multiple markers. 26. Practically, CD56 and SNAP25 will be sufficient to gate neuronal pre-synaptic events in one-step gating. 27. In principle, data generated is somewhat equivalent to data generated by conventional fluorescence-based flow cytometry, however with far more (30 to 40) analytes detected simultaneously, and without the need for compensation or spectral overlapping. 28. Choosing fewer nodes leads to insufficient visualization of the diversity of synaptic events, whereas choosing too many nodes for a given number of input events leads to unnecessarily complex trees with near empty nodes.

Acknowledgments This work was supported by grants from the NIH: P50 NS062684, P50 AG047366, RF1 AG053959, R01 AG056287, R01AG057915, DP2 EB024246, and P50 AG005136 (CDK), and by the Nancy and Buster Alvord Endowment. We thank M. Holden and M. Leipold of the Stanford Human Immune Monitoring Core for their assistance and guidance, A. Beller from the Department of Pathology, University of Washington for administrative support, and E. Fox from Department of Pathology, Stanford University for helpful discussions. References 1. Braak H, Braak E (1991) Neuropathological stageing of Alzheimer-related changes. Acta Neuropathol 82:239–259 2. Wishart TM, Parson SH, Gillingwater TH (2006) Synaptic vulnerability in neurodegenerative disease. J Neuropathol Exp Neurol 65: 733–739 3. Scott DA, Tabarean I, Tang Y et al (2010) A pathologic cascade leading to synaptic dysfunction in alpha-synuclein-induced neurodegeneration. J Neurosci 30:8083–8095 4. Bellucci A, Zaltieri M, Navarria L et al (2012) From alpha-synuclein to synaptic dysfunctions: new insights into the pathophysiology of Parkinson’s disease. Brain Res 1476:183–202 5. Overk CR, Masliah E (2014) Pathogenesis of synaptic degeneration in Alzheimer’s disease and Lewy body disease. Biochem Pharmacol 88:508–516

6. McKeith IG, Boeve BF, Dickson DW et al (2017) Diagnosis and management of dementia with Lewy bodies: fourth consensus report of the DLB consortium. Neurology 89: 88–100 7. Duyckaerts C, Potier MC, Delatour B (2008) Alzheimer disease models and human neuropathology: similarities and differences. Acta Neuropathol 115:5–38 8. Jucker M (2010) The benefits and limitations of animal models for translational research in neurodegenerative diseases. Nat Med 16: 1210–1214 9. Galli S, Lopes DM, Ammari R et al (2014) Deficient Wnt signalling triggers striatal synaptic degeneration and impaired motor behaviour in adult mice. Nat Commun 5:4992 10. Zhu H, Yan H, Tang N et al (2017) Impairments of spatial memory in an Alzheimer’s

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disease model via degeneration of hippocampal cholinergic synapses. Nat Commun 8:1676 11. Dietrich K, Bouter Y, Muller M et al (2018) Synaptic alterations in mouse models for Alzheimer disease-a special focus on N-truncated Abeta 4-42. Molecules 23:718 12. Sokolow S, Henkins KM, Williams IA et al (2012) Isolation of synaptic terminals from Alzheimer’s disease cortex. Cytometry A 81: 248–254 13. Postupna NO, Keene CD, Latimer C et al (2014) Flow cytometry analysis of synaptosomes from post-mortem human brain reveals changes specific to Lewy body and Alzheimer’s disease. Lab Investig 94:1161–1172 14. Postupna N, Latimer CS, Larson EB et al (2017) Human striatal dopaminergic and regional serotonergic synaptic degeneration with Lewy body disease and inheritance of APOE epsilon4. Am J Pathol 187:884–895 15. Bilousova T, Miller CA, Poon WW et al (2016) Synaptic amyloid-beta oligomers precede p-tau and differentiate high pathology control cases. Am J Pathol 186:185–198 16. Gylys KH, Bilousova T (2017) Flow cytometry analysis and quantitative characterization of tau in Synaptosomes from Alzheimer’s disease brains. Methods Mol Biol 1523:273–284 17. Bendall SC, Simonds EF, Qiu P et al (2011) Single-cell mass cytometry of differential immune and drug responses across a human hematopoietic continuum. Science 332: 687–696 18. Simmons AJ, Banerjee A, McKinley ET et al (2015) Cytometry-based single-cell analysis of intact epithelial signaling reveals MAPK activation divergent from TNF-alpha-induced apoptosis in vivo. Mol Syst Biol 11:835 19. Spitzer MH, Gherardini PF, Fragiadakis GK et al (2015) IMMUNOLOGY. An interactive reference framework for modeling a dynamic immune system. Science 349:1259425 20. Spitzer MH, Nolan GP (2016) Mass cytometry: single cells, many features. Cell 165: 780–791

21. Wong MT, Chen J, Narayanan S et al (2015) Mapping the diversity of follicular helper T cells in human blood and tonsils using highdimensional mass cytometry analysis. Cell Rep 11:1822–1833 22. Chevrier S, Levine JH, ZaNotelli VRT et al (2017) An immune atlas of clear cell renal cell carcinoma. Cell 169(736–49):e18 23. Hamers AAJ, Thomas GD, Kim C et al (2017) Diversity of human monocyte subsets revealed by CyTOF mass cytometry. J Immunol 198 (1 Supplement):208.5 24. Korin B, Ben-Shaanan TL, Schiller M et al (2017) High-dimensional, single-cell characterization of the brain’s immune compartment. Nat Neurosci 20:1300–1309 25. Lavin Y, Kobayashi S, Leader A et al (2017) Innate immune landscape in early lung adenocarcinoma by paired single-cell analyses. Cell 169:750–765.e17 26. Mrdjen D, Pavlovic A, Hartmann FJ et al (2018) High-dimensional single-cell mapping of central nervous system immune cells reveals distinct myeloid subsets in health, aging, and disease. Immunity 48(380–95):e6 27. Gajera CR, Fernandez R, Postupna N et al (2019) Mass synaptometry: high-dimensional multi parametric assay for single synapses. J Neurosci Methods 312:73–83 28. Bandura DR, Baranov VI, Ornatsky OI et al (2009) Mass cytometry: technique for real time single cell multitarget immunoassay based on inductively coupled plasma time-of-flight mass spectrometry. Anal Chem 81:6813–6822 29. Bendall SC, Nolan GP, Roederer M et al (2012) A deep profiler’s guide to cytometry. Trends Immunol 33:323–332 30. Schulz AR, Baumgart S, Schulze J et al (2019) Stabilizing antibody cocktails for mass cytometry. Cytometry A 95:910–916 31. Sumatoh HR, Teng KW, Cheng Y et al (2017) Optimization of mass cytometry sample cryopreservation after staining. Cytometry A 91A:48–61

Chapter 7 A Guide to Analysis of Relative Synaptic Protein Abundance by Quantitative Fluorescent Western Blotting Heidi K. Mu¨ller Abstract The introduction of fluorescent detection systems has revolutionized the applicability of Western blotting for quantitative protein expression analyses. The fundamental premise behind fluorescent Western blotting is the combination of distinct fluorescent dye-conjugated secondary antibodies and high performance digital imaging solutions in which the fluorescence signal is directly proportional to the amount of protein enabling quantitative measurements and simultaneous detection of several target proteins. This aspect of Western blotting is now widely used, especially in preclinical research, to detect quantitative changes in protein levels and phosphorylation status between experimental groups. This chapter provides a detailed step-by-step guide for best practice procedures during the entire process from sample preparation, SDS polyacrylamide gel electrophoresis to electrotransfer of proteins and highlights approaches that can be applied to increase data output. Key words Fluorescent Western Blotting, Multiplexing, Phosphoprotein detection, Nitrocellulose membrane, Synaptic proteins, Best practice, Quantitative, Normalization

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Introduction The basic principles behind Western blotting have not changed considerably since the technique was introduced 40 years ago [1, 2] and it is still one of the most commonly used methods in all aspects of life-science research as well as an important clinical immunodiagnostic tool. The essential steps of Western blotting involves separation of proteins in a complex sample according to their sizes by gel electrophoresis followed by electrotransfer of the separated proteins to a membrane for subsequent detection of proteins of interest using target specific antibodies. However, with the recent development of fluorescent Western blotting that combines fluorescent-dye conjugated secondary antibodies with digital imaging systems, the technique now offers several important

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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advances over traditional detection methods. This includes improved sensitivity, a greater range of linear detection, excellent stability, and the ability to distinguish multiple fluorophores for simultaneous detection of several target proteins without the need to strip and re-probe the blot. These features have provided a fundamental shift in signal acquisition that enables accurate and quantitative results. Changes in expression of synaptic proteins are intimately related to changes in almost all brain functions and expression abnormalities often precede the onset of neurological disorders. Using quantitative fluorescent Western blotting it is possible to discover even subtle changes in synaptic protein abundance in unpurified or enriched samples as a function of treatment, time, developmental stage, brain region, genotype, pathology, etc. This chapter describes, in detail, an optimized protocol on how to process samples prior to loading on the gel, the procedures for SDS polyacrylamide gel electrophoresis, handling of gels for optimal transfer, processing of membranes for increased data output and highlights important considerations to keep in mind when performing quantitative analyses.

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Materials Prepare all solutions using ultrapure water and analytical grade reagents. Store buffers at room temperature unless otherwise stated. Use powder-free gloves at all times.

2.1 Sample Preparation

1. Protein Assay Kit for measurement of total protein. 2. Lysis/dilution buffer compatible with the buffer used for protein extraction. 3. SDS sample buffer: Ready-to-use concentrated SDS sample loading buffers are available commercially. We routinely use homemade 3 Laemmli sample buffer (187.5 mM Tris-HCl, pH 6.8, 6% SDS, 30% glycerol, 0.03% bromophenol blue, and 200 mM DTT [added just before use]). 4. Water bath or tube block heater.

2.2 SDS Polyacrylamide Gel Electrophoresis

1. SDS Polyacrylamide Gels: Use precast polyacrylamide gels for optimal consistency and reproducibility (e.g., Criterion TGX Precast Gels, Bio-Rad or NuPAGE Bis-Tris Precast Gels, Invitrogen) (see Note 1). 2. Running Buffer: For precast gels, follow the manufacturer’s recommendations for optimal choice of electrophoresis running buffer.

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3. Prestained protein molecular weight marker: Use any commercially available prestained protein ladder covering the desired molecular weights. Dual or tricolor options are preferred for easy identification. Alternatively, choose a molecular weight marker that is specifically designed for fluorescent Western blotting. 4. Vertical electrophoresis chamber (e.g., Mini-PROTEAN Tetra/Midi Criterion-Vertical Electrophoresis Cell, Bio-Rad or XCell SureLock Mini/XCell4 SureLock Midi-Cell Electrophoresis System, Invitrogen). 5. Syringe and needle (needle diameter 0.6–0.8 mm). 6. Spatula. 2.3 Electrotransfer of Proteins

1. Nitrocellulose transfer membrane: Nitrocellulose membranes are commercially available from various vendors in ready-to-use preassembled transfer packs (e.g., Trans-Blot Turbo Transfer Pack, Bio-Rad or iBlot 2 Transfer Stacks) (see Note 2). 2. Transfer unit: Use a blotting system compatible with the choice of transfer pack (e.g., Trans-Blot Turbo Transfer System, Bio-Rad or iBlot 2 Dry Blotting System, Invitrogen. 3. Incubation trays (see Note 3). 4. 1 Tris-Buffered Saline (TBS) (50 mM Tris–HCl, 150 mM NaCl, pH 7.6) (see Note 4). 5. 20% Tween-20: Prepare a 20% stock solution by mixing 20 ml of Tween-20 with 80 ml of water by stirring. 6. Blocking solution: Use Intercept (TBS) Blocking Buffer, LI-COR (store at 4  C) (see Note 5). 7. Shaking platform (optional: tube roller mixer). 8. Flat tipped forceps. 9. Blotting roller.

2.4 Other Supplies Required

1. Pipettes and pipette tips. 2. Gloves. 3. Deionized or distilled water. 4. Power supply. 5. Primary antibodies (see Note 6). 6. Fluorescently labeled secondary antibodies (see Note 7). 7. Imaging System. 8. Quantification software.

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Methods

3.1 Sample Preparation

1. Dissect the tissue or punch out your region of interest (or collect cells if dealing with cell cultures) and proceed with the protein extraction and purification method required for your analysis (see Note 8). 2. Regardless of your end product (e.g., total lysate, synaptosomes, or purified synaptic vesicles), take out a small sample volume (typically 5–10 μl) and determine total protein concentration in each sample using a protein assay (e.g., BCA, Lowry or Bradford method) that is compatible with possible detergents present in your samples. It is important to measure protein concentration prior to mixing with SDS sample buffer. 3. Adjust sample protein concentrations (or aliquots thereof) according to the protein concentration measurements to obtain equal concentrations of proteins (see Note 9). Use the (lysis)-buffer in which your samples were prepared/resuspended for diluting the samples to maintain the same ionic strength across all samples. If unknown, use TBS containing protease inhibitors (and phosphatase inhibitors when dealing with phosphorylated proteins). 4. Mix two volumes of your sample with one volume of 3 SDS sample buffer (see Note 9). 5. Heat samples for 2 min at 98  C, 10 min at 70  C or 15 min at 55  C. We find that incubation at 55  C works well for most proteins. 6. Cool down samples at room temperature for a few minutes before loading the gel.

3.2 SDS Polyacrylamide Gel Electrophoresis

1. Assemble the gel electrophoresis chamber with the desired number of gels (1–4 depending on gel chamber). Remember to remove the tape often present at the foot of precast gels. Use a dummy plate or a spare gel to close off one side of the buffer chamber when running only one gel. Add running buffer to the central compartment. Carefully remove the comb from the gel. Use a syringe with a long thin needle to carefully rinse each well with running buffer to get rid of residual gel pieces. The needle can also be used to carefully adjust slanted walls between wells. Check for leaking caused by improper assembly of the chamber before moving on. 2. Load 1–5 μl prestained molecular weight marker (optimal volume should be optimized for each fluorescent system) to any preferred well (usually the first or the last) for monitoring protein separation and for size estimation (see Note 10).

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3. Load equal volumes of protein sample into the wells. Load any unused wells with a similar volume of 1 SDS sample buffer diluted in the same (lysis)-buffer in which the protein samples were prepared. Add the remaining running buffer to the outer compartment and place the lid on the chamber. 4. Immediately connect to a power supply and start separation at 100 V. After 10 min, voltage can be increased for faster run time. 5. Run the gel until the blue dye front nearly reaches the bottom of the gel for best resolution in the low molecular weight range. Dependent on the size of the target protein(s), the dye front can also be allowed to run off the gel to improve separation of the remaining bands. 3.3 Electrotransfer of Proteins

1. Remove the gel cassette and open it carefully with a spatula. The gel will typically stick to one of the plates. Leave the gel attached to the plate while cutting off the dye front (bromophenol blue fluoresces and can cause high background) and the top gel area in which the wells reside. Before removing the gel from the plate, wet your gloves in running buffer to minimize the risk of tearing. Transfer the gel to the membrane (keep the same orientation on the membrane as on the gel) and complete assembly of the transfer pack as recommended by the manufacturer. Alternatively, cut the gel horizontally and transfer individual gel pieces onto separate membranes for different transfer settings (e.g., settings optimized for low, mixed or high molecular weight proteins) (see Fig. 1 and Note 11). Use a blotting roller to remove air bubbles between the gel and blotting membrane. Follow the Transfer Unit manufacturer’s guidelines for optimal transfer conditions. 2. When the transfer is complete, disassemble the transfer sandwich and if necessary cut one or several corners to serve as orientation marker (never use ink or markers to write on the membrane—they fluoresce and cause background). Only handle the membrane with clean forceps and avoid touching it with your hands and gloves. Carefully remove the membrane and place it in an incubation tray containing TBS to rinse off any residual gel pieces. Incubate on an orbital shaker or rocking platform for 5 min at room temperature. 3. OPTIONAL: For normalization using total protein staining, use a staining reagent that is compatible with your image system and follow the workflow recommended by the manufacturer (see also Note 12). 4. To prevent nonspecific binding of antibodies, place the membrane in the proper blocking solution and incubate for 1 h under constant gentle agitation at room temperature.

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Fig. 1 Illustration of different options for processing of gels and membranes for increased data output using multiplex analysis

5. Incubate with primary antibodies diluted in blocking buffer containing 0.1% Tween-20 (50 μl of a 20% stock solution of Tween-20 per 10 ml blocking buffer) for 2 h at room temperature or overnight at 4  C (preferable) under constant gentle agitation on an orbital shaker. To minimize the use of costly primary antibodies, use a container that is only slightly larger than the blot or use a compartmented container for membrane strips (see Fig. 1 and Note 11). Alternatively, for membrane strips use a 10 ml or 15 ml conical tube, which require less volume and incubate on a tube roller mixer.

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6. Wash the membrane 4 5 min in TBS containing 0.1% Tween20 (5 ml of a 20% stock solution of Tween-20 per 1 l TBS) by shaking vigorously on an orbital shaker or rocking platform at room temperature. 7. Incubate with secondary antibodies diluted immediately prior to use in blocking buffer containing 0.1% Tween-20 for 1 h at room temperature under constant gentle agitation. IMPORTANT : From this point, protect the membrane from light during incubation and washing steps. 8. Wash the membrane 4 5 min in TBS containing 0.1% Tween20 by shaking vigorously at room temperature. 9. Wash the membrane 2 5 min in TBS to remove residual Tween-20. 10. Capture signals on an appropriate imaging instrument (e.g., Odyssey Imaging System, LI-COR). Use scan/capture settings according to the manufacturer’s guidelines. 11. For later reprobing (i.e., detection of different targets), rinse the membrane briefly in TBS and let it dry on an absorbent material and store in foil pouches or regular photo pockets protected from light. 12. Use appropriate imaging software to quantify Western blot signals (see Note 12).

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Notes 1. The choice of gel depends on the molecular weights of your target proteins. For optimal separation, small size proteins require higher acrylamide percentage, and large size proteins require a lower acrylamide percentage. Gradient gels offer an advantage in terms of resolution over a greater range of protein sizes, thus allowing separation of both high- and low-molecular weight proteins on the same gel. However, we find that linear gels are well-suited for most applications. For instance, when studying relative abundance of synaptic proteins you are likely to have many proteins of interest and therefore will have to run several gels. In this case, you can arrange your target proteins in groups and run your samples on different linear gels with acrylamide percentages most suitable for the range of protein sizes in each group of target proteins. Refer to the protein gel migration charts of the gel manufacturer for optimal gel selection. Also, most companies offer gels in different well formats. Select the format best suited for your application. 2. Membranes have autofluorescent properties that can cause significant background when using fluorescent detection. We find nitrocellulose membranes to be optimal for detection with the LI-COR fluorescence imaging system.

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3. Contaminated containers can cause serious background. Make sure that the containers are clean and free of any traces of Coomassie staining. If possible, avoid using incubation trays and containers for other applications than fluorescent Western blotting. Use containers with lids to protect from dust and ideally, use light-protected containers for easy handling of membranes during incubation with fluorescently labeled secondary antibodies and the subsequent washes. 4. Keep your buffer system consistent throughout the protocol for washing, blocking, and incubation with antibodies. We find that the use of TBS-based solutions is generally better than PBS. Also, for detection of phosphoproteins it is recommended to avoid PBS buffers since the phosphate ion may lead to interference. 5. To reduce costs, we routinely dilute the Intercept Blocking Buffer 1:1 or 1:2 in TBS without loss of performance. When using nitrocellulose membranes, we find that 5% BSA in TBS (always prepare fresh prior to use) also works fine for most applications. 6. The selection of primary antibodies is in many cases a matter of empirical testing. It is time well spent to check customer reviews, user-based antibody blogs, and scientific papers before choosing one antibody over another. Always check species reactivity (e.g., when analyzing samples prepared from rat; the antibody should recognize the rat immunogen sequence) and that your antibody of choice is compatible with Western blotting (the antibody has to be able to recognize the linear epitope). Dual detection of total and phosphorylated protein requires antibodies to be raised in different species (for rat samples, use for instance primary antibodies from mouse and rabbit, respectively). For multiplexing in general, always perform preliminary blots with each of the individual antibodies to determine migration properties and possible background signals. It is preferable to include appropriate positive/negative controls during this optimization process. Refer to antibody datasheets for guidance on antibody concentrations. 7. Use only secondary antibodies conjugated to high quality fluorescent dyes with excitation and emission spectra compatible with the image system. Also, use only secondary antibodies cross-adsorbed against other species. 8. All steps during extraction of proteins must be carried out at 2–8  C. For simultaneous analysis of RNA and protein from the same experimental sample, use an appropriate extraction kit (e.g., Ambion’s Protein and RNA isolation System) and proceed with an aliquot of the total cell lysate [3, 4]. For preparation of total protein lysate, homogenize the tissue (using for

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instance a mini handheld homogenizer, a Dounce homogenizer, or a Precellys homogenizer) in 10% (w/v) lysis buffer (50 mM Tris–HCl, pH 7.4, 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 1 protease inhibitor cocktail), incubate for 30 min at 4  C under gently agitation, centrifuge the samples at 12,000–14,000  g for 15 min at 4  C and collect the supernatant for further analysis. Refer to the literature for the preparation of synaptosomes [5, 6] and isolation of synaptic vesicles [7]. For downstream detection of phosphorylated proteins include phosphatase inhibitors (e.g., 5 mM NaF, 1 mM Na3VO4, 5 mM Na2PO4) during all steps in the protein extraction protocols. 9. Loading the same amount of protein to all wells is critical for quantitative measurements of relative protein abundance. To ensure that differences in fluorescent signals reflects genuine biological changes, it is essential to define the linear (quantitative) range of detection for each target- and loading control protein to determine the optimal amount of protein to be loaded in each well. Consider using total protein staining for normalization if your loading control protein is not in the same linear range of detection as your target protein(s) (see also Note 12). For most applications we find that loading of 10–20 μg protein per well falls within the linear range of both our target proteins and loading control protein(s). Consequently, we typically dilute our samples to either 3 μg/μl or 2 μg/μl to obtain a final concentration of 2 μg/μl or 1.33 μg/μl after mixing with 3 SDS sample buffer. Using a loading volume of 10 μl, this equals a total sample amount of 20 μg and 13.3 μg, respectively. Adjust sample concentrations and ratio of sample:SDS sample buffer if using SDS sample buffers with different concentrations. 10. For downstream horizontal cutting of the gel or membrane (for increased data output), it is advantageous to load protein ladder in both the first and the last well to ensure optimal cutting (see Fig. 1). Hint: use different volumes of protein ladder or even different ladders in the two wells for easy recognition of gel and membrane orientation. 11. Before cutting a gel and/or membrane, always test your sample and target antibodies on a whole gel/membrane to get familiar with band patterns and possible background signals. An increasing number of journals require full versions of blots and raw data to be uploaded in supplementary information or provided during peer review. 12. Refer to the extensive literature available online that describes the use of your analysis software of choice for Western blot quantification. Regardless of analysis tool, normalization is the

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most critical step for obtaining accurate, quantitative data from Western blots. Ideally, the loading of the same amount of protein to all wells should render normalization unnecessary but possible variabilities in sample loading and transfer efficiency calls for optimal normalization strategies. Traditionally, the signals from the proteins of interest are normalized to an internal constitutively expressed control, often a so-called housekeeping protein such as β-actin, β-tubulin, and GAPDH. Housekeeping proteins are however most often expressed at very high levels and therefore not necessarily in the same linear range of detection as the target proteins (see also Note 9). It can therefore be advantageous to use a lessabundantly expressed loading protein to normalize results. In any case, when using single protein normalization it is essential to demonstrate that the loading control protein of choice does not vary across experimental conditions. An increasingly popular alternative to single protein normalization is normalization to total protein, which entails staining of the membrane after transfer of proteins but prior to blocking. Refer to the vendor’s guidelines for optimal use. References 1. Renart J, Reiser J, Stark GR (1979) Transfer of proteins from gels to diazobenzyloxymethylpaper and detection with antisera: a method for studying antibody specificity and antigen structure. Proc Natl Acad Sci U S A 76 (7):3116–3120. https://doi.org/10.1073/ pnas.76.7.3116 2. Towbin H, Staehelin T, Gordon J (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc Natl Acad Sci U S A 76(9):4350–4354. https://doi.org/10. 1073/pnas.76.9.4350 3. Elfving B, Mu¨ller HK, Oliveras I et al (2019) Differential expression of synaptic markers regulated during neurodevelopment in a rat model of schizophrenia-like behavior. Prog NeuroPsychopharmacol Biol Psychiatry 95:109669. https://doi.org/10.1016/j.pnpbp.2019. 109669

4. Mu¨ller HK, Wegener G, Popoli M et al (2011) Differential expression of synaptic proteins after chronic restraint stress in rat prefrontal cortex and hippocampus. Brain Res 1385:26–37. https://doi.org/10.1016/j.brainres.2011.02. 048 5. Messa M (2018) Preparation of Synaptosomes from mammalian brain by subcellular fractionation and gradient centrifugation. Methods Mol Biol 1847:13–22. https://doi.org/10.1007/ 978-1-4939-8719-1_2 6. Mu¨ller HK, Wegener G, Liebenberg N et al (2013) Ketamine regulates the presynaptic release machinery in the hippocampus. J Psychiatr Res 47(7):892–899. https://doi.org/10. 1016/j.jpsychires.2013.03.008 7. Ahmed S, Holt M, Riedel D et al (2013) Smallscale isolation of synaptic vesicles from mammalian brain. Nat Protoc 8(5):998–1009. https:// doi.org/10.1038/nprot.2013.053

Chapter 8 Synaptosomes and Metamodulation of Receptors Anna Pittaluga and Mario Marchi Abstract Synaptosomes are re-sealed pinched off nerve terminals that maintain all the main structural and functional features of the original structures and that are appropriate to study presynaptic events. Because of the discovery of new structural and molecular events that dictate the efficiency of transmitter release and of its receptor-mediated control in the central nervous system, the interest in this tissue preparation is continuously renewing. Most of these events have been already discussed in previous reviews, but few of them were not and deserve some comments since they could suggest new functional and possibly therapeutic considerations. Among them, the “metamodulation” of receptors represents an emerging aspect that dramatically increased the complexity of the presynaptic compartment, adding new insights to the role of presynaptic receptors as modulators of chemical synapses. Deciphering the mechanism of presynaptic metamodulation would permit indirect approaches to control the activity of presynaptic release-regulating receptors that are currently orphans of direct ligands/modulators, paving the road for the proposal of new therapeutic approaches for central neurological diseases. Key words Synaptosomes, presynaptic receptors, Transmitter release, Receptor–receptor interaction, Functional crosstalk, Receptor trafficking, NMDA receptors, AMPA receptors, GLYT1, Somatostatin receptors metamodulation

1 An Historical Overview on Synaptosomes and their Use in the Study of Neurotransmission Neurotransmission is a central and fundamental event in the functioning of the central nervous system (CNS). Indeed neurons, for their correct functioning, must be organized to communicate with each other in a coordinated and extremely precise manner. The critical steps in the transfer of information from neuron to neuron at the synaptic level are represented by the release of neurotransmitters from the presynaptic components of one neuron into the synaptic cleft. The binding of these neurotransmitters to specific structures (receptors) present onto postsynaptic neurons induces fast or slow functional changes, thus permitting the flow of information between cells in the central nervous system [1–6].

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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There are several experimental methods that allow the study of neurotransmission and the functional events related to it [7– 11]. All these experimental approaches provide important information at this regard, but some of them, due to their peculiarities, provide significant and specific information on this topic. In this regard, for example, the possibility to use subcellular fractions of neural tissue like the isolated nerve endings (synaptosomes), which contain almost exclusively intact and functionally active presynaptic nerve endings, has represented a remarkable tool for investigation and a very significant turning point [12, 13]. Synaptosomes can be easily obtained by homogenization of brain tissues, their membranes having “pinched off” at the point of connection with the neuronal axon, followed by purification from other tissue components through density-gradient centrifugation techniques [12, 13]. Since their discovery, synaptosomes have been fully characterized both from a morphological and biochemical point of view [14, 15]. After their incubation in physiological solution, synaptosomes are able to perform most of the activities of the in vivo nerve terminals from which they derive, in an organized and integrated manner. They keep inside all the microstructures and cellular organelles visible in the intact synapse in situ and if they are maintained in thermostated and appropriate physiological glucose solutions they can allow to study most of the functional activities specific of the nerve terminals in situ. An important technological milestone in this field has been represented by the introduction of the superfusion technique [16, 17] that has permitted to overcome most, if not all, of the problems linked to the use of synaptosomes under static conditions. Basically, the apparatus consists of several identical superfusion chambers having at the bottom filter holders of porous glass. Synaptosomes are plated as very thin layers on microporous filters and up-down superfused with physiological solutions. Under these experimental conditions, the transmitters released (the endogenous molecules as well as the preloaded radiolabelled ones) are removed by the superfusion fluid before they can accumulate and activate presynaptic auto- and heteroreceptors, as well as reuptake carriers, thus excluding the possibility of indirect effects. The scenario is that of a thin layer of synaptosomes having their membrane targets (transporters, receptors, etc.) virtually free of endogenously released agonists. Each of these targets can, however, be selectively activated by adding the appropriate ligand, at the desired concentration, to the superfusion medium. Another characteristic of this methodological approach is that any effect on the release of one neurotransmitter can be attributed exclusively to an action on the nerve terminal releasing that neurotransmitter. Moreover, this fact allows the unambiguous anatomical localization of a given protein (receptors, transporters, ion channels, etc.) on the nerve terminal.

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Therefore, several aspects of neurotransmission such as the synthesis, compartmentation, and release itself can be investigated with this preparation at the subcellular as well as at the molecular level [18–20]. It is important to specify that some of the mechanisms above described are not easy to study and characterize in the nervous tissue with other methodological approaches due to the obvious difficulties represented by the complexity of the neuronal interactions. As already introduced, the up-down superfusion of a thin layer of synaptosomes permits the unequivocal anatomical localization of a receptors present on the nerve terminal which play a fundamental role in the modulation of the neurotransmitter under investigation [3, 21]. Indeed at the presynaptic levels, neurotransmitter release is tightly regulated by autoreceptors that, throughout a negative feed-back mechanism, can tune the exact amount of released transmitter needed for the information to pass to postsynaptic elements [18, 19]. In addition, the release of a given transmitter is presynaptically modulated by other neurotransmitters by the activation of heteroreceptors located onto the membrane of the releasing nerve terminal, providing a mean of integration of the original signal [22]. One may therefore conclude that this preparation is the ideal method to study the receptors modulating neurotransmitter release present on synaptic terminals as well as their classic pharmacological quantification. Last but not least, an important review published in 2000 (Synaptosomes Still Viable after 25 Years of Superfusion) highlighted, with the support of significant literature, the versatility of the use of synaptosomes in superfusion, pointing out several aspects of the nerve endings function that have been previously investigated by means of this approach [9]. In particular, several aspects of synaptic neurochemistry have been investigated including the multiplicity of the carrier systems transport present on one single nerve terminal and some peculiarities of neurotransmitter release which occur through reversal of the uptake carrier [19, 20]. In the recent years the research in this field, thanks to different experimental approaches, which include electrophysiological studies, immunochemical analysis of the protein expression and content and others, has brought to a new and even more complex picture of the mechanisms of neurotransmission. Our most recent research has been focused in particular on the study of modulation of the release of neurotransmitters and we here report some new findings that concern the mechanism of “metamodulation of neuromodulation” and the “in-out trafficking of receptors” in synaptosomal plasma membranes which can be studied almost exclusively using our methodological approach. Our results might have significant implications not only with regard to the physiology of the synapse but may pave the way to further investigations to clarify the role of synaptic plasticity on aging, drug dependence, and acute or chronic pharmacological treatments.

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“Metamodulation” of Release-Regulating Presynaptic Receptors The amount of transmitter released from nerve terminals does not exclusively depend on the intensity of the stimulus applied at nerve endings, but also on other concomitant events elicited by the transmitters themselves. This neuromodulation account for the tuning of the chemical neurotransmission at the synapsis level and it is exerted acting at both auto- and/or heteroreceptors which control transmitter exocytosis. The event is pivotal to define the strength of the chemical contact and therefore of the synaptic plasticity. However, neuromodulation is insufficient to describe the complexity of the mechanisms of regulation of transmitter release. Indeed, we have to consider the presence on a single nerve terminal of a plethora of different receptors and carriers that control transmitter exocytosis not only directly through their receptors but also indirectly modulating the function of other coexisting receptors which modulate neurotransmitter release. Therefore, several receptors which are present and coexist on the same terminals may interact one to each other exerting a “modulation of neuromodulation” which is referred to as “metamodulation.” The concept of metamodulation originates from the study of Fuxe and Agnati [23] and the term was adopted by Katz and Edwards [24] to refer to the mechanism of tuning of neurotransmission involving different receptors which are colocalized on the same cells and functionally interact. Briefly, the neurotransmitter released in the synaptic cleft can modulate its own release as well as the release of other transmitters by acting on auto- or heteroreceptors, respectively. The same molecule, however, can also modulate the activity of other presynaptic release-regulating receptors colocalized on the same terminals [25]. The complexity that derives from this multi-target system is impressive but well consistent with the multiplicity of the neuronal interactions and accounts, in part, for the mechanisms of synaptic plasticity. Our experimental approach with synaptosomes in superfusion has been very useful to decipher the mechanism(s) of the “metamodulation” of presynaptic events. By using the “up-down superfusion of a thin layer of synaptosomes” we demonstrated the coexistence of distinct receptors on the same structure and investigated the impact of their functional crosstalk that drives the terminals activities.

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3 Metamodulation of Presynaptic Release-Regulating Receptors: The Case of Presynaptic Release-Regulating NMDA Receptors N-methyl-D-aspartate (NMDA) receptors are ionotropic glutamate receptors that mediate most of the excitatory glutamatergic signaling in the central nervous system (CNS). They preferentially adopt a tetrameric subunit assembly and associate to a voltageoperated ionic channel permeant to mono- and divalent cations, sterically blocked by Mg2+ ions in resting physiological conditions. The receptor consists of GluN1 and GluN3 subunits (which bear the glycine binding site) and of GluN2 subunit (that possess the glutamate binding site). In the presence of physiological Mg2+ ions in the biophase, the associated ionic channel is sterically blocked and the in-flowing of the positive charges minimized, a condition that impedes the agonist-induced activation of the receptor. Accordingly, starting from the early 1980s, most of the studies dedicated to quantify the releasing activity of NMDA receptors in synaptosomes were carried out in non-physiological low [Mg2+]out in order to assure the NMDA receptor-mediated signal [26, 27]. Starting from the 1992, the study concerning the presynaptic release-regulating NMDA auto- and heteroreceptors in nerve endings demonstrated that these receptors can be “metamodulated” by colocalized receptors and/or carrier, whose activation and/or blockade enables or disenables the NMDA-mediated functional responses, representing therefore an indirect mechanisms of activation/inhibition of the receptors in physiological condition (i.e., in the presence of millimolar Mg2+ ions) [28]. Also in these cases, our approach was pivotal to describe the mechanisms of the physical/functional crosstalk at the basis of the “conditional” control of the NMDA receptors, paving the road to the proposal of new therapeutic approaches to modulate the NMDA-mediated signaling in central neurological diseases. The opportunities that derive from our studies are impressive when considering that NMDA receptors represent the preferential target of therapies for the cure of most of the central diseases but that this therapeutic approach is unmet because of the lack of drugs acting on the NMDA receptors with a safe profile. In order to describe the complexity and the pleiotropic mechanisms of metamodulation of the presynaptic releaseregulating NMDA receptors, we here briefly resume some findings obtained by using synaptosomes in superfusion that allowed to describe the colocalization and the functional interaction linking the NMDA receptor complex with ionotropic receptors, metabotropic receptors, and carrier in purified synaptosomes. The results from these studies would suggest new classes of Indirect Allosteric Modulator (IAM) of the NMDA receptor for therapeutic purposes.

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3.1 Metamodulation Involving Ionotropic Receptors

It is known that the NMDA receptors are associated to voltageoperated calcium channels, which gating is triggered by the local membrane depolarization, while rating is controlled by glutamate and glycine acting at the GluN subunits participating to the receptor expression. The local depolarization of the plasma membrane bearing the NMDA receptors also can be triggered by the activation of ligand-gated ionotropic receptors localized nearby the NMDA receptors in nerve terminals. It is the case of the presynaptic AMPA and nicotinic receptors. Both receptors associate to ligand-sensitive ionic channels that are permeant to monovalent cations, whose activation elicit the exocytosis of transmitters in physiological condition (i.e., the presence of physiological Mg2+ ions in the external milieu). Results in the literature unveiled that the exposure in superfusion of synaptosomes to non-saturating concentration of the respective agonists (namely, AMPA and nicotine, respectively) causes a transmitter overflow that could be further increased by the concomitant addition of NMDA agonists that on their own failed to cause transmitter overflow. The transmitter release elicited by AMPA or nicotine plus NMDA was partially inhibited by NMDA antagonists and totally prevented by AMPA or nicotine antagonists, compatible with conclusion that the two non-NMDA receptors exerted a permissive role on the colocalized NMDA receptors, unveiling their releasing activity despite the presence of physiological [Mg2+]out [29–31]. Quite interestingly, the receptor-evoked activation of the presynaptic release-regulating NMDA receptors caused functional changes in the NMDA component that emerged as changes in the affinity and the efficacy of agonists/antagonists at this receptor as well as in modification of the subunit composition [32, 33]. Comparable results were obtained when quantifying the release of transmitter evoked by nicotine alone and nicotine in the presence of NMDA/glycine [31, 34]. Notably, by using synaptosomes it was possible to correlate the metamodulation of the NMDA receptors to the synaptosomal exocytosis efficiency, to the influx of calcium ions into the isolated particles and last but not least to the in-out movements of GluN subunit proteins in synaptosomal plasma membranes (as discussed below).

3.2 Metamodulation Involving Metabotropic Receptors

Synaptosomes also possess metabotropic receptors that finely tune the mechanisms of vesicular exocytosis [21, 35]. The data from the studies with superfused synaptosomes indicates that presynaptic metabotropic receptors often fail to elicit on their own a releasing activity but in most cases finely tune the molecular events leading to exocytosis, reinforcing or inhibiting this event [36, 37]. Again, the possibility to discriminate between basal and depolarized conditions represents one of the unique features of the technique of the up-down superfusion of synaptosomes. The approach permits to highlight whether the activation of a

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presynaptic release-regulating receptor (in this case the metabotropic one) (1) can trigger transmitter release, (2) can potentiate transmitter exocytosis or (3) cannot modify transmitter outflow from nerve endings. The possibility is particularly relevant since it allows to define the impact of a selected receptor ligands in resting or depolarized conditions, improving the knowledge on their main role in controlling synaptic plasticity. Starting from the early 2000s, evidence was provided supporting the notion that in isolated nerve endings metabotropic receptors colocalize with NMDA receptors and, depending on the intraterminal pathway they are linked to, they favor, potentiate, or inhibit the NMDA-mediated functions, including the releasing activity. In particular, when focusing on those metabotropic receptors that couple facilitatory G proteins, the data available in the literature clearly demonstrated the existence of G protein coupled receptors (GPCRs) colocalized with the NMDA receptors the activation of which strengthens the NMDA-mediated releasing activity (i.e., the mGlu1/mGlu5 heterodimer that coexists with NMDA heteroreceptors in hippocampal noradrenergic nerve endings), [21, 38] or discloses the NMDA-mediated releasing efficiency despite the presence of physiological concentration of Mg2+ ions in the external milieu (i.e., the somatostatinergic subtype 5 receptors, sst5, colocalized with the NMDA heteroreceptors) [39, 40]. In both cases, the metabotropic receptors control the functions of the colocalized NMDA receptors by affecting the phosphorylation of the GluN subunits, although through differente cascade of events: the PLC-dependent, PKC-mediated src-induced phosphorylation of tyrosine residues in the inner sides of the GluN subunits for the former case [38]; the CaMKIIdependent PKC-mediated phosphorylation of serine and threonine residues in the latter case, [40]. The different phosphorylative pathways were dissected by using selective permeant kinases inhibitors and mimicked by entrapping (see for technical details on this approach the review by Raiteri and Raiteri, [9]) the phosphorylative enzymes in the activated form [40] correlating the induced changes of the receptor-evoked release of transmitter to the functional modulation of the enzymes targeted by the kinase modulators/ substrates. In this case, the use of superfused synaptosomes allowed to decode the intraterminal enzymatic pathways linking the occupancy by agonists of the outer binding sites at plasma membrane receptors with the intraterminal mechanisms that control the vesicles recruitment, docking, and fusion with plasma membranes. It is worth stressing that these events can be monitored independently on the density of the synaptosomal subpopulation involved in the transmitter release, since the use of the radioactive tracer permits to functionally isolate a defined subpopulation of terminals (i.e., those able to actively taken up the radioactive molecule) independently

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on its consistency and to monitor the events occurring in that subpopulation, leaving out the other synaptosomal subpopulations, also if prevalent, in the entire synaptosomal preparations. This is not true for the immunochemical approaches, as in this case the signal supporting the correlation between the structural changes of intraterminal proteins and the activation of membrane receptors is diluted because of the relative low percentage of the responsive synaptosomal subpopulation with respect to the entire one. The use of synaptosomes in superfusion therefore allows a “precise analysis” of the events occurring in nerve terminals. 3.3 Metamodulation Involving Transmitter Transporter

Besides the functional and physical interactions linking two receptors, metamodulation of receptors also could be driven by other membrane structures which might influence the functions of the colocalized receptor protein in different way. As far as the presynaptic release-regulating NMDA receptors are concerned, glycine transporters (GLYTs) are membrane proteins that finely metamodulate the glutamate receptors by exerting a complex mechanism of control of the releasing activity. We are referring particularly to the GLYT type 1 (GLYT1) that, beside its well-known expression in astrocytes, also localized in neurons, on presynaptic terminals, where it closely associates to the NMDA receptors [41]. The GLYT1 takes up the endogenous glycine present in the synaptic cleft, then controlling its concentration nearby the colocalized NMDARs and maintaining it to non-saturating level, indirectly impeding the overactivation of the glutamate receptor [42]. GLYT1, however, is a Na(+)/Cl( )-dependent transporters with electrogenic properties. The uptake of glycine implies the concomitant net influx of positive charges nearby the colocalized NMDA receptors that causes a transient membrane depolarization [42, 43]. We investigated whether this local modification of plasma membrane potential might influence the function on the colocalized NMDA receptors and accordingly to the hypothesis found that the release of transmitter elicited by NMDA in the presence of saturating glycine and physiological Mg2+ ions was dramatically reduced by the concomitant presence of GLYT1 blockers or by reducing the [Na+]out [41]. Taking into account that the synaptosomal subpopulation under study was glutamatergic, which represents the prevailing synaptosomal subfamily in the entire hippocampal synaptosomal preparation, it was possible to confirm the functional observations with an immunochemical data that definitively proved the colocalization of the GLYT1 and the NMDA receptor subunits. We confirmed the colocalization of the GLYT1 and the GluN2B and GluN1 subunits in the synaptosomal presynaptic plasma membranes fraction with Western blotting analysis. Furthermore, we demonstrated that the GluN2B immunoprecipitated component

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of the synaptosomal presynaptic plasma membranes subfraction is immuno-positive for the glycinergic carrier, confirming the physical association between the receptor complex and the glycine transporter [41]. 3.4 Metamodulation and Presynaptic Receptors Trafficking

It is known that ionotropic glutamatergic receptors (namely, AMPA and NMDA receptors) traffic in-out the plasma membranes and that their movements occur constitutively or can be speeded up by concomitant events [44–47]. Ionotropic glutamate receptors undergo a constitutive trafficking that assure the continuous renewing of the receptor proteins in plasma membranes [44]. By using synaptosomes, we demonstrated that presynaptic release-regulating AMPA receptors controlling noradrenaline, dopamine, and glutamate traffic constitutively [46]. Furthermore, we demonstrated that forcing the insertion/ deletion of receptor subunits in synaptosomal plasma membranes was paralleled by quantitative changes in the efficiency of transmitter release, allowing a direct correlation between the insertion of receptor subunits in synaptosomal plasma membranes and the amount of transmitter released. The constitutive trafficking was highlighted by a functional point of view by using peptides that compete for the binding sites of receptor subunits to cytosolic proteins that drive the endocytosis of receptors as well as their insertion in membranes. Altogether these results confirmed the superfused synaptosomes as a useful approach to study the mechanisms of active insertion or removal of receptors from presynaptic neuronal membranes. An interesting finding of our research was that the constitutive in-out movement that typify the trafficking of receptors in presynaptic plasma membranes is also dynamically regulated by the concomitant activation of colocalized receptors (i.e., the nicotinic receptors and the NMDA receptors as well) [31, 33, 34]. The trafficking of the NMDA receptor elicited by metamodulators, however, emerged when the activation of the colocalized receptors was prolonged during time, consistent with the view that the sustained activation of one of the two counterparts of the receptor-receptor complex causes adaptative changes that drastically affect the receptor crosstalk, changing their impact at the presynaptic level. It is the case of the nicotine receptors, whose acute activation discloses the signaling of colocalized NMDA receptors in conditions that usually impedes their releasing activity (i.e., the presence of physiological [Mg2+]), [47] without affecting the GluN expression in synaptosomal membranes, but drastically reduces the insertion of GluN2B subunits in synaptosomal plasma membranes as well as the releasing activity of the ionotropic glutamate receptor when the exposure of synaptosomes to nicotine is prolonged during time [31, 34]. The shift from facilitation to inhibition was also observed turning from the acute to the

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prolonged activation of the chemokine receptors CXCR4 colocalized with the NMDA receptors in glutamatergic and noradrenergic nerve endings. In this case, the prolonged activation of the CXCR4 did not modify the insertion of GluN subunits in plasma membranes but permitted the phosphorylation of residues in the GluN1 subunits that were not targeted during the acute activation of the colocalized receptors [35, 48]. These observations seem best interpreted by assuming that the metamodulation of the receptor trafficking is an event that determines long-lasting adaptative changes in nerve terminals that are preferentially induced by conditioning stimuli and might account for the mechanisms of synaptic plasticity.

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Concluding Remark Starting from the proposal of the “up-down superfusion” of a synaptosomal monolayer, the knowledge and the use of this technique has largely increased, allowing the answer to emerging questions and unexpected observations pivotal to comprehend synaptic transmission. The results reviewed in this article suggest the superfusion of synaptosomes as an approach of choice to investigate the mechanisms of metamodulation of receptors presynaptically located, to clarify how do they interact, the intraterminal pathways accounting for the receptor coupling, and which are the consequences of their concomitant activation, permitting the precise decoding of the receptor–receptor coupling in selected subpopulations of nerve terminals. By a functional point of view, the technique permits to isolate the events that occur presynaptically, avoiding interferences due to postsynaptic signaling, which represents the main limit of other experimental techniques including the electrophysiological approaches. Of course, the availability of selective tools for the receptor subtypes is fundamental to dissect functionally the receptor-receptor crosstalk and the intimate correlation linking the different receptors but, for some receptors, this condition is unsatisfied limiting the investigation of the receptor–receptor interaction. Conversely, the functional decoding of the relationship linking the receptors involved in the metamodulation of neuromodulation indirectly provides an experimental model for the screening of molecules whose target is unknown and that could represent new therapeutics for central diseases. Last but not least, a so far poorly explored possibility is that aberrant metamodulation could account for the development of neurological disorders. In this case, the synaptosomes isolated from the CNS of animals suffering from that disease would represent a suitable model to analyze the altered metamodulation of presynaptic receptors and the corrective actions to restore the functional crosstalk. All these considerations suggest that the superfusion of synaptosomes is a “no-ending” story that will assure in the future new advance in the study of the CNS functions.

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Acknowledgments This article is dedicated to all our colleagues and friends that starting from the 1980s (40 years ago, when the superfusion technique was first introduced at the Institute of Pharmacology, Faculty of Pharmacy, University of Genova) have carried out their studies using this methodological approach contributing to improve the knowledge of the presynaptic events of the chemical transmission in the central nervous system. References 1. Langer SZ (1974) Presynaptic regulation of catecholamine release. Biochem Pharmacol 23 (13):1793–1800 2. Langer SZ (1997) 25 years since the discovery of presynaptic receptors: present knowledge and future perspectives. Trends Pharmacol Sci 18(3):95–99 3. Raiteri M, Marchi M, Maura G et al (1989) Presynaptic regulation of acetylcholine release in the CNS. Cell Biol Int Rep 13 (12):1109–1118 4. Wonnacott S, Drasdo A, Sanderson E et al (1990) Presynaptic nicotinic receptors and the modulation of transmitter release. Ciba Found Symp 152:87–101; discussion 102-105 5. von Ku¨gelgen I, Kurz K, Bu¨ltmann R et al (1994) Presynaptic modulation of the release of the co-transmitters noradrenaline and ATP. Fundam Clin Pharmacol 8(3):207–213 6. Raiteri M (1994) Functional studies of neurotransmitter receptors in human brain. Life Sci 54(22):1635–1647 7. Raiteri M (1987) Release in vitro as a model to study neurotransmitter receptors. Pharmacol Res Commun 19(12):927–941 8. Dunant Y, Israe¨l M (1998) In vitro reconstitution of neurotransmitter release. Neurochem Res 23(5):709–718 9. Raiteri L, Raiteri M (2000) Synaptosomes still viable after 25 years of superfusion. Neurochem Res 25:1265–1274 10. Bennett MR, Kearns JL (2000) Statistics of transmitter release at nerve terminals. Prog Neurobiol 60(6):545–606 11. Pinheiro PS, Mulle C (2008) Presynaptic glutamate receptors: physiological functions and mechanisms of action. Nat Rev Neurosci 9 (6):423–436 12. Gray BE, Wittaker VP (1962) The isolation of nerve endings from brain: an electronmicroscopic study of cell fragments derived by

homogenization and centrifugation. J Anat 96:79–91 13. Jones DG, Bradford HF (1971) Observations on the morphology of mammalian synaptosomes following their incubation and electrical stimulation. Brain Res 28(3):491–499 14. Harrison SM, Jarvie PE, Dunkley PR (1988) A rapid Percoll gradient procedure for isolation of synaptosomes directly from an S1 fraction: viability of subcellular fractions. Brain Res 441 (1–2):72–80 15. Dunkley PR, Jarvie PE, Robinson PJ (2008) A rapid Percoll gradient procedure for preparation of synaptosomes. Nat Protoc 3 (11):1718–1728 16. de Belleroche JS, Bradford HF (1972) Synaptosome beds: a method for the study in vitro of the metabolism and function of nerve endings. Biochem J 127(2):21P 17. Raiteri M, Angelini F, Levi G (1974) A simple apparatus for studying the release of neurotransmitters from synaptosomes. Eur J Pharmacol 25(3):411–414 18. Langer SZ (2008) Presynaptic autoreceptors regulating transmitter release. Neurochem Int 52(1–2):26–30 19. Raiteri M (2008) Presynaptic metabotropic glutamate and GABAB receptors. Handb Exp Pharmacol 184:373–407 20. Raiteri L, Raiteri M (2015) Multiple functions of neuronal plasma membrane neurotransmitter transporters. Prog Neurobiol 134:1–16 21. Pittaluga A (2016) Presynaptic releaseregulating mGlu1 receptors in central nervous system. Front Pharmacol 7:295 22. Pittaluga A (2019) Acute functional adaptations in isolated presynaptic terminals unveil Synaptosomal learning and memory. Int J Mol Sci 20(15):3641 23. Fuxe K, Agnati LF (1985) Receptor-receptor interactions in the central nervous system. A

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new integrative mechanism in synapses. Med Res Rev 5(4):441–482 24. Katz PS, Edwards DH (1999) Metamodulation: the control and modulation of neuromodulation. In: Kats PS (ed) Beyond neurotransmission. Neuromodulation and its importance for information processing. Oxford University Press, New York, pp 349–382 25. Sebastia˜o AM, Ribeiro JA (2015) Neuromodulation and metamodulation by adenosine: Impact and subtleties upon synaptic plasticity regulation. Brain Res 1621:102–113 26. Paoletti P, Neyton J (2007) NMDA receptor subunits: function and pharmacology. Curr Opin Pharmacol 7:39–47 27. Stroebel D, Casado M, Paoletti P (2017) Triheteromeric NMDA receptors: from structure to synaptic physiology. Curr Opin Physiol 2:1–12 28. Pittaluga A, Pattarini R, Feligioni M et al (2001) N-methyl-D-aspartate receptors mediating hippocampal noradrenaline and striatal dopamine release display differential sensitivity to quinolinic acid, the HIV-1 envelope protein gp120, external pH and protein kinase C inhibition. J Neurochem 76:139–148 29. Pittaluga A, Raiteri M (1992a) N-methyl-Daspartic acid (NMDA) and non-NMDA receptors regulating hippocampal norepinephrine release. I. Location on axon terminals and pharmacological characterization. J Pharmacol Exp Ther 260:232–237 30. Marchi M, Grilli M (2010) Presynaptic nicotinic receptors modulating neurotransmitter release in the central nervous system: functional interactions with other coexisting receptors. Prog Neurobiol 92:105–111 31. Zappettini S, Grilli M, Olivero G et al (2014) Nicotinic α7 receptor activation selectively potentiates the function of NMDA receptors in glutamatergic terminals of the nucleus accumbens. Front Cell Neurosci 8:332 32. Pittaluga A, Raiteri M (1992b) N-methyl-Daspartic acid (NMDA) and non-NMDA receptors regulating hippocampal norepinephrine release. III. Changes in the NMDA receptor complex induced by their functional cooperation. J Pharmacol Exp Ther 263:327–333 33. Summa M, Di Prisco S, Grilli M et al (2011) Hippocampal AMPA autoreceptors positively coupled to NMDA autoreceptors traffic in a constitutive manner and undergo adaptative changes following enriched environment training. Neuropharmacology 61(8):1282–1290 34. Salamone A, Zappettini S, Grilli M et al (2014) Prolonged nicotine exposure down-regulates presynaptic NMDA receptors in dopaminergic

terminals of the rat nucleus accumbens. Neuropharmacology 79:488–497 35. Olivero G, Cisani F, Vergassola M et al (2019) Prolonged activation of CXCR4 hampers the release-regulating activity of presynaptic NMDA receptors in rat hippocampal synaptosomes. Neurochem Int 126:59–63 36. Parodi M, Patti L, Grilli M et al (2006) Nicotine has a permissive role on the activation of metabotropic glutamate 5 receptors coexisting with nicotinic receptors on rat hippocampal noradrenergic nerve terminals. Neurochem Int 48(2):138–143 37. Luccini E, Musante V, Neri E et al (2007) Functional interactions between presynaptic NMDA receptors and metabotropic glutamate receptors co-expressed on rat and human noradrenergic terminals. Br J Pharmacol 151:1087–1094 38. Longordo F, Feligioni M, Chiaramonte G et al (2006) The human immunodeficiency virus-1 protein transactivator of transcription up-regulates N-methyl-D-aspartate receptor function by acting at metabotropic glutamate receptor 1 receptors coexisting on human and rat brain noradrenergic neurones. J Pharmacol Exp Ther 317:1097–1105 39. Pittaluga A, Bonfanti A, Raiteri M (2000) Somatostatin potentiates NMDA receptor function via activation of InsP(3) receptors and PKC leading to removal of the mg(2+) block without depolarization. Br J Pharmacol 130:557–566 40. Pittaluga A, Feligioni M, Longordo F et al (2005) Somatostatin-induced activation and up-regulation of N-methyl-D-aspartate receptor function: mediation through calmodulindependent protein kinase II, phospholipase C, protein kinase C, and tyrosine kinase in hippocampal noradrenergic nerve endings. J Pharmacol Exp Ther 313(1):242–249 41. Musante V, Summa M, Cunha RA et al (2011) Pre-synaptic glycine GlyT1 transporter-NMDA receptor interaction: relevance to NMDA autoreceptor activation in the presence of Mg2+ ions. J Neurochem 117:516–527 42. Arago´n C, Lo´pez-Corcuera B (2005) Glycine transporters: crucial roles of pharmacological interest revealed by gene deletion. Trends Pharmacol Sci 26:283–286 43. Roux MJ, Supplisson S (2000) Neuronal and glial glycine transporters have different stoichiometries. Neuron 25(2):373–383 44. Collingridge GL, Isaac JT (2003) Functional roles of protein interactions with AMPA and kainate receptors. Neurosci Res 47(1):3–15

Synaptosomes and Metamodulation 45. Marchi M, Grilli M, Pittaluga AM (2015) Nicotinic modulation of glutamate receptor function at nerve terminal level: a fine-tuning of synaptic signals. Front Pharmacol 6:89 46. Pittaluga A, Feligioni M, Longordo F et al (2006) Trafficking of presynaptic AMPA receptors mediating neurotransmitter release: neuronal selectivity and relationships with sensitivity to cyclothiazide. Neuropharmacology 50(3):286–296

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Chapter 9 A Novel Method to Monitor GABA Loading into Synaptic Vesicles by Combining Patch Pipette Perfusion and Intracellular, Caged-GABA Photolysis in Brain Slice Preparations Manami Yamashita and Tetsuya Hori Abstract A given concentration of GABA can be introduced into a presynaptic terminal by patch clamping the soma of a presynaptic neuron, if the neuron has a relatively short axon. By combining patch pipette perfusion or intracellular, caged-GABA photolysis, it is possible to measure various parameters related to synaptic vesicle filling with GABA. Key words Synaptic vesicles, GABA, Slice preparation, Pipette perfusion, Intracellular photolysis

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Introduction One of the features that makes whole-cell patch clamping useful is that molecules in a patch pipette diffuse rapidly into the cell. While whole-cell patch clamping makes it possible to vary the intracellular environment, artificial replacement of cytoplasmic composition using whole-cell patch clamping has the disadvantage of creating non-physiological conditions within the cell. Nonetheless, manipulation of intracellular composition is a powerful experimental method that has enabled historically important discoveries. In one famous experiment, the cytoplasm of a squid giant axon was replaced with an artificial intracellular perfusate [1]. In another, cytoplasmic Ca2+ concentration was elevated at the presynaptic terminal of a squid giant synapse using intracellular photolysis of DM-nitrophen [2]. Whole-cell patch clamping while introducing new molecules via patch pipette perfusion or by intracellular photolysis allows us to perform “single cell intracellular pharmacology” by changing intracellular composition during an experiment.

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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At chemical synapses, neurotransmitters such as glutamate, GABA, acetylcholine, and monoamines are essential for neurotransmission. These neurotransmitters are stored in synaptic vesicles (SVs) and are released by exocytosis. After exocytosis, SVs are retrieved by endocytosis, refilled with neurotransmitters, and recycled to be reused in a subsequent round of transmission. For recycled vesicles to contribute to synaptic transmission, they must be refilled with sufficient quantities of neurotransmitter before being reused; therefore, efficient filling of synaptic vesicles with neurotransmitter is essential. Each neurotransmitter has specific transporters that pump a given neurotransmitter into synaptic vesicle lumens [3]. Numerous biochemical studies have investigated properties of neurotransmitter transporters; however, until the present decade, transport speed measurements had been limited to studies using isolated or reconstituted synaptic vesicles (glutamate: [4, 5]; GABA: [6, 7]). In 2012, Hori and Takahashi [8] directly determined the vesicle filling rate at central glutamatergic synapses. Six years later, such measurements were accomplished at GABAergic synapses in rat brain slices, using simultaneous presynaptic and postsynaptic whole-cell recording, combined with patch pipette perfusion and caged glutamate/GABA photolysis in presynaptic neurons [9]. In the present paper, we explain methods for both GABA perfusion and GABA photolysis experiments at inhibitory synapses in the molecular layer of the cerebellum.

2

Materials Prepare all solutions using ultrapure water.

2.1

Slice Preparation

1. Cutting solution: 250 mM sucrose, 2.5 mM KCl, 26 mM NaH2PO4, 0 mM CaCl2, 6 mM MgCl2, 10 mM glucose, 3 mM myo-inositol, 2 mM sodium pyruvate, and 5 mM ascorbic acid. 2. Artificial cerebrospinal fluid (aCSF): 125 mM NaCl, 2.5 mM KCl, 26 mM NaH2PO4, 2 mM CaCl2, 6 mM MgCl2, 10 mM glucose, 3 mM myo-inositol, 2 mM sodium pyruvate, and 5 mM ascorbic acid (pH 7.3 when bubbled with 95% O2/5% CO2). 3. Wistar/ST rats (postnatal day 12–16) of either sex. 4. Microtome.

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2.2 Electrophysiological Recording (Simultaneous Presynaptic and Postsynaptic Recordings)

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1. EPC-10/2 amplifier controlled by the Pulse Program. 2. Postsynaptic pipette solution: 170 mM CsCl, 10 mM HEPES, 5 mM EGTA, 1 mM MgCl2, 5 mM QX314 (adjusted to pH 7.3–7.4 with CsOH). 3. Presynaptic pipette solution: 97.5 mM potassium methanesulfonate, 32.5 mM KCl, 12 mM sodium phosphocreatine, 3 mM Mg-ATP, 0.5 mM Na-GTP, 0.5 mM EGTA, 40 mM HEPES (adjusted to pH 7.3–7.4 with KOH) (see Note 1). 4. Patch Pipette: Glass capillaries, normal wall pipette with filament.

2.3

Pipette Perfusion

1. Thin glass capillary. 2. Picopump. 3. Presynaptic solution containing defined concentration of GABA.

2.4 DPNI-GABA Uncaging

1. 1-(4-Aminobutanoyl)-4-[1,3-bis(dihydroxyphosphoryloxy)propan-2-yloxy]-7-nitro-indoline, (DPNI-GABA, Tocris Cookson). 2. GSH stock solution: 100 mM Glutathione, reduced form and 10 mM HEPES, adjusted to pH 7.3–7.4 with KOH. 3. Mercury lamp light source. 4. Microscope shutter. 5. Presynaptic solution 2: 130 mM potassium methanesulfonate, 0 mM KCl, 12 mM sodium phosphocreatine, 3 mM Mg-ATP, 0.5 mM Na-GTP, 0.5 mM EGTA, 40 mM HEPES (adjusted to pH 7.3–7.4 with KOH). 6. Presynaptic solution 3: 0 mM potassium methanesulfonate, 130 mM KCl, 12 mM sodium phosphocreatine, 3 mM MgATP, 0.5 mM Na-GTP, 0.5 mM EGTA, 40 mM HEPES (adjusted to pH 7.3–7.4 with KOH).

3 3.1

Methods Slice Preparation

3.2 Electrophysiological Recording (Pair Recording)

After decapitation, sagittal slices (200 μm) are cut in cutting solution from cerebellar cortices of Wistar/ST rats (postnatal day 12–16, either sex) using a microtome. After cutting, slices are superfused with aCSF. 1. Whole-cell patch-clamp recordings are made simultaneously from a presynaptic BC and a postsynaptic PC (Fig. 1b) under voltage clamp at holding potentials of 60 mV (BC) and 80 mV (PC), using an EPC-10/2 amplifier controlled by the Pulse Program (see Notes 2 and 3). For postsynaptic

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Fig. 1 Paired recordings in cerebellar cortex. (a) Schematic illustration of a paired recording from a presynaptic BC (b) and a postsynaptic PC (P). S denotes a stellate cell. (Adopted with permission from Yamashita, Kawaguchi, Hori et al. [9]). (b) Simultaneous patch-clamp recording from a Purkinje cell and an inhibitory interneuron in the molecular layer of a cerebellar slice. (c) Visualization of an interneuronal axon with Alexa488 loading. Scale bar, 10 μm. The oval indicates the synaptic site

IPSC recordings, patch pipettes (resistance, 2.0–3.5 MΩ) contain CsCl solution. For presynaptic recordings, patch pipettes (3–4 MΩ) contain potassium methanesulfonate solution. 2. To evoke IPSCs in PCs, APs are elicited in presynaptic BCs with a 2-ms depolarizing command pulse to 0 mV. 3.3

Pipette Perfusion

1. Heat-pull an Eppendorf “yellow tip” to make the plastic tube. 2. Cut the plastic tube made in previous step 1 at a diameter of 350–400 μm, allowing the glass tube (φ: 300 μm) to fit inside it. 3. Insert the glass tube into the plastic tube and seal the junction with gentle heating (Fig. 2a). 4. Mount the tube assembly onto a three-port (Y-shape) pipette holder. 5. Heat-pull the glass pipette using a horizontal pipette puller to achieve a tip diameter of 10–20 μm (Fig. 2b) (see Note 4).

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Fig. 2 Patch pipette perfusion system. (a) Thin glass tube inserted into a fine plastic tube heat-pulled from a yellow tip. (b) An enlarged view of the tip of the infusion glass tube (a), processed with a pipette puller. (c) The infusion glass tube inserted into a presynaptic patch pipette

6. Backfill the presynaptic solution containing GABA into the infusion glass tube. 7. Backfill the presynaptic solution without GABA into the same infusion glass tube (Fig. 2c) to minimize dilution by the control solution. This leaves the presynaptic solution without GABA at the tip of the pipette and the GABA-containing solution behind that. 8. Fill the presynaptic solution without GABA into patch pipette.

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Fig. 3 Data collected during pipette perfusion, showing the IPSC rundown after GABA washout and recovery following GABA reloading. After IPSCs ran down to a low steady level, GABA was infused into a presynaptic BC via pipette perfusion: at 2 mM GABA (a) and 5 mM GABA (b). (Adopted with permission from Yamashita, Kawaguchi, Hori et al. [9])

9. Insert the infusion glass tube in the patch pipette with its tip recessed 150–200 μm inside the tip of the patch pipette (Fig. 2c), which enables quick infusion of GABA into a presynaptic soma. 10. Collect control data (Fig. 3) (see Note 5). 11. GABA-containing solution is infused from the capillary into a patch pipette under a positive pressure (8–12 psi) using a Picopump PV820. 3.4 DPNI-GABA Uncaging

1. Mix 740 μL presynaptic solution 2, 300 μL presynaptic solution 3, and 260 μL GSH stock solution, to make a presynaptic pipette solution cocktail for GABA uncaging. The solution cocktail contains 20 mM reduced glutathione to minimize cell toxicity associated with UV uncaging. 2. Dissolve DPNI-GABA (10 mM) in the presynaptic pipette solution cocktail made in the previous step 1 on the day of experiments, and load into a BC soma through whole-cell pipettes (see Note 6). 3. Deliver a 1-s UV light pulse from a mercury lamp light source through an objective lens by opening a shutter controlled by a shutter driver (Fig. 4) (see Note 7).

4

Notes 1. Store the solution containing ATP and/or GTP at

80  C.

2. Basket cells (BCs) and stellate cells are distinguished based on their positions within the molecular layer. Interneurons in the lower third are considered BCs, and those in the upper two-thirds are considered stellate cells (Fig. 1a) [10].

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Fig. 4 Data illustrating DPNI-GABA uncaging. Recovery of IPSCs from rundown with a UV pulse (1 s) that uncaged DPNI-GABA. The UV pulse was applied at time 0. (Adopted with permission from Yamashita, Kawaguchi, Hori et al. [9])

3. A basket cell sends axons that form collaterals to the somas of Purkinje cells. Basket cells are filled with Alexa488 which is directly infused through the patch pipette. It helps to identify the synaptic pair (Fig. 1c). 4. If the tip diameter is too small for smooth injection, break the tip with tweezers. 5. Use low-resistance pipettes to replace cytoplasmic GABA to the pipette concentration as quickly as possible. 6. Experiments should be performed in the dark and transillumination from the microscope is turned off after establishing a paired recording for GABA uncaging. 7. Control UV light intensity (weak and long) for DPNI-GABA uncaging. References 1. Hodgkin AL, Keynes RD (1956) Experiments on the injection of substances into squid giant axons by mean of a microsyringe. J Physiol 131(3):592–616 2. Delaney KR, Zucker RS (1990) Calcium released by photolysis of DM-nitrophen stimulates transmitter release at squid giant synapse. J Physiol 426:473–498 3. Takamori S (2016) Presynaptic molecular determinants of quantal size. Front Synaptic Neurosci 8:2 4. Maycox PR, Deckwerth T, Hell JW et al (1988) Glutamate uptake by brain synaptic vesicles. Energy dependence of transport and functional

reconstitution in proteoliposomes. J Biol Chem 263(30):15423–15428 5. Carlson MD, Kish PE, Ueda T (1989) Characterization of the solubilized and reconstituted ATP-dependent vesicular glutamate uptake system. J Biol Chem 264(13):7369–7376 6. Kish PE, Fischer-Bovenkerk C, Ueda T (1989) Active transport of γ-aminobutyric acid and glycine into synaptic vesicles. Proc Natl Acad Sci U S A 86(10):3877–3881 7. Hell JW, Maycox PR, Jahn R (1990) Energy dependence and functional reconstitution of the γ-aminobutyric acid carrier from synaptic vesicles. J Biol Chem 265(4):2111–2117

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8. Hori T, Takahashi T (2012) Kinetics of synaptic vesicle refilling with neurotransmitter glutamate. Neuron 76(3):511–517 9. Yamashita M, Kawaguchi SY, Hori T et al (2018) Vesicular GABA uptake can be rate

limiting for recovery of IPSCs from synaptic depression. Cell Rep 22(12):3134–3141 10. Palay SL, Chan-Palay V (1974) Cerebellar cortex. Springer-Verlag, Berlin, Heidelberg, New York

Chapter 10 Rapid Isolation of Functional Synaptic Vesicles from Tissues Through Cryogrinding, Ultracentrifugation, and Size Exclusion Chromatography Huinan Li Abstract Many biochemical and biophysical related questions require the isolation of functional synaptic vesicles. Isolated synaptic vesicles can be used for transporter kinetics studies, synaptic vesicle content analysis and immuno-labeling of specific synaptic vesicle proteins, etc. Here I describe a fast and reliable isolation procedure to allow researchers to isolate a large amount, as well as physiologically functional synaptic vesicles, by following the subsequent order of cryogrinding, gradient ultracentrifugation, and size exclusion liquid chromatography. This process enriches over 90% of the synaptic vesicle population, with low contamination of Golgi or endoplasmic reticulum vesicles. Key words Synapses, Presynaptic terminal, Synaptic vesicles, Neurotransmitter, Synaptic vesicular protein, Neurotransmitter transporter, Vesicular content, Organelle isolation, Gradient centrifugation, Size exclusion chromatography

1

Introduction Synaptic transmission enables neuronal communication in the central nervous system, neurons communicating with muscle cells or secretory glands in the peripheral nervous system. Structurally, a cell-to-cell connection named synapse is responsible for this process [1]. There are two major types of synapses: electrical synapses and chemical synapses. Chemical synapses function when neurotransmitters are released from the presynaptic side to the postsynaptic side. Neurotransmitters can bind their receptors on the postsynaptic membrane to trigger downstream effects [2, 3]. With the advancement of transmission electron microscopy (TEM), Heuser and colleagues [4, 5] were able to demonstrate that neurotransmitters are released in a quantal fashion. The individual packet containing neurotransmitters is termed synaptic vesicle. Under transmission electron microscopy, one can observe that there

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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are abundant number of synaptic vesicles at each presynaptic terminal [6]. Some of these synaptic vesicles are docked at the electrondense area named active zone material [6]. At the arrival of an action potential, exogenous calcium enters the presynaptic terminal through electrically gated calcium channels, triggering some of the docked synaptic vesicles to fuse with the presynaptic membrane, and the subsequent release of the neurotransmitter into the synaptic cleft for binding postsynaptic receptors. It has been relatively well understood now that it is through a series of synaptic vesicle proteins interacting with presynaptic membrane proteins that makes synaptic vesicles accessible for release [7]. Many aspects of the studies to address synaptic transmission-related questions have been done in tissue preparation or cell culture. However, to understand how a synaptic vesicle is built requires the isolation of synaptic vesicles from the tissue. Ohsawa and colleague were among the first group of people that isolated relatively pure synaptic vesicles from organism directly and studied the lipid and protein content of synaptic vesicles [8]. Takamori and colleague isolated synaptic vesicles from mouse brains and identified the species of many proteins residing on the synaptic vesicles, as well as the ones inside and associated with them [9]. Mutch and colleague used isolated synaptic vesicles to quantify each synaptic vesicle protein’s copy numbers on an individual synaptic vesicle level. Many of the isolation protocols take a long time to isolate the vesicles and result in low yield. Here, we describe a detailed optimized protocol of isolating intact functional synaptic vesicles from frozen tissue. We have used this protocol and demonstrated that this method preserves epitopes of the synaptic vesicle protein for immunolabeling [10]. In addition, these synaptic vesicles can be used for TEM as well as TEM tomography at both room temperature and cryo-TEM preparation, and even the content inside can be analyzed with further mass spectrometry or RNA content sequencing [11]. They can also be used for vesicle loading to understand neurotransmitter transporters residing on individual synaptic vesicles. This protocol addresses many biochemical and biophysical properties of this fascinating subcellular organelle.

2 2.1

Materials Sucrose Beads

1. Sucrose Beads Buffer (at least 125 mL for making sucrose beads and warming up cryogrinding powder to 4
C): 320 mM sucrose, 10 mM Tris–HCl, pH 7.4 (see Notes 1 and 2). 2. Liquid nitrogen. 3. Colander. 4. Ice bucket.

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5. Glass pipettes. 6. 50 mL canonical tubes. 7. Sucrose gradient buffer: 0.6 M and 1.2 M sucrose buffer in 10 mM Tris–HCl, pH 7.4 (can be stored at 4
C for future use for up to 3 months). 2.2

Cryogrinding

For centrifugation, thaw materials depicted in steps 2–9 at 4
C. 1. Spex Freezer Mill 6800 (Spex Sample Prep, Metuchen, NJ) or similar model, ceramic pestle and mortar. 2. Chemical fume hood. 3. Stainless steel chemical spoon. 4. Magnetic stir bar. 5. Magnetic stir plate. 6. 500 mL glass beaker. 7. Cryoprotection gloves. 8. Goggles. 9. Lab coat.

2.3

Centrifugation

1. Beckman Coulter JA-20 rotor. 2. Avanti J25 centrifuge (4
C storage). 3. 2 JA-20 rotor compatible Beckman Tubes. 4. Gram grade scale. 5. Micropipette and tips. 6. Ice bucket with ice. 7. Beckman Coulter 70ti rotor (4
C storage). 8. Optima 980 centrifuge. 9. 2 70ti compatible Beckman Tubes. 10. Gram scale. 11. Micropipette and tips. 12. 10 mL and 5 mL serological pipettes. 13. Beckman silicon grease. 14. Ice bucket with ice. 15. Eppendorf tubes (see Note 3).

2.4 Fast Protein Liquid Chromatography (FPLC)

1. 0.22 μm spin-X tube filters. 2. Pharmacia LC500 plus Fast Protein Liquid Chromatography (FPLC). 3. 25 cm 4% agarose bead column (Bioscience Beads, West Warwick, RI).

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4. Elute buffer: 0.2 M NaCl, 10 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), pH 7.4. 5. Bradford Assay. 6. Stirred Cell apparatus with a 100 kDa filter (PLHK02510; Millipore, Billerica, MS).

3

Methods

3.1 Preparation of Sucrose Beads

1. Measure 25 mL of sucrose beads buffer, then use glass pipette to make small drops of sucrose beads buffer into liquid nitrogen container (see Note 3), then collect with a colander (sucrose beads can be stored in 50 mL canonical tubes, at 80
C).

3.2

1. Pre-cool Spex Freezer Mill by filling in liquid nitrogen in its full capacity for at least 30 min prior to cryogrinding.

Cryogrinding

2. Pre-cool the cryogrinding apparatus (two medal lids and the magnetic bar) by pouring in liquid nitrogen for 15 min (see Note 4). 3. Pre-cool the pestle and mortar by pouring liquid nitrogen inside for 5 min. 4. Load 25 g tissue samples into the mortar (see Note 5) and break down into 1–3 cm3 sized chunks. 5. Assemble the cryogrinding apparatus with 25 g sample chunks and 25 g pre-made sucrose beads with the magnetic stir bar, as well as the lids at two sides. Then load the apparatus into the freezer mill (see Note 6). 6. Fill liquid nitrogen until the Spex Freezer Mill is full again, before closing the lid. Be careful when closing the lid. 7. Set 30 s grinding and 1 min rest for 3 cycles (see Note 7). 8. During the last round of grinding, set up a stir plate in the chemical hood, with 50 mL 4
C sucrose beads buffer in a 500 mL glass beaker. Add a stir bar inside with mid-speed. 3.3 Warming of the Solution

1. After the grinding, retrieve the apparatus out from the freezer mill with cryoprotection (see Note 8), use the stainless steel chemical spoon to take out the cryogrinding powder in the chemical hood (see Note 9), and put it in the sucrose solution beaker. 2. The magnetic stir bar may not stir immediately after the powder is added in. Keep the speed and stir with the chemical spoon every 5 min, for around 30 min. The sucrose solution turns slurry with a little bit of ice and is then named slurry (Sl). Then, they are ready to be loaded into the centrifuge tubes.

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1. Pre-cool Beckman Avanti centrifuge to 4
C at least 15 min prior to centrifugation. 2. Pre-cool the high-speed Beckman Optima 980 centrifuge to 4
C. 3. Load 50 mL slurry solution into Beckman tubes, balance two tubes with a gram scale, and use a micropipette to adjust the volume. 4. Tubes are centrifuged at 48,297  g for 15 min to pellet tissue debris. Collect the tubes on ice and keep the tubes on ice at all time. The resulting supernatant is named S1, the pellet is named P1.

3.5 High-Speed Centrifugation

1. Slowly take out the supernatant with a 5 mL serological pipette, avoiding the big pellet, by leaving some supernatant in the tubes. The pellets from the low-speed centrifuge mostly consist of myelin and other lipids. 2. Lay the supernatant into the 70ti rotor tube and place the centrifuge tubes on ice at all time. 3. Balance the two tubes on the scale, adjust with micropipette. 4. Load the tubes into the 70ti rotor centrifuge for 40 min at 118,666  g at 4
C. With 20 min left, prepare the sucrose gradient. Place 70ti rotor compatible tubes in the ice bucket when sucrose buffers are taken out from the fridge. 5. Lay 8 mL of 1.2 M sucrose buffer at the bottom of the tubes, followed by 8 mL 0.6 M sucrose buffer (see Note 10). 6. Take out the tubes from the rotor to the ice bucket. Tilt the tube to get the supernatant out and avoid the bottom pellet area, there should be enough to lay on the sucrose gradient tube. The resulting supernatant is named S2, the pellet is named P2. 7. Balance the tubes with micropipette on the scale to make sure they are balanced (see Note 11). 8. Put a small amount of Beckman silicon grease on the rubber ring of the rotor lid to create a vacuum seal. 9. Load the rotor into the ultracentrifugation at 236,510  g for 2 h at 4
C. 10. After centrifugation, unload the tubes to the ice bucket. Use a micropipette to take out 1 mL by 1 mL into Eppendorf tubes, then the most enriched area is the fluffy layer (FL, with pink to red color right above 1.2 M sucrose). 11. Label the numbers, as well as other information like sample name and date. Store the 1 mL fraction into the 80
C freezer for future use.

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3.6 Fast Protein Liquid Chromatography (FPLC) (See Note 12)

1. Prior to running samples, run the sucrose column at low speed (1 mm light path, you may dilute this stock further to a low concentration stock and determine its concentration instead (see Table 1). 15. 22 mW laser intensity at 800 nm are the values used in our laboratory. The given laser intensity might not be optimal for your new setup. To find the optimal excitation intensity, the fluorescence intensity should be plotted against a series of laser powers ranging from, for example, 0.1 to 30 mW. The fluorescence intensity should increase with a quadratic dependency of the laser power to confirm TPE. Too high laser powers will lead to saturation and the fluorescence increase will be reduced. Any laser power that lies for certain in the range of quadratic dependency and results in a good signal-to-noise ratio can be used for the measurements. The wavelength was chosen to give similar signal intensites for liposomes prepared as described in Chap. 13, using the fluorophores Oregon Green and Texas Red. 16. Prepare the inverted microscope by carefull putting a droplet of ultrapure water on top of the objective lens without touching it. Then, put a single coverslip directly onto the objectie. Place the sample over the lens onto the coverslip. Putting the coverslip directly on top of the objective is an easy way as it avoids any stages or holders for the coverslip. Therefore, the coverslip needs to sit directly on top of the lens with just a thin water layer in-between. 17. The maximum signal intensity is mainly dependent on the used PCM and detectors. The maximum count-rates can be found in their specifications. In most cases, the detectors will be the bottleneck. To give an example: For the APDs in our setup, the maximum count rate is given with 5 Mc/s and a peak light intensity of 104 photons per pulse with a width less than 1 ns. Higher intensities may reduce the observed count rate due to saturation and can damage the detector. Therefore, for the calibration procedure, we recommend starting with the lowest dye concentration. With higher concentrations, the relation between mean count rate and dye concentration should stay linear. For measurements including lifetime analysis, saturated detectors introduce severe artifacts into the observed lifetime decay. Hence, we avoid measuring above a count rate one order of magnitude lower than the maximum count rate—500 kc/s in our case. In addition, the count rate should not exceed 1–2 % of the repitition rate of the laser. Thereby for each reference pulse, there is at most one detected photon to register and pileup effects are avoided.

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18. Most of the time, single-photon counting modules come with software with competent correlation functions which do all these steps for you. However, in some cases, we found it useful to analyze the data with home-written scripts, e.g., in MATLAB or python using the given formulas. 19. Binning the photons into μs-slots will typically result in just a few photons per time interval. The correlation curve will start therefore at μs time lags, which is enough to resolve the behavior of liposomes and most fluorescent dyes. The bin time could be reduced down to the resolution limit of the counting module, which will increase the calculation time. 20. The linear range is setup-specific and depends on the laser intensity. Usually, Eq. 20 stays true for N < 100. Plotting the obsered N against the concentrations given in Table 1 should allow to determine this linear range. References 1. Magde D, Elson E, Webb WW (1972) Thermodynamic fluctuations in a reacting system--measurement by fluorescence correlation spectroscopy. Phys Rev Lett 29(11):705–708. https://doi.org/10.1103/PhysRevLett. 29.705 € Widengren J et al (1993) 2. Rigler R, Mets U, Fluorescence correlation spectroscopy with high count rate and low background: analysis of translational diffusion. Eur Biophys J 22(3):169–175 3. Rigler R, Mets U (1993) Diffusion of single molecules through a Gaussian laser beam, vol 1921. Laser Spectroscopy of Biomolecules: 4th International Conference on Laser Applications in Life Sciences. SPIE 4. Schwille P, Meyer-Almes FJ, Rigler R (1997) Dual-color fluorescence cross-correlation spectroscopy for multicomponent diffusional analysis in solution. Biophys J 72 (4):1878–1886. https://doi.org/10. 1016/S0006-3495(97)78833-7 5. Elson EL (2011) Fluorescence correlation spectroscopy: past, present, future. Biophys J 101(12):2855–2870. https://doi.org/10. 1016/j.bpj.2011.11.012 6. Berland KM, So PT, Gratton E (1995) Two-photon fluorescence correlation spectroscopy: method and application to the intracellular environment. Biophys J 68(2):694–701. https://doi.org/10.1016/S0006-3495(95) 80230-4

7. Schwille P, Haupts U, Maiti S et al (1999) Molecular dynamics in living cells observed by fluorescence correlation spectroscopy with one- and two-photon excitation. Biophys J 77(4):2251–2265. https://doi.org/10.1016/ S0006-3495(99)77065-7 8. Walla PJ (2014) Single-biomolecule techniques, Vol 2. In: Modern biophysical chemistry, Wiley-VCH, Weinheim. pp 203–256. https:// doi.org/10.1002/9783527683505.ch09 9. Kim SA, Heinze KG, Bacia K et al (2005) Two-photon cross-correlation analysis of intracellular reactions with variable stoichiometry. Biophys J 88(6):4319–4336. https://doi. org/10.1529/biophysj.104.055319 10. Datta R, Heaster TM, Sharick JT, Gillette AA, Skala MC (2020) Fluorescence lifetime imaging microscopy: fundamentals and advances in instrumentation, analysis, and applications. J Biomed Opt 25 (7):1–43. https://doi.org/ 10.1117/1.JBO.25.7.071203 11. Walla PJ (2014) Basic Fluorescence Techniques, Vol 2. In: Modern Biophysical Chemistry, Wiley-VCH, Weinheim. pp 61–104. https://doi.org/10.1002/ 9783527683505.ch03 12. Mu¨ller CB, Loman A, Pacheco V et al (2008) Precise measurement of diffusion by multicolor dual-focus fluorescence correlation spectroscopy. EPL (Europhysics Letters) 83(4):46001. https://doi.org/10.1209/ 0295-5075/83/46001

Chapter 13 Fluorescence Lifetime and Cross-correlation Spectroscopy for Observing Membrane Fusion of Liposome Models Containing Synaptic Proteins Tobias Grothe and Peter J. Walla Abstract Watching events of membrane fusion in real time and distinguishing between intermediate steps of these events is useful for mechanistic insights but at the same time a challenging task. In this chapter, we describe how to use fluorescence cross-correlation spectroscopy and Fo¨rster-resonance energy transfer to resolve the tethering and fusion of membranes by SNARE proteins (syntaxin-1, SNAP-25, and synaptobrevin-2) as an example. The given protocols can easily be adapted to other membrane proteins to investigate their ability to tether or even fuse vesicular membrane. Key words FCCS, FRET, Fluorescence lifetime, Membrane fusion, Proteoliposomes, Small unilamellar vesicles, SNARE proteins

1

Introduction The membrane protein family of soluble N-ethylmaleimide sensitive factor attachment protein receptors (SNAREs) is the key player for all fusion events of the vesicular trafficking in eukaryotes [1, 2]. To investigate the mechanism and kinetics of membrane fusion by these proteins, it is useful to look at highly purified systems [3, 4]. The involved proteins can be isolated and reconstituted into artificial vesicles called liposomes. From there on, fluorescence-based bulk assays are commonly used. In a detailed example, the neuronal SNAREs syntaxin-1 and SNAP-25 are reconstituted into one population of liposomes and the corresponding R-SNARE synaptobrevin-2 into another. One of the liposomes is labeled with a lipid-anchored self-quenching fluorescent dye. When mixed, the liposomes bump into each other and the SNARE proteins form a membrane bridging trans-complex

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with the N-terminal end of their SNARE motifs [5]. Subsequently, the SNAREs act in a zipper-like fashion pulling the membranes closer together thus they fuse while forming the more stable cis-complex [6, 7]. The mixing of the lipids of both membranes then dilutes the concentration of the self-quenching dye, which increases the fluorescence. This versatile technique goes back to the early 1980s [8], is widely used in several variants and can easily be complemented by the addition of effectors as soluble proteins or relevant ions into the buffer medium to characterize their catalytic or inhibitory effects. As the fluorescence only increases when lipid mixing occurs, a major downside of this dequenching technique is that it cannot resolve intermediate states of the fusion events, such as membrane docking. This makes it difficult to get further mechanistic insights into the regulation of membrane fusion. The proximity of the membranes is an obvious prerequisite for fusion and often many auxiliary proteins are regulating membrane docking. In case of synaptic vesicle fusion, the docking of synaptic vesicles and subsequent priming of the fusion apparatus, while preventing fusion with the presynaptic membrane, is an essential stable intermediate to ensure the synchronous release of neurotransmitters [6, 9]. To resolve this intermediate state while keeping the simplicity of this assay, fluorescence cross-correlation spectroscopy (FCCS) can be used [10]. This method uses the same probes as in dequenching assays with two minor adaptations: first, instead of labeling one liposome population, both populations are labeled with either the donor or the acceptor of a fluorescent dye pair that can undergo Fo¨rster resonance energy transfer (FRET). Therefore, both liposome populations are fluorescently labeled and can be detected. Lipid mixing will be monitored by an increase of the FRET efficiency. The mixing of the lipid membranes brings the dye pairs close together, which makes FRET possible. Second, the liposomes are further diluted to nanomolar concentrations. With FCCS then both liposome populations are observed in a femtolitersized volume by two separate detectors (see Chapter 12). The fluctuations in both fluorescence signals are caused by single or few liposomes diffusing in and out of the detection volume. A high temporal correlation of both fluctuations is found, if both fluorophores co-diffuse. This is the case for docked and fused liposomes, but not for non-interacting. To distinguish between docked and fused liposomes, we use the FRET efficiency, which is only increased if lipid mixing took place. The FRET efficiency, however, is not measured by a change in the fluorescence intensity, as large intensity changes occur when observing only few fluorescently labelled liposomes. Instead, the fluorescence lifetime of the FRET donor is observed, which is a parameter independent of intensity fluctuations.

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In this chapter, we describe the simple and reproducible procedure of reconstituting neuronal SNARE proteins into small unilamellar liposomes (SUVs) with gravity-driven size-exclusion chromatography. We found this to be a reliable source of proteoliposomes that can be easily transferred to other membrane proteins. However, for different preparation techniques and further processing we want to point the reader to the publications of Rigaud and Le´vi [11] and Wang and Tonggu [12]. After dilution to nanomolar concentrations, the fluorescence signals of the SUVs are recorded, binned and correlated. Finally, the results are fitted to a model function and the key parameters are extracted. A more detailed description of the background of fluorescence cross-correlation spectroscopy and fluorescence lifetime analysis, as well as a description of the fluorescence microscope setup needed for the simultaneous monitoring of co-diffusion and membrane fusion can be found in Chapter 12.

2

Material Prepare all solutions with analytical grade reagents and ultrapure water. Prepare and store all reagents at room temperature unless indicated otherwise. Follow all waste disposal regulations when disposing waste materials. Use glass tools and work under the chemical hood when working with chloroform. Glass tools need to be cleaned without detergent.

2.1

Lipid Film

1. Phospholipids L-α-phosphatidylcholine (PC), L-α-phosphatidylethanolamine (PE), L–αphosphatidylinositol (PI) and L-α-phosphatidylserine (PS) as well as Cholesterol (Chol), solubilized in chloroform (for concentrations, see Table 1, rows 1 and 2) (see Note 1). Stored at
20  C (see Note 2). 2. Lipid-anchored fluorescent dyes Oregon Green™ 488 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine (OG-DHPE) and Texas Red™ 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine, Triethylammonium Salt (TR-DHPE), solubilized in chloroform (1 mg/mL, see Notes 1 and 2). Stored at
20  C. 3. Microliter glass syringes, gastight, for volumes up to 25 μL (see Note 3). 4. Vacuum-driven drying chamber. 5. Resuspension buffer: 20 mM HEPES/KOH, 150 mM KCl, TCEP 0.1 mM, 3% CHAPS, pH 7.4 (see Note 4). Stored at 4  C.

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Table 1 Calculation scheme for the lipid mixture Lipids

PC

PE

PS

PI

Cholesterol OG-DHPE TR-DHPE

Concentration stock solution (mg/mL)

25.0

25.0

10.0

10.0

10.0

Molecular weight (g/mol)

1.00

1.00

785.6 743.6 809.5 879.6 386.4

1086

1382

Molar ratio

50.0

18.5

10

10

10

1.50

1.00

μL needed

11.8

4.13

6.07

6.60

2.90

12.2

15.5

Final volume (μL)

50.0

Final lipid concentration (mM)

15.0

Lipid:protein ratio

300:1

Final protein amount (nmol)

2.50

2.2 Formation of Proteoliposomes

1. Resuspended lipid films for green and red SUVs (see Subheading 2.1). 2. Purified synaptobrevin-2 and ΔN-complex of Syntaxin-1A (183-288) and SNAP-25A and synaptobrevin-2(49-96): Purify as reported [10, 13]. Stored at
80  C. 3. Two glass chromatography columns, 3 mL column volume, 0.5  15 cm (see Note 5). 4. Sephadex® G-50 Superfine. 5. Equilibration and dilution buffer: 20 mM HEPES/KOH, 150 mM KCl, TCEP 0.1 mM, pH 7.4, 0.2 μm filtered and degassed (see Note 4). Stored at 4  C.

2.3 FCCS Measurement

1. FCCS-setup, adjusted and calibrated (see Chapter 12). 2. Proteoliposome solutions (see Subheading 2.2). 3. Equilibration and dilution buffer: 20 mM HEPES/KOH, 150 mM KCl, TCEP 0.1 mM, pH 7.4 (see Note 4). Stored at 4  C. 4. Coverslips of No.1 thickness.

3

Methods Carry out all procedures at room temperature unless specified otherwise.

3.1

Lipid Film

1. Remove the lipid solutions from the freezer and put them on ice to reduce the evaporation of chloroform.

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2. Wash the syringes three times with chloroform. For the green SUVs, transfer the specific amount of each lipid solution except TR-DHPE into a clean glass tube (see Table 1, μL needed). Wash the used syringe after each lipid transfer three times with chloroform. For the red SUVs, repeat the transfer with TR-DHPE instead of OG-DHPE into a second glass tube. Wash the syringes three times with chloroform when finished. 3. Pre-dry the lipid mixtures under a gentle stream of nitrogen under constant spinning (see Note 6). Remove chloroform traces in a vacuum-driven drying chamber at room temperature for 2 h (see Note 7). 4. Completely solve the lipid films in 50 μL resuspension buffer each by vigorous vortexing for a few minutes (see Note 8). Store the solved lipids at
20  C or use them immediately (see Note 9). 3.2 Formation of Proteoliposomes

1. Remove an aliquot each of 2.5 nmol of the purified synaptobrevin-2 and the ΔN-complex of Syntaxin-1A (183-288) and SNAP-25A and synaptobrevin-2(49-96) from the
80  C freezer and let them thaw on ice for a few minutes. Add synaptobrevin-2 to the red lipid solution and the binary complex to the green lipid solution. Let both solutions incubate for 30 min at room temperature. After 30 min, put the solution back on ice. 2. In the meanwhile, prepare the chromatography columns. Weigh two times 0.3 g of Sephadex® G-50 resin into a small beaker each, add 10 mL equilibration buffer, and let the resin soak for 15 min. 3. Rinse the chromatography columns with equilibration buffer. Slowly transfer the soaked resin into the columns with a plastic pipette and let it settle (see Note 10). Add buffer to keep the resin wet. 4. After the incubation of step 1 is finished, let the remaining buffer sink into the resins. Carefully add one of the proteinlipid mixtures onto each column without disturbing it. Let the mixture sink into the resin completely. Carefully add approximately 1 mL buffer onto the column without disturbing the resin (see Note 11). 5. The proteoliposomes are in the running fronts which can be identified by their red/green color. Collect the fronts in one or multiple plastic tubes as they elutes from the column (see Note 12). Label the proteoliposome solution and put it on ice (see Note 13). 6. Discard the resin and clean the columns by reverse flushing with water for a few minutes.

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3.3 FCCS Measurement

If you are working without automated laser blocking: Make sure that the laser beam is blocked. Inform your colleagues not to work on the laser table as long as you are working on the microscope. Wear appropriate eye and skin protection. 1. Prepare the excitation source by adjusting the wavelength and the laser power according to the used fluorophores (see Chapter 12, Note 15). 2. Dilute 1 μL of the synaptobrevin-2 proteoliposome solutions (R) 1:100 with dilution buffer. Put 20 μL onto the spectroscope (see Note 14). 3. Cover the objective to avoid background light disturb the measurement and block the excitation light. Shut down all background lights in the room. Turn on the detectors and open the laser shutter. 4. Start the data acquisition of single photon events tagged by their absolute time. Measure at least 3 times for 15 s (see Note 15). 5. Repeat steps 1–4 for the binary complex SUV solution (G). 6. Histogram the absolute time-tagged photon data of the three measurements of both proteoliposomes in 1-μs-steps. Autocorrelate the histograms with Eq. 1 and fit the correlation curves to Eq. 2. Extract hNi and calculate the corrected dilution factor for both proteoliposomes according to Eq. 3 (see Notes 16 and 17). See Figs. 1 and 2. hI Fl ðt Þ∙I Fl ðt þ τÞi D E
1 I Fl ðt Þ2

ð1Þ

1 1 1 ∙ qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ∙ hN i 1 þ ττD 1 þ w2 τ

ð2Þ

G ðτÞ ¼ G ðτ Þ ¼

τD

hN i∙

used dilution ¼ needed diluton factor needed hN i

ð3Þ

7. Repeat steps 1–6 with the new dilution factor until you have dilution factors which result in hNi ¼ 10 for both proteoliposome solutions. 8. Dilute 1 μL of the proteoliposome solutions by half of their final dilution factor resulting in a doubled concentration. Mix 20 μL of each solution into a fresh plastic tube and put 20 μL of the mixture onto the coverslip (see Note 14). 9. Cover the objective to avoid background light disturb the measurement. Shut down all background lights in the room. Turn on the detectors and open the laser shutter (see Note 18).

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Fig. 1 Typical Histogram data of the recorded photons in 1 μs bins for red and green channel. Per bin, only a few photons are recorded. Most bins show 0 or 1 photon as shown in the zoomed inlet of the first 50 bins

Fig. 2 Auto- and cross-correlations of histograms such as shown in Fig. 1. Red and green circles show the autocorrelations of the respective channels, calculated by Eq. 1. Blue circles show the cross-correlations of both channels, according to Eq. 4. Fits according to Eqs. 2 and 5 are shown as straight lines. The increasing cross-correlation amplitude over time is indicated by the arrows

10. Start the time-tagged data acquisition of single photon events. Make sure to acquire both the absolute time and the time relative to the laser pulse. Measure 240 times for 15 s to get an overall acquisition time of 60 min (see Note 19). 11. Histogram the absolute time-tagged photon data of each of the 240 measurements in 1-μs-steps. Autocorrelate and crosscorrelate the histograms with Eqs. 1 and 4. Fit the correlation curves to Eqs. 2 and 5 (see Notes 16 and 20). Extract hNi and calculate the relative amount of co-diffusing proteoliposomes with Eq. 6 (see Note 21). See Figs. 1 and 2.

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Fig. 3 Histogram of the arrival time of the recorded photons of the green channel relative to the laser pulse, binned by 164 ps. Green circles show the histogram after 1 and 10 min, indicated by the arrow. Fits by Eq. 7 are shown as straight lines



 R IG Fl ðt Þ∙I Fl ðt þ τÞ G X ðτÞ ¼  G   R 
1 I Fl ðt Þ ∙ I Fl ðt Þ

ð4Þ

1 1 ∙ qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi 1 þ ττD 1 þ w2 τ

ð5Þ

2 G X ,0 ∙hN G i∙hN R i ½%  ðhN G i þ hN R iÞ

ð6Þ

G X ðτÞ ¼ G X ,0 ∙

τD

N corr ¼

12. Histogram the relative time-tagged photon data of each of the 240 measurements by their smallest resolution accordingly to the used hardware. See Fig. 3. Fit the tail of the curve to Eq. 7 and extract τFl (see Notes 18 and 22). Do the same for the measurements of the green liposomes during steps 2–5. The mean of the latter is used as τFl, 0 in Eq. 8. Calculate the percentage of fused liposomes according to Eq. 8 (see Note 21). I Fl ðt Þ ¼ I Fl ð0Þ∙e
τFl ∙t N fused ðt Þ ¼

1 1 τFl ðt Þ
τFl,0 1 1 τFl ð60 min Þ
τFl,0

ð7Þ ð8Þ

13. Plot all data points of Ndocked and Nfused over time (see Notes 23 and 24 and Fig. 4).

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Fig. 4 Time course of docked (Ncorr, red) and fused (Nfusion, black) proteoliposomes, calculated by Eqs. 6 and 8. The error bars show the standard deviation of a 90-s time period for two independent liposome preparations. Open circles show the results of a control experiment where the interaction was inhibited with a soluble fraction of synaptobrevin 2. Reprinted with permission from Cypionka et al. [10]

4

Notes 1. We usually work with natural phospholipids from brain material but we also made good experiences with synthetic 1,2-dioleoyl-sn-glycero-3-phospholipids. Lipid solutions are commercially available or can be homemade by solving the lipid powder in chloroform. Work on ice to reduce chloroform evaporation. A determination of the lipid concentration by total phosphorus determination should be performed for homemade solutions [14, 15]. For cholesterol, which has no phosphor, enzymatic assays are commercially available. For other lipid compositions, the melting point of the lipid membrane should be calculated. All steps from resuspension to liposome formation (size exclusion chromatography) need to be performed at temperatures above the melting point of the final lipid membrane. In our case, the melting point is below room temperature. 2. Lipid solutions are stored in tightly sealed amber glass vials with screw caps. To reduce chloroform evaporation, the screw can be reinforced with a few layers of Teflon band.

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3. We recommend at least two syringes with a maximum volume of 10 and 25 μL. We cannot recommend common air cushion pipettes for handling chloroform solutions as chloroform is dripping off the pipette tip due to its low vapor pressure. 4. The resuspension buffer only differs from the equilibration or dilution buffer by the addition of 3% (w/v) CHAPS. A stock of 10 dilution buffer without TCEP (HEPES/KOH 200 mM, KCl 1500 mM, pH 7.4) can be stored protected from light at 4  C and diluted when needed. After the addition of TCEP and CHAPS later, check the pH again. We are using these buffers as a general buffer for neuronal proteins. They provide a physiological ion strength and use potassium ions instead of sodium ions as in neuronal cells. The formation of liposomes, however, is robust against buffer changes and can be performed in any buffer as far as we know. 5. Any other chromatography columns can be used. We recommend glass columns as they are reusable. We prefer a small diameter of the column as an even packing is easily achieved. 6. Holding the tube in a roughly 30 angle and slowly spinning it results in an evenly spread lipid film. The warmth of the hand accelerates the chloroform evaporation and a chill tube temperature is a good indicator that there is still chloroform to remove. 7. Usually, an evenly spread lipid film is dry after 30 min. The drying time can be extended to overnight. 8. It is of great importance to resuspend and solve all of the lipid film as any residual traces of the film would drastically change the lipid concentration and therefore the resulting lipid-toprotein ratio. Moreover, the solution needs to be clear: Unsolved lipids as multilamellar particles may be identified by sedimentation after centrifugation and should be dissolved by carefully adding detergent. 9. Although we usually prepare the lipid mixes fresh, it is possible to prepare a greater amount and store it aliquoted at
20  C to improve the day-to-day reproducibility. However, lipid molecules undergo hydrolysis in aqueous suspensions. Therefore, we avoid storing resuspended lipid mixes for more than a few weeks. 10. Usually, the gel settles in two separate fronts: a loosely packed front at the top and a little slower developing more tightly packed thus slightly darker front raising from the bottom. In the end, the gel should look completely evenly colored and without any air bubbles. To get rid of any air bubbles, the buffer-degassing step can be elongated.

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11. Too fast addition of buffer would disturb the running front. Dropwise addition out of a 200 μL-pipette works well in our hands. After the first 200 μL, the remaining buffer can be added a little faster. With the given amount of Sephadex G-50, the bulk volume of the column is approximately 1 mL. Thus, the proteoliposomes should elute after the added buffer has sunk completely into the gel. 12. Especially when establishing a new protocol, it is important to confirm detergent removal in the liposome fractions. The presence of most detergents can be easily indicated by blowing air into the solution and check for bubble formation. Ideally, an UV-absorption profile shows two distinct peaks, the first corresponding to the liposomes, the second to detergentmicells. Liposome fractions with incomplete detergent removal should not be used for further analysis, as the detergent will introduce severe artifacts. Instead, the length of the column can be increased. Alternatively, a second run through a fresh column can be applied. 13. We do not recommend snap freezing and thawing liposomes. SUVs without protein are stable for a few days at 4  C. However, fresh liposomes show a more uniformly distributed size and less aggregation, which improves the FCCS measurement. For proteoliposomes, the degradation effects can be even more severe. Additionally, the amount of functional protein will decrease over time. Therefore, we recommend using proteoliposomes on the same day of preparation. 14. Prepare the inverted microscope by carefully putting a droplet of ultrapure water on top of the objective lens without touching it. Then, put a single coverslip directly onto the objective. Place the sample over the lens onto the coverslip. Putting the coverslip directly on top of the objective is an easy way as it avoids any stages or holders for the coverslip. Therefore, the coverslip needs to sit directly on top of the lens with just a thin water layer in-between. 15. Usually, the FCCS-setup gives robust results with measurement times short as 10 s in a concentration range of 1 to 100 particles in the focal volume. A final concentration of hNi ¼ 10 (hNi ¼ 20 for the individual solutions before mixing) worked well in our lab. To get to this concentration, usually, a 1:100 dilution is a good starting point. 16. Most of the time, single-photon counting modules come with software with competent correlation functions which do all these steps for you. However, in some cases, we found it useful to analyze the data with home-written scripts, e.g., in MATLAB or python using the given formulas.

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17. Binning the photons into μs-slots will typically result in just a few photons per time interval. The correlation curve will start therefore at μs time lags, which is enough to resolve the behavior of liposomes. 18. This procedure usually results in missing the first 30 s of the fusion reaction, which can be tolerated because of the slow in-vitro fusion kinetics of SNARE proteins. 19. Measuring 240 times 15 s each is a trade-off between getting a sufficient time resolution and less noisy data that works well in our hands. Measuring 60 times for 60 s, for example, would give less data points but may decrease the noise. In our hands, there are always a few measurements unusable because of, e.g., dirt particles. These particles disturb the obtained correlation function by huge jumps in the fluorescence intensity, but can therefore usually be identified by looking at histograms with millisecond bin-times. Those measurements are considered as outliers and taken off the analysis. However, these events effect 60-s measurements as well as 15-s measurements. Imagine the extreme case of one outlier each 60 s would thus result in a completely messed up data acquisition in case of 60-s measurements, while 75% of the 15-s measurements would still be usable. Therefore, we recommend short measurement times, that, nevertheless, should be optimized for each setup. 20. Note that GX(0) is not equal to hNXi
1 like for autocorrelations. Instead hNXi can be calculated by multiplying GX(0)with hNGi and hNRi. 21. For the direct comparison with Nfused, Ndocked needs to be normalized by dividing through its value at the end of the reaction, Nfused(60 min). The docking of liposomes is measured straight forward due to the character of the correlation analysis, and can easily be adapted to new systems. The lifetime of the FRET donor, however, is more sensible to changes. For the given fluorophores in the given concentrations we have shown the linear dependence in various control measurements [10]. 22. The histogram is a convolution of the instrument response, timing jitter introduced by the detectors and the photon counting module for example, and the fluorescence signal. It is important to fit only the part of the histogram that shows the exponential decay of the fluorescence signal. One can easily identify this part by looking at the residuals plot of the fit. If the ends of the fitted region have higher residuals, a shortening of the fitted region should be considered. In a more sophisticated approach, the instrument response function is convoluted with the fitting function during the fit. A free software package for MATLAB was published for example by Enderlein and

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Erdmann [16]. The instrument response has to be recorded in advance, for example by measureing a scattering solution like diluted mild. 23. To be precise, the here presented assay does not show fusion but lipid mixing. However, for these SNAREs fusion is shown by, e.g., content mixing assays [17–19]. When adatping to other proteins, actual fusion should be proven by similar approaches but mostly these can be combined with FCCS as well. 24. When adapting this method to proteins that interact with each other or with lipids without further fusion, the docking of liposomes can be thought of as a steady state equilibrium of this binding. Measuring after the equilibrium state is reached for 5–10 min in total and averaging the data points over this time span is a useful alternative [20–22]. References 1. Kloepper TH, Kienle CN, Fasshauer D (2007) An elaborate classification of SNARE proteins sheds light on the conservation of the eukaryotic endomembrane system. Mol Biol Cell 18(9):3463–3471. https://doi.org/10.1091/ mbc.e07-03-0193 2. Jahn R, Scheller RH (2006) SNAREs—engines for membrane fusion. Nat Rev Mol Cell Biol 7(9):631–643. https://doi.org/10.1038/ nrm2002 3. Brunger AT, Weninger K, Bowen M et al (2009) Single-molecule studies of the neuronal SNARE fusion machinery. Annu Rev Biochem 78:903–928. https://doi.org/10.1146/ annurev.biochem.77.070306.103621 4. Weber T, Zemelman BV, McNew JA et al (1998) SNAREpins: minimal machinery for membrane fusion. Cell 92(6):759–772. https://doi.org/10.1016/s0092-8674(00) 81404-x 5. Vites O, Florin EL, Jahn R (2008) Docking of liposomes to planar surfaces mediated by transSNARE complexes. Biophys J 95(3):1295–1302. https://doi.org/10.1529/ biophysj.108.129510 6. Jahn R, Fasshauer D (2012) Molecular machines governing exocytosis of synaptic vesicles. Nature 490(7419):201–207. https://doi. org/10.1038/nature11320 7. Matos MF, Mukherjee K, Chen X et al (2003) Evidence for SNARE zippering during Ca2+
triggered exocytosis in PC12 cells. Neuropharmacology 45(6):777–786. https://doi. org/10.1016/s0028-3908(03)00318-6

8. Struck DK, Hoekstra D, Pagano RE (1981) Use of resonance energy transfer to monitor membrane fusion. Biochemistry 20(14):4093–4099. https://doi.org/10. 1021/bi00517a023 9. Martens S, McMahon HT (2008) Mechanisms of membrane fusion: disparate players and common principles. Nat Rev Mol Cell Biol 9(7):543–556. https://doi.org/10.1038/ nrm2417 10. Cypionka A, Stein A, Hernandez JM et al (2009) Discrimination between docking and fusion of liposomes reconstituted with neuronal SNARE-proteins using FCS. Proc Natl Acad Sci U S A 106(44):18575–18580. https://doi.org/10.1073/pnas.0906677106 11. Rigaud J-L, Le´vy D (2003) Reconstitution of membrane proteins into liposomes. In: Methods in enzymology, vol 372. Academic, pp 65–86. https://doi.org/10.1016/S00766879(03)72004-7 12. Wang L, Tonggu L (2015) Membrane protein reconstitution for functional and structural studies. Sci China Life Sci 58(1):66–74. https://doi.org/10.1007/s11427-0144769-0 13. Stein A, Radhakrishnan A, Riedel D et al (2007) Synaptotagmin activates membrane fusion through a Ca2+
dependent trans interaction with phospholipids. Nat Struct Mol Biol 14(10):904–911. https://doi.org/10.1038/ nsmb1305

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14. Fiske CH, Subbarow Y (1925) The colorimetric determination of phosphorus. J Biol Chem 66(2):375–400 15. Carles J (1956) Colorimetric microdetermination of phosphorus. Bull Soc Chim Biol 38(1):255–257 16. Enderlein J, Erdmann R (1997) Fast fitting of multi-exponential decay curves. Opt Commun 134(1):371–378. https://doi.org/10.1016/ S0030-4018(96)00384-7 17. Rizo J, Chen XC, Arac D (2006) Unraveling the mechanisms of synaptotagmin and SNARE function in neurotransmitter release. Trends Cell Biol 16(7):339–350. https://doi.org/10. 1016/j.tcb.2006.04.006 18. Zimmerberg J, Chernomordik LV (1999) Membrane fusion. Adv Drug Deliv Rev 38(3):197–205. https://doi.org/10.1016/ s0169-409x(99)00029-0

19. van den Bogaart G, Holt MG, Bunt G et al (2010) One SNARE complex is sufficient for membrane fusion. Nat Struct Mol Biol 17(3):358–364. https://doi.org/10.1038/ nsmb.1748 20. Hubrich BE, Kumar P, Neitz H et al (2018) PNA hybrid sequences as recognition units in SNARE-protein-mimicking peptides. Angew Chem Int Ed Engl 57(45):14932–14936. https://doi.org/10.1002/anie.201805752 21. Lin CC, Seikowski J, Perez-Lara A et al (2014) Control of membrane gaps by synaptotagminCa2+ measured with a novel membrane distance ruler. Nat Commun 5:5859. https:// doi.org/10.1038/ncomms6859 22. Vennekate W, Schroder S, Lin CC et al (2012) Cis- and trans-membrane interactions of synaptotagmin-1. Proc Natl Acad Sci U S A 109(27):11037–11042. https://doi.org/10. 1073/pnas.1116326109

Chapter 14 Synaptic Vesicle Pool Monitoring with Synapto-pHluorin Marc Dahlmanns and Jana Katharina Dahlmanns Abstract Synaptic vesicle exocytosis can be monitored with genetically encoded pH sensors in an in vitro fluorescence microscopy setup. Here, we describe a workflow starting with preparation of a primary cell culture to eventually estimate synaptic vesicle pool sizes based on electrical current-evoked vesicle release, which is reported by the synaptobrevin 2-EGFP fusion protein synapto-pHluorin (spH) that is expressed inside the synaptic vesicle membrane. The readily releasable pool and the recycling pool of synaptic vesicles are released separately in response to electrical stimulation. As vesicle reacidification is blocked in this experimental design, every released vesicle is counted only once. This spH-based approach offers different information than styryl-dye (FM dyes)-based approaches because the total synaptic pool size is measured by an alkalinization step. This provides a normalization constant for quantifying and comparing the synaptic vesicle pool sizes. In addition to investigation of basic research questions, spH-reported vesicle release is valuable to determine presynaptic effects of, e.g., pharmacological drug treatments. Key words Live-cell fluorescence imaging, Synapto-pHluorin, Synaptic vesicle pools, Readily releasable pool, Recycling pool

1

Introduction Synaptic transmission strongly relies on presynaptic release of neurotransmitter-filled clear vesicles. Synaptic vesicles are exocytosed and neurotransmitters diffuse into the synaptic cleft [1], where they bind a postsynaptic receptor. The remaining empty vesicle membrane is then recycled and new vesicles are formed through endocytosis and reacidification [2]. Synaptic vesicles are organized in three different functional pools (Fig. 1a): the readily releasable pool (RRP), the recycling pool (RP), and the resting pool (RP). Vesicles from the RRP are docked and fuse rapidly after stimulation [5], vesicles from the RP are released after the RRP during prolonged release [6], and the resting pool is stimulationinsensitive and does not get released [7]. To experimentally access these different pools, two common methods for exocytosis-dependent staining of synaptic vesicles are known: FM

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Schematic drawing of synaptic vesicle pools and synapto-pHluorin fluorescence: (a) Schematic drawing of the different synaptic vesicle pools in a synaptic bouton: reserve pool, A recycling pool B, and readily releasable pool docked to the active zone C. Reused with permission from [3]. (b) Schematic of a synaptic vesicle recycling cycle and the pH-dependent fluorescence: 1. Synaptic vesicles carry synapto-pHluorin proteins that do not fluoresce in their acidic lumen (blue); 2. During vesicle exocytosis neutral medium from the synaptic cleft enters the vesicle and the synapto-pHluorin proteins fluoresce; 3. After a vesicle reuptake, the membrane is still filled with neutral medium and emitting synapto-pHluorin; 4. Reacidification (blue) through vesicular ATPase rebuilds the vesicles acidic pH and causes a de-quenching of fluorescence. This fourth step is prevented in the described experiments with vesicular ATPase inhibitor Concanamycin A, so that once fluorescent proteins will stay fluorescent throughout the rest of the recording. Reused with permission from [4]

dyes and genetically encoded pH-sensitive fusion proteins. Whereas FM dyes are independent from genetic modifications and allow the acute staining of synapses to estimate their RRP and RP sizes, the estimation of the resting pool to calculate the total pool size remains impossible. In contrast, genetically encoded pH-sensitive fusion proteins such as synapto-pHluorin (spH; pH-sensitive EGFP fused to synaptobrevin2, located in the vesicular membrane) require transfection but allow for total pool sizes estimation. The vesicular lumen is acidic, which quenches spH-reported fluorescence (Fig. 1b). Exocytosis leads to a direct access of spH to the pH-neutral synaptic cleft, thereby increasing fluorescence [2, 8]. In the experimental setup, fluorescence remains elevated even during vesicle membrane recycling, because concanamycin A blocks the v-ATPase so that vesicles will not be reacidified. The size of RRP and RP are measured in the experiment by separately evoking their release through electrical field stimulation of the neuronal cultures (Fig. 2a). The total pool sizes of a given synapse is visualized by bath-application of an alkaline substance, such as ammonium chloride (Fig. 2b). In summary, our method provides the necessary information to culture and transfect primary neurons, to use livecell fluorescence microscopy for exocytosis imaging, and, lastly, we provide exemplary MATLAB codes to analyze the gathered microscopic data. In previous studies, we have shown that this method is for example suitable to investigate the mechanism behind antipsychotic medications ([3]; Fig. 3a) or to detect cancer drug-induced

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Fig. 2 Recording of synapto-pHluorin fluorescence during evoked release of different synaptic vesicle pools: (a) Exemplary raw recording of neurons during vesicle pool size measurement. Cells were transfected with synapto-pHluorin and treated with 80 nM haloperidol (HAL) prior to the experiment. Cells were perfused with imaging buffer containing 80 nM haloperidol during the entire recording as well; a: Raw image showing baseline fluorescence; b: Raw image showing fluorescence after electrical stimulation with 40 pulses at 20 Hz, which corresponds to the readily releasable pool vesicles being fluorescent; c: Raw image showing fluorescence after further electrical stimulation with 1200 pulses at 40 Hz, which corresponds to the recycling pool vesicles being fluorescent additionally; d: Raw image showing fluorescence during perfusion with imaging buffer containing additional 50 mM ammonium chloride, which corresponds to all synaptic vesicles being fluorescent; Scale bar ¼ 25 μm. Reused with permission from [3]. (b) Typical fluorescence trace from a vesicle pool size recording. Electrical stimulation times are shaded in gray; the time of ammonium chloride perfusion is shaded in blue; dashed horizontal lines mark the fluorescence levels of baseline, readily releasable pool, recycling pool, and total pool. Reused with permission from Dahlmanns DA

side effects on a synaptic level ([9]; Fig. 3b). As our recordings last around 5 min, many recordings may be performed in relatively short time, which yields high sample sizes and reliable results.

2

Materials

2.1 Preparation of Primary Hippocampus Cultures from Newborn Rats

1. Scissor. 2. Micro-scissor. 3. Newborn rats: 2–5 days old (see Note 1). 4. Sterile working bench. 5. Petri dishes: two small dishes plus one additional dish per animal.

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Fig. 3 Exemplary applications of synapto-pHluorin-based vesicle pool size measurement: (a) Effect of haloperidol treatment in vesicle recycling pool size in hippocampal cells after 1 h (P ¼ 0.0132, n ¼ 9 recordings) or 6 days (P ¼ 0.0052, n ¼ 14 recordings) and 14 days (P ¼ 0.0009, n ¼ 21 recordings) of treatment compared with vehicle (DMSO, control group). Effect of haloperidol treatment on readily releasable pool size in hippocampal cells compared with controls (P ¼ 0.0136, n ¼ 21 recordings). Reused with permission from [3]. (b) After a 24 h 10 μM erastin treatment (or control), the readily releasable pool (RRP) size (released upon electrical stimulation with 40 stimuli at 20 Hz) and the synaptic vesicle recycling pool (RP) size (released upon electrical stimulation with 1200 stimuli at 40 Hz) were measured relative to the total vesicle population (perfusion with 50 mM ammonium chloride) for each synapse. The bar plot shows means with standard errors of the mean. Number of experiments: n ¼ 32 for control, n ¼ 19 for 10 μM erastin; unpaired two-sided t-test, **** P < 0.0001 (RRP), P ¼ 0.512 (RP). Reused with permission from [9]

6. Cooling block. 7. Falcons, 20–50 ml. 8. Stereoscopic microscope. 9. Syringe. 10. Centrifuge. 11. Culture medium: Minimum Essential Medium supplemented with 1% Gibco™ B-27™ (Thermo Fisher Scientific). 12. Water bath. 13. Fetal calf serum (FCS, 10%).

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14. Trypsin (5 mg/ml): 0.11 g trypsin +22 ml digestion solution. Store 2 ml aliquots at 20  C. 15. DNase solution: 500 μl water, 25 mg DNase (1 mg/20 μl; 375,000 units). Store 10 μl aliquots at 20  C. 16. Phosphate Buffered Saline (PBS). 17. Hank’s Balanced Salt Solution (HBBS, Thermo Fisher Scientific). 18. Cell counting chamber. 19. Matrigel: 1 ml Matrigel, 49 ml MEM (see Note 2). 20. 12-well culture plates inserted with glass cover slips (18 mm, Corning Inc.), pre-coated with Matrigel: For coating, use Matrigel (4  C, 500 μl per 12-well plate) and place it into one well. Shake it. Take up the Matrigel and add it into the next well. Repeat until everything is coated with Matrigel. Incubate at 37  C for 1 h. Remove the rest of the Matrigel. 21. Glutamine solution: 0.2 M L-glutamine, 100 ml aqua dest. (store 5 ml aliquots at 20  C). 22. Cytarabine (AraC) solution (4 mM, in aqua dest.): store at 20  C. 23. Insulin solution (12.5 mg/ml): 100 mg insulin in 8 ml 10 mM HCl (2 ml 1 M HCl + 200 ml aqua dest.), filtered sterile. Store at 20  C. 24. Basic medium (500 ml): 500 ml MEM, 2.5 g Glucose, 0.1 g NaHCO3, 0.05 g Transferrin (after sterile filtration, store at 4  C; see Note 3). 25. Starting medium (500 ml): 500 ml basic medium, 50 ml FCS, 5 ml 0.2 M L-glutamine, 1 ml insulin solution (after sterile filtration, store at 4  C, see Note 3). 26. Growth medium (500 ml): 500 ml basic medium, 25 ml FCS, 1.25 ml 0.2 M L-glutamine solution, 10 ml B27™-supplement, 750 μl AraC (6 μM), 535 μl penicillin/streptomycin (100; after sterile filtration, store at 4  C; see Note 3). 27. Dissociation solution: 100 ml Hank’s solution, 12 mM MgSO4* 6 H2O (after sterile filtration, store at 4  C). 28. Digestion solution: 100 ml aqua dest., 137 mM NaCl, 5 mM KCl, 7 mM Na2HPO4, 25 mM HEPES, adjust pH to 7.2 at room temperature with 1 M NaOH and 1 M HCl (after sterile filtration, store at 4  C). 29. 2 BES-buffered saline (BBS) solution: in 500 ml H2O, 50 mM BES, 280 mM NaCl, 1.5 mM Na2HPO4. Adjust pH to 7.05 with 1 M NaOH and 1 M HCl. 30. Light microscope.

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2.2 Transfection of Primary Hippocampus Cultures (for Three 12-Well Plates)

1. Plasmid for synapto-pHluorin (PlasmidFactory, Bielefeld, Germany), 90 μl for three 12-well plates (see Note 4). 2. Neurobasal A: at 320–330 mOsm (18 ml for three plates). 3. Transfection solution (containing 60 μl spH-DNA, 60 μl CaCl2, 480 μl H2O, 2 BBS 600 μl, 10.8 ml NBA). For preparation of transfection solution for three 12-well plates, add DNA to the sterile water in one eppendorf cup (EPC). Then add drops of CaCl2, carefully mix it on the vortex. In a second EPC, vortex 900 μl BBS. During shaking, drop the DNA solution into the BBS-filled EPC. Keep it at room temperature and in the dark for 30 min. 4. HBSS for washing. 5. Sterile water. 6. Cell growth medium. 7. HBBS. 8. Water bath. 9. Vortex.

2.3 Fluorescence Live-Cell Imaging of Exocytosis

1. Inverted fluorescence microscope: optimally equipped with a perfect focus system to maintain focus during the superfusion of solution during the experiment. 2. 60 objective (or higher). 3. Dichroic mirror: cut off wavelength of 488 nm for EGFP (see Note 5). 4. Imaging chamber with parallel field electrodes (see Note 6). 5. Perfusion equipment: including a programmable stepper, a small glass capillary to direct the flow of the experimental solution above your field of view, a suction pump (see Note 7). 6. Stimulation equipment: a programmable stimulator and a stimulus isolator, connected to the imaging chamber. 7. Imaging buffer: 140 mM NaCl, 2.4 mM KCl, 10 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), 10 mM Glucose, 2 mM MgCl2, 2 mM CaCl2, Adjust pH to 7.5 using 1 M NaOH and 1 M HCl (see Note 8). Add mannitol to set osmolarity to 320  10 mOsm. 8. Ammonium chloride buffer: 140 mM NaCl, 2.4 mM KCl, 10 mM HEPES, 10 mM Glucose, 2 mM MgCl2, 2 mM CaCl2 and 50 mM NH4Cl, pH to 7.5 using 1 M NaOH and 1 M HCl (see Note 8). Add mannitol to set osmolarity to 320  10 mOsm. 9. Concanamycin A (80 nM), add freshly to imaging buffer before the experiment.

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2.4 Data Analysis of Fluorescence Images

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1. Imaging software to convert fluorescence images into tagged image format (tif) files (e.g., Andor Solis or NIS elements). 2. MATLAB (Mathworks Inc.). Download exemplary MATLAB code for processing vesicle pool size recordings at https:// github.com/janawrosch/VesiclePoolSizes.

Methods

3.1 Preparation of Primary Hippocampus Cultures (Four 12-Well Plates)

1. Decapitate the newborn rat pups (P2-P4, n ¼ 5–6, see Note 9) rapidly with a scissor. 2. Place small forceps in the eyes for fixation. Remove the skull above the brain with a micro-scissor, remove the entire brain and place it into a cooled petri dish filled with Hank’s solution with FCS. 3. Remove the cerebellum, separate the hemispheres, and place the brain under the stereoscopic microscope. 4. Turn the brain so that the basis is on top. Under microscopic assistance, carefully remove subcortical basal ganglia structures until you can see the hippocampus below the cortex. Cut the hippocampus at both of its ends and carefully roll it away from the remaining cortex structure. 5. Repeat steps 1–3 for every animal until you have collected enough tissue for your cell culture. 6. Collect all hippocampi in a small dish and clean them from other adhering structures. Cut them into smaller pieces and put them into a 15 ml falcon filled with Hank’s solution without FCS. Let them sink down and remove the supernatant. Add Hank’s solution (5 ml), let the brains sink and remove the supernatant again. 7. Add trypsin into a syringe, then add DNase (10 μl) and filter sterile. Use a big pipette to transfer everything into a small petri dish. Incubate at 37  C for 15 min. After that, transfer into a 15 ml falcon containing Hank’s solution with FCS. 8. Wash 2–3 times with Hank’s solution, as described in step 6. 9. After removing Hank’s supernatant, add 1 ml dissociation solution and DNase (10 μl, filtered sterile). Separate the cells carefully by pipetting up and down. Add Hank’s solution with 20% FCS to stop the reaction. 10. Centrifugate for 10 min with low speed. Take off supernatant and add 2.4 ml starting medium for resuspension. 11. Use a counting chamber to calculate cell number (see Note 9). 12. Plate the cells onto the Matrigel-coated cover slips (see Note 2). Use 50 μl cell suspension in the falcon per well, distribute it well by shaking it. Incubate at 37  C for 1 h. After 1 h, add 950 μl warmed starting medium per well. If cover slips start to float, press them down with a pipette.

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13. Renew growth medium at DIV 2 by removing 500 μl of old medium and adding 1 ml of fresh, warm growth medium. Check for correct adherence of the cells to the cover slip. Pack all prepared 12 well plates into cling film with a distilled water-soaked paper below and put them into an incubator (37  C, 5% CO2). 3.2 CalciumPhosphate Transfection of Primary Hippocampus Cultures

1. Use a light microscope to check all plates for vitality and contaminations. Use best plates for further transfection procedure. 2. Collect the medium of all plates and store it in a petri dish (one petri dish for each plate). Following that, directly add 500 μl NBA onto the cells so that they do not dry. 3. Add 3 ml fresh growth medium per petri dish filled with old growth medium to compensate pipetting losses. Incubate petri dishes and 12-well plates at 37  C in the incubator. 4. Slowly drip the previously prepared DNA solution (Subheading 2.2, item 3) into slowly vortexing NBA (16.2 ml). Let every drop fall as a single drop into the solution and keep solution in motion during the whole procedure. 5. Take off NBA from the cover slips and add DNA-containing NBA (500 μl). 6. Incubate at 37  C for 30 min. 7. After incubation, remove DNA-NBA and directly add HBSS (750 μl). Take off old HBSS and add new HBSS (750 μl). Then remove this HBSS and add 1500 μl of the previously saved medium from the petri dish. 8. Put the plates back in the incubator.

3.3 Fluorescence Live-Cell Imaging of Exocytosis

1. Perform the experiment between DIV 20 to 25. Prepare the imaging buffer with and without ammonium chloride and warm them in the incubator at 37  C. Add concanamycin A (80 nM) to the imaging buffer. Make sure the experiment is prepared so that recording can start directly after transferring the cells onto the microscope (correct channel for EGFP etc., see Note 5). 2. Place the cover slip with transfected cells into an imaging chamber filled with imaging buffer (containing concanamycin A) and transfer it to the microscope. Place all required equipment (glass pipette of the perfusion nearby the cells, suction pump etc., see Note 10). 3. Choose your field of view. Take a region with clearly separated EGFP-stained synapses and avoid big clusters of synapses as an increase in fluorescence leads to difficulties during analysis (Fig. 2a).

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4. Set up the illumination parameters, note how much filter is inside the light path, etc. Ensure that baseline fluorescence is high enough but also not too high, so that an increase in fluorescence later after alkalinization does not saturate the camera (Fig. 2b). 5. Record a baseline of around 2 min, allowing concanamycin A to enter the synapses and block the v-ATPase. 6. Stimulate your culture via field electrode (50 mA, 1 ms, alternating current) with 40 pulses at 20 Hz (duration 2 s), resulting in the release of the readily releasable pool (RRP). 7. Thirty seconds to one minute after RRP release, stimulate with 1200 pulses at 40 Hz to release the recycling pool (RP). 8. One minute after RP release, use the programmable perfusion system to switch from imaging buffer to ammonium chloride buffer (50 mM) for around 5–10 s, allowing illumination of the entire vesicle population. 9. Save the experimental data. 10. Remove the imaging chamber and repeat the experiment with the next cover slip until you have a sufficient sample size (usually around at least 10 good recordings). In case some part of the experiment failed (e.g., electrical field stimulation) the cultures cannot be reused, as concanamycin A prevents the fluorescence to go back to baseline after the experiment. Hence, always use a new cover slip. 3.4 Analysis of Fluorescence Images

1. Choose an image during the experiment that contains electrical stimulation, e.g., the release of the recycling pool. Take two to five frames at this time point (see Note 11), average them, and create a subtraction image from this stimulation image and a baseline image prior to electrical stimulation at the end of the 2 min period that allows concanamycin A wash-in (see Note 12). 2. Use a feature point detection algorithm to detect electrically excitable regions of interest (ROI)—these are the synapses (Fig. 4). 3. For each ROI that is chosen in step 2, calculate fluorescence curves. 4. Average the baseline fluorescence for each ROI, then subtract it from its fluorescence trace (see Note 13). 5. Normalize the fluorescence trace to ammonium chloridereported total vesicle pool. For this purpose, average fluorescence values corresponding to the total pool and divide the entire trace by average fluorescence (see Note 14).

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Fig. 4 Image segmentation process: The original image (left) is filtered with a Laplacian kernel (middle) and fluorescent synapses are detected (right); Scale bars span 20 μm. Reused with permission from [4]

6. Calculate the relative sizes of the vesicle pools. For RRP estimation, calculate two to five images before and after completion (1) of the 40 pulses/20 Hz-stimulation (RRP) and (2) of the 1200 pulses/40 Hz-stimulation (RP). Subtract the two averages around RRP release to get a percentage values corresponding to RRP size. Repeat this procedure for the RP (see Note 15) (Fig. 2b). 7. Average the RRP and RP sizes of all synapses in one recording.

4

Notes 1. This protocol is also suitable to be conducted with mouse tissue, which has the advantage of using transgenic mouse lines. 2. Alternatives: 0.1 mg/ml Poly-L-Lysin, 0.1 mg/ml Poly-LOrnithine, 50 μg/ml Laminin, or even without coating. 3. The basic medium can be prepared in larger amounts and be stored at 4  C for several months. Before the preparation of the rat brains, freshly prepare starting medium and growth medium because L-glutamine may degrade over time. 4. Beyond this synapto-pHluorin construct, also other genetical reporters can be used if they fulfill necessary parameters, e.g., if they do not affect normal vesicle recycling. pH-luorins have also been fused to other synaptic proteins as synaptophysin [10, 11]. 5. If using other fluorescent proteins with a different excitation and emission wavelength, use different mirrors, filters, etc. 6. Other setups such as multi electrode arrays may be used as well. Alternating polarity in parallel field electrodes prevents hydrolysis. In our setup, 50 mA sufficiently stimulated the cells,

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which may differ in your recording setup. Be careful as too high intensities would damage the cells. Keep your stimulating intensity constant across all recordings. 7. In your setup you need reservoirs for solutions, tubes, and a small glass pipette outlet. Tubes are best controlled with automatic vales to shut/open, and a variable clip allows calibration of the flow rate (0.5–1 ml/min). 8. These buffers can be prepared as 10 stock solution, stored at 4  C. If stored for a long time, filter it under sterile conditions and adjust osmolarity if necessary. 9. How many animals are required is dependent on your desired cell density in the culture well. In our lab, we made best experiences with six animals in four plates, as this leads to cultures with enough synapses in the field of view and avoids undesired clustering. 10. Be careful to position the glass pipette of the perfusion, the water level control, and the suction pump. None of them should point directly into your field of view. The perfusion pipette is best positioned outside but near the field of view, in direct opposition to the suction pump. This allows a constant linear flow across the synapses you record from. Also take care of the angle between your horizontal cover slip and your glass pipette, it should be around 45  to avoid de-attachment of your cells. 11. Averaging several images reduces the camera noise and gives more exact values, which is particularly important in case of the smaller readily releasable pool. 12. Subtracting of images clears the recording from most autofluorescent artifacts unresponsive synapses, spH surface expression [12], as constant fluorescence yields zero after subtraction. 13. Average five to ten fluorescence values to yield baseline fluorescence. Subtract this baseline level from the recorded values in the entire recording. If unstable, check if the fluorescence trace has a constant slope, so that a linear fit to the baseline may be applied. This is subtracted from the trace and allows for correction of constant fluorescence increase. Otherwise, discard the recording. 14. Normalization compensates for different synapse sizes and makes the relative vesicle pool sizes comparable. 15. Calculate the average fluorescence of a ROI before (e.g., pre-RRP) and after the electrical stimulation that releases RRP or RP (e.g., post-RRP). Then subtract pre-RRP and post-RRP to obtain the final size of the respective vesicle pool (in % of total pool size).

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References 1. Ryan TA, Smith SJ (1995) Vesicle pool mobilization during action potential firing at hippocampal synapses. Neuron 14(5):983–989 2. Wienisch M, Klingauf J (2006) Vesicular proteins exocytosed and subsequently retrieved by compensatory endocytosis are nonidentical. Nat Neurosci 9(8):1019–1027. https://doi. org/10.1038/nn1739 3. Amato D, Canneva F, Cumming P et al (2020) A dopaminergic mechanism of antipsychotic drug efficacy, failure, and failure reversal: the role of the dopamine transporter. Mol Psychiatry 25(9):2101–2118. https://doi.org/10. 1038/s41380-018-0114-5 4. Dahlmanns JK (2019) Development of quantitative functional analysis tools and models for neuroscientific applications. Dissertation; Friedrich-Alexander-Universit€at ErlangenNu¨rnberg urn:nbn:de:bvb:29-opus4-103312 5. Rosenmund C, Stevens CF (1996) Definition of the readily releasable pool of vesicles at hippocampal synapses. Neuron 16(6):1197–1207 6. Wilhelm BG, Groemer TW, Rizzoli SO (2010) The same synaptic vesicles drive active and spontaneous release. Nat Neurosci 13(12):1454–1456. https://doi.org/10. 1038/nn.2690

7. Atwood HL, Karunanithi S (2002) Diversification of synaptic strength: presynaptic elements. Nat Rev Neurosci 3(7):497–516. https://doi. org/10.1038/nrn876 8. Hua Y, Sinha R, Thiel CS et al (2011) A readily retrievable pool of synaptic vesicles. Nat Neurosci 14(7):833–839. https://doi.org/10. 1038/nn.2838 9. Dahlmanns M, Yakubov E, Chen D et al (2017) Chemotherapeutic xCT inhibitors sorafenib and erastin unraveled with the synaptic optogenetic function analysis tool. Cell Death Discov 3:17030. https://doi.org/10.1038/ cddiscovery.2017.30 10. Tagliatti E, Fadda M, Falace A et al (2016) Arf6 regulates the cycling and the readily releasable pool of synaptic vesicles at hippocampal synapse. eLife 5:10116. https://doi.org/10. 7554/eLife.10116 11. Royle SJ, Granseth B, Odermatt B et al (2008) Imaging phluorin-based probes at hippocampal synapses. Methods Mol Biol 457:293–303 12. Sankaranarayanan S, De Angelis D, Rothman JE et al (2000) The use of pHluorins for optical measurements of presynaptic activity. Biophys J 79(4):2199–2208. https://doi.org/10.1016/ S0006-3495(00)76468-X

Chapter 15 Imaging Neuropeptide Release at Drosophila Neuromuscular Junction with a Genetically Engineered Neuropeptide Release Reporter Yifu Han and Keke Ding Abstract Despite the important roles of neuropeptides in a variety of physiological processes, there still lacks a method to probe neuropeptide release events in vivo with satisfying temporal and spatial resolution. Neuropeptide Release Reporter (NPRR) was recently introduced as a novel genetically encoded indicator of neuropeptide release with a high temporal resolution and peptide specificity based on GCaMP molecule. Here we describe a method for using NPRR to image selective neuropeptide release at Drosophila neuromuscular junction in semi-dissected larvae. This method provides a quantitative analysis of activitydependent neuropeptide release as real-time changes in fluorescence intensity of GCaMP reporter with sub-second temporal resolution and single bouton specificity. Key words Drosophila, Neuromuscular junction, Dense core vesicle, Neuropeptide release, Calcium imaging

1

Introduction Neuropeptides participate in extremely diverse and complex physiological processes and behaviors. Visualizing and measuring the cellular location and kinetics of peptide release is crucial for understanding the release and modulation of neuropeptide in vivo. A series of techniques have been established for measuring neuropeptide release including biochemical probes and genetically encoded indicators. In Drosophila, a GFP indicator fused with Atrial Natriuretic Peptide (ANP) has been applied to study the transportation and release of dense core vesicle (DCV) at NMJs. However, all of these techniques lack either high temporal resolution or peptide specificity with in vivo preparations.

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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A novel neuropeptide indicator, NPRR was characterized as an optimal tool to detect the release of selective neuropeptides in intact neural tissues, with subcellular spatial, sub-second spatialtemporal resolution. Neuropeptides are synthesized as precursors in cell bodies, packaged in DCVs and transported to nerve terminals or dendrites for releasing. To probe DCV release events, NPRR combines two important modules as a fused protein: (1) a neuropeptide precursor that can be sorted into DCVs and transported to releasing sites, (2) a GCaMP reporter that can reflect the different chemical environments in DCV (pH ¼ 5.5–6.75, [Ca2+]~ 30 μM) and extracellular space (pH ¼ 7.3, [Ca2+] ~ 2 mM) [1, 2] as fluorescent changes (Fig. 1a). To ensure the NPRRs can be sorted and transported properly as the endogenous Drosophila neuropeptides, different engineering strategies and reporters have been evaluated [3]. The optimal designing was by substituting the GCaMP reporter for the neuropeptide precursor C-terminal domain that follows the final peptide (Fig. 1a). Two GCaMP6sbased NPRR have been investigated for measuring electrically triggered DCV release at Drosophila NMJs: a rat ANP reporter (NPRRANP) as a surrogate DCV indicator at fly NMJs and a Drosophila Tachykinin reporter (NPRRdTK) that functions as an endogenous fly neuropeptide. This protocol focuses on harnessing NPRRs to study the patterns of electrically triggered release of selective neuropeptides in semi-dissected Drosophila NMJs. Drosophila NMJs contain several native neuropeptides [4–6] and have been utilized as an excellent model to study the transportation and release of neuropeptides [7]. Furthermore, NMJs are large, individually identifiable and amenable to genetic tools for manipulating individual neuronal terminals. At muscle 12, there are limited neuronal terminals, including (1) one type Ib and one type Is containing predominantly synaptic vesicles (SV) and few DCVs and (2) one type III containing only DCVs [8] (Fig. 1b). Here we use specific GAL4 drivers to drive UAS-NPRRANP in type III motor neuron terminals at muscle 12. The motor neurons are stimulated with 70 Hz current pulses to induce DCV release at type III terminals. This method includes a calcium imaging process to detect an activity-dependent NRPP response. Electrically induced NPRR response shows a tri-phasic pattern, including three phases: initial rising, following undershoot and final recovery. We provide an analysis protocol to quantify the fluorescent changes representing each phase of NPRR response. The same method was applied on both NPRRANP and NPRRdTK to confirm that the DCV release at NMJs is stimulation intensitydependent and to show that type Ib and type III require distinct stimulation thresholds for DCV release. Thus, this method is a powerful approach for characterizing the releasing patterns of selective neuropeptides and can be generalized to different peptidergic neurons in the nervous system.

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Before stimulation

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After stimulation

Low pH, low Ca2+

Type

Ib

GCaMP Reporter domain

Is

II

III

+++ + + -

+ + + -

+++ +

bouton

SV +++ DCV + Glutamate + TypeIII-GAL4 -

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Type III – GAL4 > UAS-NPRRANP vGlut Bursicon Merge

b

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GFP

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GAL4

Fig. 1 Schematic design and expression of NRPP at Drosophila NMJs. (a) Schematic illustrating the principle of NPRRs (Neuropeptide Release Reporters). NPRR molecules in the DCV lumen (low pH/low calcium, left) exhibit increased fluorescence when released by fusion into the extracellular space (neutral pH/high calcium, right). NPRR fluorescent signal is expected to decay following diffusion into the synaptic cleft. New NPRR-containing DCVs are produced by synthesis and transport from the soma, not by recycling. NP neuropeptide. DCV dense core vesicle. SV synaptic vesicle. (b) Distinct motor neuron subtypes at the Drosophila NMJ (muscle 12/13) have different proportions of DCVs vs. SVs. Light gray circles, black lines, and dark gray shading represent boutons, inter-bouton intervals, and subsynaptic reticulum, respectively. The studies in this paper focus on Type III neurons (in red rectangles). (c) Expression patterns of NPRRANP using Type III-GAL4. Triple immunostaining labels GFP for NPRR (green), Bursicon (blue), and vGluT (red). Arrows indicate boutons in Type III neurons, which contain no vesicular glutamate transporter (vGlut). Note that anti-vGluT stains other types of motor neurons, which are not labeled by the Type III-specific driver used in this experiment. Anti-Bursicon stains exclusively type III motor neurons. Scale bar, 5 μm

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Materials Fly Strains

1. UAS-NPRRANP [3]. 2. Type III motor neuron driver: R20C11-GAL4 (BDSC: 48887).

2.2 Experiment Reagents

1. HL-3 solution: 70 mM NaCl, 5 mM KCl, 1.5 mM CaCl2, 20 mM MgCl2, 10 mM NaHCO3, 5 mM trehalose, 115 mM sucrose, 5 mM sodium HEPES, pH 7.2. 2. Dissection solution: HL-3 solution with zero calcium. 3. High glutamate solution: HL-3 solution with 10 mM glutamate. 4. Calcium imaging solution: HL-3 solution with 1 mM glutamate.

2.3

Equipment

1. Customized larval dissection chamber (Fig. 2a) [9]. 2. Suction glass micropipette (WPI, model: TW120-4). 3. Micro-electrode puller (Sutter Instrument, model: P-97) with pulling program: Heat ¼ Ramp - 15; Pull ¼ 10; VEL. ¼ 10; Time ¼ 200; Pressure ¼ 500. 4. Microforge (Narishige, model: MF-830). 5. Microfill (WPI, model: MF28G67-5) for filling suction electrode. 6. 10 ml syringe for filling up suction pipette and applying negative pressure.

a

b magnetic tape

magnetic pins

c

3” x 4” glass cover slip

d

Fig. 2 Drosophila larval dissection and tools. (a) Representative schematic (top) and picture (bottom) of the recording chamber used for larval dissection and electrophysiology with magnetic pins. (b) Representative suction pipette tip pulled by the program (detailed in Subheading 2.3) and polished to a diameter of ~5 μm. Representative images of a third-instar larvae before (c) and after dissection (d) using this system

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7. Scissors and forceps for dissection. 8. Master-9 pulse generator (A.M.P.I., Israel). 9. Iso-flex stimulator electrode.

(A.M.P.I.,

Israel)

with

stimulation

10. Nikon A1R confocal microscope with resonant scanner. 11. 60 APO 1.4 N.A. water immersion objective, 10 Apo air objective and 488 nm laser. 2.4

Software

1. NIS-Elements software. 2. ImageJ and ImageJ plugins [3]. 3. Customized MATLAB codes [3].

3

Methods

3.1 Drosophila Larval NMJ Preparation

1. Larval collection: Cross fly strains carrying type III-GAL4 (detailed in Subheading 2.1) with fly strains carrying UAS-NPRRANP (see Note 10). Grow flies on regular fly food for 7–8 days until third-instar wandering larvae appear. 2. Pull suction pipettes using the electrode puller with the program stated above (detailed in Subheading 2.3). Polish the tip of the pipette to achieve an inner diameter of ~5 μm or with a resistance of ~0.5 MΩ for sucking in a single motor neuron nerve bundle (Fig. 2b). 3. Fill in the suction pipette with HL-3 solution with microfill and syringe (detailed in Subheading 2.3). Load the suction pipette on the stimulation electrode holder (Fig. 3a). 4. Place a third-instar larva with dorsal body wall facing up in the customized dissection chamber, loosely hold the head and tail with magnet pins (Fig. 2c). 5. Bath the larva in ice cold dissection solution (see Note 1). Cut the dorsal body wall and excise nervous system (see Note 2). Pin down the cleaned body wall with four pins on the magnet dissection chamber (Fig. 2d; see Note 3).

3.2 Electric Stimulation

1. Incubate the larval body wall sample with ice cold high glutamate HL-3 solution (detailed in Subheading 2.2) for 5 min to desensitize postsynaptic receptors. Shift the sample to calcium imaging solution (see Note 4). 2. Orient the larval preparation on the microscope stage at optimal position for imaging (Fig. 3a). 3. Under 10 air objective, place the tip of the suction electrode to center of the field of view. Then lower down the suction pipette close to the larval preparation.

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Fig. 3 Schematic illustrating the resonant imaging and electrical stimulation configuration of the third-instar larval NMJ. (a) Picture of the electrophysiology setup, with electrodes placed to stimulate and record from the larval preparation. (b) Schematic of a dissected third-instar larvae with a detailed view of the muscle 12 NMJ after the stimulation electrode is properly positioned. A suction pipette is used to suck the severed motor nerve innervating the muscle segment to be recorded from. The presynaptic nerve terminal of type Ib motor neurons (gray) and type III motor neurons (green) are shown in a classical “beads on a string” bouton structure

4. Switch to 60 APO 1.4 N.A. water immersion objective. Then move muscle 12 at abdominal segment A2 or A3 close to the suction electrode tip (Fig. 3b) at the center of the visual field. 5. Apply a slight negative pressure from the syringe connected to the suction electrode holder to aspirate the nerve end in the pipette tip (see Note 5). 6. Program electrical stimulation with Master-9 pulse generator (Fig. 4a, b, see Note 6). 3.3

Calcium Imaging

1. Place the type III motor neuron terminal of muscle 12 to the center of field of view with 60 water immersion objective. Type III motor neuron terminals labeled by NPRRANP are visible with 488 nm laser excitation (Fig. 3b). 2. Set up time lapse imaging protocol. Set the duration of each time lapse imaging to 5 min with NIE Element and deliver four repetitive 70 Hz electrical stimulations following a 30 s delay during each imaging section. Perform imaging with a resonant

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Fig. 4 Protocol of electrical stimulation and sample selection of NPRR imaging data. (a) Representative repetitive electrical stimulations paradigm (above) and detailed pattern of 70 Hz pulse (below) for triggering NPRRANP release at type III terminals. (b) A representative program of 70 Hz pulse with Master-9 stimulator. (c) Representative parameters for movement correction in Image Stabilizer. (d) Representative selection of ROIs (yellow) selected for an individual NMJ sample with ImageJ. For details, see Subheading 3.4. Scale bar, 5 μm

scanner at a frequency of 1 fps with an acquisition resolution of 512  512 pixels. Acquire Z-stacks simultaneously with step length varying from 1 to 1.5 μm (see Note 7). 3. Acquire calcium imaging data from at least six independent NMJs from at least five animals. 3.4 Data Analysis and Interpretation

1. Perform maximum intensity projection on z-stacks images with NIS-Elements. Remove data with severe drifting or deformation due to muscle contractions from data set to avoid unreliable measurements. 2. Apply image registration using ImageJ with proper plugin (Image stabilizer) to correct planner movements from slight muscle contractions (see Note 8). 3. Load the corrected data in ImageJ. Manually select ROIs in ImageJ following image registration. Outline one individual ROI along the edge of single branch of type III motor neuron boutons for each sample using the selection tool in ImageJ (Fig. 4d). 4. Average the mean fluorescence intensity of all pixels of each ROI in ImageJ and copy the results to excel spreadsheets (see

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Fig. 5 Representative data showing the patterns of NPRRANP release at type III motor neuron terminals. (a) Normalized data from a representative experiment showing changes in NPRRANP fluorescence intensity (ΔF/F) in type III motor neurons at the larval NMJs evoked by electrical stimulation. BG: background. S1–S4: Stimulation trials 1–4. I1–I4: Inter-stimulation Intervals (ISIs) 1–4. Green line: ΔF/F averaged across all boutons in the field of view. Gray shading: s.e.m. envelope. Red bar: electrical stimulation trials (70 Hz). The three typical phases of the response are indicated in S4. The peak height of the response on the first trial is characteristically lower (see also (c)), and may reflect competition with unlabeled DCVs in the readily releasable pool. (b) Integrated NPRRANP ΔF/F values during trials S1–4 and intervals I1–4. A.U. arbitrary units. n ¼ 8. ***, p < 0.001. (c) Average NPRRANP ΔF/F peak heights for trials S1–4. n ¼ 8. *, p < 0.05. Plotted values in (b, c) are mean  s.e.m

Note 9). Transform arbitrary units of fluorescent intensity to normalized value by calculation: ΔF/F. Calculate baseline fluorescent intensity F by averaging the mean intensity of 30 s period before the first trial of electrical stimulation. 5. Plot normalized data over time. Represent NPRRANP release by the increase of fluorescence, triggered by 70 Hz electrical stimulations (Fig. 5a). Characterize the NPRR diffusion response as the negative peak amplitude during the interval of stimulations (Fig. 5a). Average peak values and negative peak values for each trial across different NMJ samples (Fig. 5b, c). 3.5 Representative Results

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For illustration, chose representative results that exhibit the temporal release pattern of NPRR (Fig. 5a, see Note 11).

Notes 1. Using zero calcium HL-3 solution for dissection could help prevent spontaneous muscle movement during dissection. 2. Cut through the dorsal body wall from tail to head and remove the gut and trachea using scissors and forceps. Excise all motor nerves between ventral nerve cord and body wall muscle to fully remove the brain and central nerve cord from the sample. The motor nerves should be remained sufficiently long to enable its proper positioning in the suction pipette.

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3. Stretch and pin the body wall to reduce muscle contraction during electric stimulation. A video protocol for larval dissection is detailed online [10]. 4. This step is aimed to minimize muscle contraction during electric stimulation [3]. 1 mM glutamate in the imaging solution is for keeping minimizing muscle contractions during imaging session. An alternative method to avoid muscle contraction is to substitute 100 μM 1-naphthylacetyl spermine trihydrochloride, which blocks the postsynaptic glutamate receptors with 1 mM glutamate in the imaging solution. Then the incubation step with 5 mM glutamate is dispensable [11]. 5. Make sure that the nerve end is tightly sealed within the tip of pipette. The motor neuron nerve that innervates the specific muscle segment mostly reaches out from the medial anterior corner of muscle 7 in that muscle segment (Fig. 3b). 6. 70 Hz electrical stimulation is required to repetitively induce significant DCV release at type III motor neurons terminals [1, 11]. Set the duration of single pulse to 1 ms. For inducing repetitive rising phase and following recovery phase of NPRR response, deliver 4 repetitive 70 Hz stimulations (20 s each) with intervals of 40s (Fig. 4a). Program the stimulation protocol in Master-9 pulse generator before experiments (Fig. 4b). Calibrate the stimulation intensity for each motor neuron before experiments. To provide enough current to induce action potentials in motor neurons, the stimulation intensity should be set to double the intensity needed for inducing muscle contractions on the iso-flex stimulator. 7. Z-stacks are required to cover the entire NMJ in the z direction with typical depth ranging from 10 to 20 μm. Sampling rate depends on the number of z steps and the number of pixels per section. Number of z steps (step length) should be tuned for each NMJ to maintain consistent sampling rate of 1 fps (one entire z-stack per second) among all samples. 8. Maximum projected data are saved as nd2 format, which can be directly imported to FIJI distribution of ImageJ. Imaging registration plugins (Image Stabilizer) should be pre-installed in ImageJ. After loading data files in ImageJ, select Plugins— Image Stabilizer. Set proper parameters for Image Stabilizer (Fig. 4c), then select OK. Save the corrected data as nd2 format after image registration. 9. After ROIs selected, select Analyze—tools—ROI Manager— Add. Then individual ROI is ready for analysis. In ROI manager, select More—Multi Measure—OK. Then mean intensity of the selected ROI is plotted over time. The mean intensity data can be copied into excel spreadsheet for further analysis.

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10. R20C11-GAL4 has been used for driving expression of UAS-operated genes exclusively in type III motor neurons at Drosophila NMJs [3, 12]. Other types of motor neurons, except type III, are glutamatergic, which express vesicular glutamate transporter (vGlut) at boutons. Thus, co-staining of vGlut (for type Ib, Is and II terminals) and GFP (for NPRR) and bursicon (for type III terminals) at NMJs could be used to verify the expression patterns of NPRRANP in type III terminals [3]. 11. Electrically triggered NPRR response exhibits a tri-phasic temporal pattern (Fig. 5a). An initial rising phase, which peaks at 0.5–5 s after the stimulation onset. NPRR release response is defined as the peak amplitude within the period of individual electrical stimulation. The latency indicates the kinetic difference between calcium influx and DCV exocytosis due to the loose association between DCVs and calcium channels [13]. A falling phase starts at 1–5 s before the end of each stimulation trial. This early decline might be due to the depletion of readily releasable DCVs. During the final recovery phase, NPRR signals fall below the baseline intensity and then recover to baseline level. This further decline reflects diffusion of released fluorescent NPRR molecules into the synaptic cleft [7], while the recovery process indicates DCV replenishments in synaptic terminals.

Acknowledgments Work in the Dickman lab is funded by a grant from the National Institutes of Health (NS091546). We thank Taylor Seid (California Institute of Technology), David Anderson (California Institute of Technology), and Dion Dickman (University of Southern California) for insightful discussions and comments. Competing Interests The authors declare no conflicts or other competing interests. References 1. Mitchell KJ, Pinton P, Varadi A et al (2001) Dense core secretory vesicles revealed as a dynamic Ca2+ store in neuroendocrine cells with a vesicle-associated membrane protein aequorin chimaera. J Cell Biol 155(1):41–51. https://doi.org/10.1083/jcb.200103145 2. Sturman DA, Shakiryanova D, Hewes RS et al (2006) Nearly neutral secretory vesicles in Drosophila nerve terminals. Biophys J 90(6): L45–L47. https://doi.org/10.1529/ biophysj.106.080978

3. Ding K, Han Y, Seid TW et al (2019) Imaging neuropeptide release at synapses with a genetically engineered reporter. elife 8:e46421. https://doi.org/10.7554/eLife.46421.001 4. Gorczyca M, Augart C, Budnik V (1993) Insulin-like receptor and insulin-like peptide are localized at neuromuscular junctions in Drosophila. J Neurosci 13(9):3692–3704. https://doi.org/10.1523/jneurosci.13-0903692.1993

Imaging Neuropeptide Release Using NPRR ˜ a LA (1995) A novel synaptic 5. Zhong Y, Pen transmission mediated by a PACAP-like neuropeptide in drosophila. Neuron 14(3):527–536. https://doi.org/10.1016/0896-6273(95) 90309-7 6. Anderson MS, Halpern ME, Keshishian H (1988) Identification of the neuropeptide transmitter proctolin in Drosophila larvae: characterization of muscle fiber-specific neuromuscular endings. J Neurosci 8(1):242–255. https://doi.org/10.1523/jneurosci.08-0100242.1988 7. van den Pol AN (2012) Neuropeptide transmission in brain circuits. Neuron 76 (1):98–115 8. Menon KP, Carrillo RA, Zinn K (2013) Development and plasticity of the Drosophila larval neuromuscular junction. Wiley Interdiscip Rev Dev Biol 2(5):647–670 9. Sullivan WA, Ashburner M, Hawley RS (2000) Drosophila protocols. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY

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10. Imlach W, McCabe BD (2009) Electrophysiological methods for recording synaptic potentials from the NMJ of drosophila larvae. J Vis Exp. https://doi.org/10.3791/1109 11. Levitan ES, Lanni F, Shakiryanova D (2007) In vivo imaging of vesicle motion and release at the Drosophila neuromuscular junction. Nat Protoc 2010(12):pdb.prot5529. https://doi. org/10.1038/nprot.2007.142 12. Koon AC, Budnik V (2012) Inhibitory control of synaptic and behavioral plasticity by octopaminergic signaling. J Neurosci 32 (18):6312–6322. https://doi.org/10.1523/ JNEUROSCI.6517-11.2012 13. Xia X, Lessmann V, Martin TFJ (2009) Imaging of evoked dense-core-vesicle exocytosis in hippocampal neurons reveals long latencies and kiss-and-run fusion events. J Cell Sci. https:// doi.org/10.1242/jcs.034603

Chapter 16 Imaging Synaptic Glutamate Release with Two-Photon Microscopy in Organotypic Slice Cultures Ce´line D. Du¨rst and Thomas G. Oertner Abstract The strength of an excitatory synapse relies on the amount of glutamate it releases and on the amount of postsynaptic receptors responding to the released glutamate. Here we describe a strategy to investigate presynaptic release independently of postsynaptic receptors, using a genetically encoded glutamate indicator (GEGI) such as iGluSnFR to measure synaptic transmission in rodent organotypic slice cultures. We express the iGluSnFR in CA3 pyramidal cells and perform two-photon glutamate imaging on individual Schaffer collateral boutons in CA1. Sparse labeling is achieved via transfection of pyramidal cells in organotypic hippocampal cultures, and imaging of evoked glutamate transients with two-photon laser scanning microscopy. A spiral scan path over an individual presynaptic bouton allows to sample at high temporal resolution the local release site in order to capture the peak of iGluSnFR transients. Key words Genetically encoded glutamate indicators (GEGIs), iGluSnFR, Synaptic transmission, Glutamate, Organotypic slice cultures, Two-photon imaging

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Introduction In the mammalian cortex, glutamatergic excitatory synapses convert presynaptic action potentials into chemical signals that are sensed by postsynaptic glutamate receptors. The smallest information transmission unit at chemical synapses is the release of a single synaptic vesicle filled with neurotransmitters. Its release occurs in a probabilistic manner due to the stochastic nature of the release machinery. Release probability is typically assessed from electrophysiological recordings at the soma of a postsynaptic neuron. However, a pair of connected neurons is usually connected by more than one synapse, precluding the isolation of single-synapse responses. Furthermore, postsynaptic changes such as phosphorylation, desensitization, saturation, lateral diffusion, and internalization of postsynaptic receptors further contribute to the variability of responses measured in the postsynaptic cell. Optical measurements

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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based on fluorescence microscopy are therefore ideal to assess synaptic physiology, as responses from individual synapses can be isolated despite other synapses on the same neuron being active simultaneously. Previous imaging studies used postsynaptic calcium transients as a read-out of synaptic efficacy. However, more recently, thanks to the development of genetically encoded glutamate indicators (GEGIs) with suitable affinity and kinetics to detect cleft glutamate, it is now possible to monitor glutamate transients at the synaptic cleft and thereby isolating presynaptic release of glutamate from postsynaptic parameters. iGluSnFR [1] is an intensity-based glutamate sensing fluorescent reporter constructed from E. coli periplasmic glutamate binding protein Gltl and a circularly permuted (cp) EGFP and is the first GEGI that gained great popularity among neuroscientists for investigating glutamate dynamics. In particular, it is well suitable for monitoring cleft glutamate transients thanks to its high fluorescence dynamic range and affinity (ΔF/ Fmax of 4.5 and Kd of ~4 μM). Further variants with varied biophysical properties displaying different kinetics, affinities, and emission profiles [2–4] enable researchers to select the most appropriate sensor depending on the biological question (bulk tissue vs. single synapse) and imaging system (camera, galvanometric laser scanner, or resonant scanner). Here, we introduce a methodology to investigate presynaptic release, using the GEGI iGluSnFR, to measure glutamate release in the synaptic cleft (Fig. 1). To conduct a singlesynapse study, sparse and cell-specific labeling are a prerequisite, making single-cell electroporation the method of choice for GEGI expression. We express the iGluSnFR in CA3 pyramidal cells of rat hippocampal organotypic slice cultures. We elicit individual action potentials in iGluSnFR-expressing CA3 cells and simultaneously monitor released glutamate from individual Schaffer collateral boutons in CA1. To accurately determine the amplitude of individual glutamate transients, we use spiral scans on individual boutons, which provide an increased temporal resolution and a better signalto-noise ratio.

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Materials Experiments involving organ explant must conform to relevant Institutional and National regulations. Follow all waste disposal regulations for disposing waste materials. All solutions have to be prepared using ultrapure water.

2.1 Organotypic Slice Cultures

1. Slice cultures from rodent hippocampus. 2. Slice culture medium: 394 mL MEM, 20% (v/v) Heatinactivated horse serum, 1 mM L-glutamine, 0.01 mg/mL Insulin, 14.5 mM NaCl, 2 mM MgSO4, 1.44 mM CaCl2,

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Fig. 1 Experimental workflow of cleft glutamate transient measurements with GEGIs in organotypic hippocampal slice cultures

0.00125% Ascorbic acid, 13 mM D-glucose. Medium has to be sterile filtered (0.2 μm pore size) and stored at 4
C for up to 4 weeks. 2.2 Single-Cell Electroporation 2.2.1 Electroporation Equipment

1. Upright microscope with motorized stage, CCD camera and IR-DIC (infrared differential interference contrast) or Dodt contrast. 2. 20 or 40 water immersion objective. 3. 4 zoom lens system (0.5–2.0 magnification range). 4. Vibration isolation table. 5. Axoporator 800A with HL-U pipette holder. 6. Motorized micromanipulators. 7. Microscope chamber made of a glass microscope slide (70  100  1 mm) onto which a Teflon ring (inner diameter ~ 35 mm, 2 mm high) is fixed with silicone aquarium sealant.

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8. Plastic syringe body (1 mL) as disposable mouthpiece, connected through a Luer 1-way stopcock and thin silicone tubing to the electrode holder. 9. Headphones. 10. Silver wire (~0.25 mm diameter). 11. Forceps. 12. Hot bead sterilizer. 13. Incubator (37
C; 5% CO2) with rapid humidity recovery and copper chamber. 14. Vertical micropipette puller. 15. Thin-walled borosilicate glass capillaries. 16. Tissue culture dishes (60 mm, sterile). 17. Ultrafree centrifugal filter units. 18. Slice culture transduction solution: 10 mM HEPES, 145 mM NaCl, 25 mM D-glucose, 2.5 mM KCl, 1 mM MgCl2, 2 mM CaCl2. Measure the pH using a pH-meter and adjust to pH 7.4 by adding NaOH or HCl. Measure the osmolality with a micro-osmometer and ensure that the osmolality is between 310 and 320 mOsm/kg. Solution must be sterile filtered (0.2 μm pore size), stored at 4
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1. Perfusion chamber with a quartz glass bottom. 2. In-line heating of the perfusion solution. 3. Vertical micropipette puller. 4. Micropipettes for whole-cell recording (borosilicate glass with filament, 1.5 mm O.D.). 5. Recording solution, artificial cerebrospinal fluid (ACSF): 25 mM NaHCO3, 1.25 mM NaH2PO4, 127 mM NaCl, 25 mM D-glucose, 2.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, pH adjusted to 7.4, ACSF has to be saturated with 95% O2 and 5% CO2. Osmolality should be between 310 and 320 mOsm/ kg. Store for max. 1 week at 4
C. Bubble with carbogen (95% O2, 5% CO2) during warm-up to prevent Ca2+ precipitation. Maintain perfusion reservoir at 34
C to prevent bubble formation in recording chamber. 6. K-gluconate-based intracellular solution: 10 mM HEPES, 135 mM K-gluconate, 0.2 mM EGTA, 4 mM MgCl2, 4 mM Na2-ATP, 0.4 mM Na-GTP, 10 mM Na2-phosphocreatine, 3 mM L-ascorbic acid, pH adjusted to 7.2 with KOH. Osmolality should be between 290 and 300 mOsm/kg. Solution has to be sterile filtered (0.2 μm pore size), stored at 20
C. Aliquot in Eppendorf tubes can be stored at 80
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Fig. 2 Setup for simultaneous imaging and somatic induction of individual action potentials. A Ti:Sapphire laser system with dispersion compensation is coupled in through an electro-optical modulator (EOM). Red and green fluorescence is detected through the objective and through the oil immersion condenser (NA 1.4) using two pairs of photomultiplier tubes. 560 DXCR dichroic mirrors and 525/50 and 607/70 emission filters are used to separate green and red fluorescence. Excitation light is blocked by short-pass filters. A high-power blue LED coupled in through the epifluorescence port excites the red morphological marker allowing visualization of the transfected cells. During epifluorescence illumination, PMTs are protected by a mechanical shutter (S) 2.3.1 Imaging Setup

1. Two-photon microscope based on an Olympus BX51WI microscope (Fig. 2). 2. Ti:Sapphire laser system with dispersion compensation. 3. Electro-optical modulator. 4. pE-4000 LED light source for epifluorescence. 5. 3 telescope, 5 mm scan mirrors. 6. Compound scan lens ( f ¼ 50 mm) [5]. 7. Dual camera port with IR mirror. 8. Water immersion objective (60, NA 1.0). 9. Two pairs of photomultiplier tubes. 10. Oil immersion condenser (NA 1.4) (see Note 1). 11. 560 DXCR dichroic mirrors. 12. 525/50 and 607/70 emission filters. 13. Short-pass filters. 14. NS45B shutter. 15. MP-285 micromanipulator. 16. Motorized stage. 17. Peltier-heating of the oil immersion condenser.

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18. Data acquisition boards. 19. Image acquisition software ScanImage [6]. 2.3.2 Electrophysiology Setup

1. pE-4000 LED light source for epifluorescence. 2. Infrared Dodt contrast. 3. Patch-clamp amplifier MultiClamp 700B. 4. Electrophysiology software Ephus [7] or Wavesurfer written in MATLAB. 5. Microelectrode manipulator.

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Methods Single-cell electroporation is the method of choice to achieve very sparse expression of glutamate sensors in organotypic culture of brain tissue (Figs. 3 and 4b). Sparse expression is necessary to follow the axon of a patch-clamped sensor-expressing neuron to a distal projection area such as CA1. Two-photon microscopy is typically used to detect weak functional signals deep in intact tissue. For expression in neurons, we clone the GEGIs behind the human synapsin 1 promoter and electroporate single neurons in organotypic slice cultures of rat hippocampus. GEGIs are relatively dim in

Fig. 3 Electroporation set up. An upright microscope with DIC (Differential interference contrast) and a low magnification objective (20 with a variable magnifier tube before the camera) is used for the electroporation procedure. The electroporation setup is under a laminar flow cabinet with HEPA (high-efficiency particulate arrestance) filter blowing clean air downwards to minimize contaminations. The microscope is on an active anti-vibration table, mechanically isolated from the vibrations generated by the fan in the filter

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Fig. 4 iGluSnFR expression in CA3 pyramidal cells in organotypic slice cultures of rat hippocampus. (a) Maximum intensity projection of two-photon images of CA3 pyramidal neurons expressing a GECI (here, iGluu [9]) 4 days after electroporation. The GEGI has its fluorescence mainly localized to the plasma membrane over the entire cell. The scale bar represents 50 μm. (b) Transmitted light image (dark field) of a transfected organotypic culture merged with a wide-field fluorescence image showing three transfected CA3 neurons co-expressing tdimer2 and iGluSnFR. The area for synaptic imaging is indicated (red dotted box). Scale bar represents 500 μm. (c) Maximum intensity projection of two-photon images of CA3 axons co-expressing tdimer2 and iGluSnFR in CA1 stratum radiatum (cells not identical to panel b). Scale bar represents 10 μm (left panel) and 1 μm (right panel). (d) Cartoon of a typical Schaffer collateral synapse showing a transfected CA3 bouton (b) synapsing onto a dendritic spine (s). The red fluorescent protein tdimer2 labels the axoplasm while the membrane-anchored iGluSnFR is exposed to the extracellular space. (e) Schematic representation of a hippocampal slice with transfected and patch-clamped CA3 pyramidal cell and the optically monitored area in CA1. (f) Action potentials are elicited in a transfected neuron by somatic current injections, and glutamate release is simultaneously optically recorded (GEGI fluorescence) from a single Schaffer collateral bouton in

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the absence of glutamate, making it difficult to focus on small structures such as axonal boutons. Therefore, we routinely use co-expression of a bright red fluorescent protein (tdimer2 or tdTomato) to label the cytoplasm and follow the axon through the tissue. 3.1 Culture Preparation

For slice cultures preparation from rodent hippocampus, refer to [8] which provides a detailed step-by-step description. Change the medium of the slice culture twice a week (see Note 2).

3.2 Choice of GEGI Variant

The affinity of the GEGI, expressed as Kd, must be appropriate for the expected glutamate concentration in the cellular or tissue environment. If the goal of the experiment is to distinguish synaptic failures (no glutamate release, no GEGI signal) from successes (stimulation-induced glutamate release), a very high affinity is desirable. If a linear response is important, for instance to estimate the number of vesicles released simultaneously, a slightly lower affinity might be advantageous.

3.3 Neuron Transfection 3.3.1 DNA and Plasmids Preparation

3.3.2 Electroporation

1. Sterile filter an aliquot (0.5 mL) of K-gluconate-based intracellular solution through a Millipore Ultrafree centrifugal unit by centrifugation at 16,000  g for several seconds in a table-top centrifuge at 4
C (see Note 3). 2. Remove the filter insert and add the plasmid DNA to the desired concentration. Use 40–50 ng/μL for pCI.syn. iGluSnFR (a gift from Loren Looger, addgene plasmid #41732) (see Note 4). For proper visualization of the transfected cells, mix a plasmid encoding for a red fluorescent protein (for example tdimer2; 20 ng/μL) with the GEGI plasmid to achieve co-expression (see Note 5). 1. Ground the electrodes and coat silver wire tips by bathing them in a saturated Cl3Fe solution for at least 30 min or overnight prior to first usage. 2. Pull electroporation pipettes using a micropipette puller. Pull thin-walled borosilicate capillaries to obtain a resistance of 10–15 MΩ when filled with the K-gluconate-based intracellular solution (see Note 6). 3. Back-fill an electroporation pipette with ~2 μL of plasmid mix solution for each slice to electroporate. Store the remaining

ä Fig. 4 (continued) CA1 showing a broad distribution of amplitudes and occasional failures. Images were acquired at 500 Hz at 34
C. Individual trials are classified as successes if the peak amplitude of the GEGI transient is >2σ (green traces) and as failures when the peak amplitude is 1/2 max (ΔF) are analyzed (ROI, region of interest) resulting in the extracted fluorescence transient (before bleach correction). (e) iGluSnFR response amplitude (black markers) of an individual bouton stimulated with single action potentials every 10 s. Note that response amplitudes were constant over time. A short time window before stimulation is analyzed to estimate imaging noise (white markers). The histogram of response amplitudes shows separation between failures of glutamate release (overlap with the baseline histogram) and successes. Figure and legend reproduced from [2] with permission from Nature Protocols 3.5 GEGI Signal Extraction

1. For amplitude extraction, linearize the spiral scans and display them as xt-plots (Fig. 5c). 2. To distinguish successful glutamate release events from failures, perform a statistical comparison of fluorescence fluctuations before stimulation (ΔF baseline, n ¼ 64 columns/locations) and response amplitude (ΔF response, n ¼ 64 columns/locations). A significant difference suggests a success, whereas lack of significance a failure trial. This classification is preliminary; the final failure analysis is performed after amplitude extraction (steps 8–11).

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3. Assign a new region of interest (ROI) for each success trial to compensate for eventual lateral drift between individual trials. 4. The spiral scan covering the entire bouton may hit the GEGI transient once or several times per line. Sort the pixel columns (i.e., spatial positions) according to the change in fluorescence (ΔF) in each column (Fig. 5d) (see Note 22). 5. Include columns with the largest signal (ΔF > ΔFmax/2) in the region of interest (see Note 23). 6. In failure trials, evaluate identical columns/locations than in the last success trial (see Note 24). 7. For each bouton, extract the characteristic decay time constant (τ) by fitting a mono-exponential function to the average GEGI fluorescence transient. 8. Estimate the glutamate transient amplitude for every trial by fitting an exponential function to the decay of the fluorescence transient (fixed τ, amplitude as the only free parameter). 9. For each trial, determine the imaging noise (σ) from the baseline of the extracted fluorescence time course (see Note 25). 10. Classify as “success” trials where average ΔF/F0 > 2σ above baseline imaging noise, otherwise classify as “failure.” 11. Calculate the release probability as the ratio of total number of successes/total number of stimulations (Fig. 5e).

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Notes 1. To minimize bleaching by excessive excitation, the microscope must be designed to detect emitted photons more efficiently. Using only the objective for fluorescence detection is not enough to achieve single-vesicle sensitivity. Condenser detection (oil immersion, 1.4 NA, large field of view) is essential for the success of single synapse experiments with many trials (Fig. 2). The condenser detection is sensitive to the refractive index of the immersion oil and correct (Ko¨hler) alignment. The oil immersion condenser must be permanently heated (day and night) if a recording temperature above room temperature is desired. This can be achieved with flexible heating pads or Peltier elements. 2. Avoid changing the medium prior to an experiment (~16 h) as this might affect cell excitability. 3. It is important that the DNA-containing solution is not passed through the Millipore Ultrafree centrifugal filter unit. 4. Avoid using very strong promoters (for instance CMV) as they might drive the expression of the plasmids to toxic levels.

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5. For different cell types and iGluSnFR variants, the final concentration and optimal expression time have to be determined empirically (typical range: 1–100 ng/μL with this protocol). 6. Ensure constant pipette resistance for reproducible expression. A too high pipette resistance leads to low expression of the plasmids, whereas a too low resistance causes extreme expression levels leading to toxicity problems. 7. Ensure that there are no bubbles in the back-filled pipettes. The back-filled pipettes can be kept in an upright position for up to 2 h before use. 8. Work on an electrophysiology microscope setup in a laminar flow box to prevent contamination (Fig. 3). 9. For reproducible expression level of the plasmids between different electroporation sessions, ensure a constant pipette resistance. 10. If the tip of the pipette seems dirty, withdraw the tip from the tissue and apply strong positive pressure using a syringe. Only once the tip is clean, penetrate again into the tissue while keeping a positive pressure. 11. For more reproducible expression levels of the plasmids between different cells, try to wait for the resistance to increase to a similar value before applying the pulse train. 12. The optimal settings may differ depending on the cell type to electroporate and the desired expression level. Ensure that the pulse is negative otherwise the negatively charged DNA will not be electroporated. 13. To ensure that the constructs are incorporated into the cells, add a fluorescent dye such as Alexa Fluor 594 to the DNA mix. After applying the pulse train to the target neuron, take a fluorescence image to ensure that the DNA solution and fluorescent dye were successfully electroporated. 14. If positive pressure is applied and the tissue surrounding the tip of the electroporation pipette does not appear to be gently blown away, the pipette might be clogged. Clean the tip (away from the tissue) by applying a strong positive pressure with a syringe or use a fresh pipette to pursue the electroporation. 15. To facilitate the search of a bouton that belongs to the stimulated cell, restrict the number of electroporated neurons to three or four in order to make the experiment more effective. 16. The optimal time for a cell to express a given plasmid before starting the experiment has to be determined empirically. 17. As new GEGIs variants (blue, yellow) [3, 4], different combination of GECIs and morphological marker colors can be used according to the available equipment (i.e., laser, filter sets).

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18. Eliciting individual action potential in whole-cell configuration of an individual transfected cell as opposed to field stimulation ensures that the GEGI transients originate from the imaged terminal and are not a consequence of glutamate spillover from neighboring terminals. 19. If a galvanometric scanning system is used, the microscope software has to support arbitrary line scans. The code for arbitrary line scans that we developed for our original study is now incorporated in the ScanImage software (Version 2016 and later). 20. Image only for the duration of the GEGI transients to minimize laser exposition. However, the number of trials that can be obtained from a single bouton is limited by the unavoidable bleaching of the indicator molecules and eventual destruction of the release machinery by toxic photoproducts (e.g., oxygen radicals). By using lower laser power, this number can be extended to 300 trials at the cost of a slightly lower SNR. Longer intervals between trials allow replenishing indicator molecules by lateral diffusion, but this strategy is limited by the need for stable whole-cell access during the entire experiment for reliable action potential generation. Furthermore, manual refocusing between trials can lead to substantial bleaching of the indicator. Automated refocusing between trials can minimize this. 21. If the slice is strongly drifting, lower the perfusion rate and ensure that the temperatures of the perfusion solution and of the imaging chamber of the microscope are stable to avoid thermal expansion during the experiment. 22. As opposed to a static ROI, this analysis procedure is robust against minute lateral drift of the tissue between trials and does not require a priori knowledge of the fusion location. As the point spread function (PSF) of a two-photon microscope is typically around 1.8–2 μm along the optical axis, the upper and lower surface of the bouton gets sampled simultaneously and therefore small drifts along the optical axis are unlikely to affect the amplitude of the iGluSnFR transients. 23. The threshold to select pixel columns to be kept in the region of interest is once adjusted according to the noise of the imaging system but should be kept constant for amplitude comparisons between different experiments. 24. If necessary, correct traces for GEGI bleaching prior to amplitude extraction. To this end, fit a mono-exponential or double exponential decay time constant to non-stimulated trials and subtract to the signal of individual trials.

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25. In some trials, the baseline fluorescence may show large fluctuations caused by green fluorescent vesicles passing through the axon. Remove these trials from further analysis.

Acknowledgments The authors thank Iris Ohmert and Sabine Graf for the preparation of organotypic cultures and excellent technical assistance and Mauro Pulin for critical comments on the manuscript. This study was supported by the German Research Foundation through Research Unit FOR 2419 P4, Priority Programs SPP 1665, and Collaborative Research Center SFB 936 B7. References 1. Marvin JS, Borghuis BG, Tian L et al (2013) An optimized fluorescent probe for visualizing glutamate neurotransmission. Nat Methods 10(2):162–170. https://doi.org/10.1038/ nmeth.2333 2. Durst CD, Wiegert JS, Helassa N et al (2019) High-speed imaging of glutamate release with genetically encoded sensors. Nat Protoc 14(5):1401–1424. https://doi.org/10.1038/ s41596-019-0143-9 3. Marvin JS, Scholl B, Wilson DE et al (2018) Stability, affinity, and chromatic variants of the glutamate sensor iGluSnFR. Nat Methods 15(11):936–939. https://doi.org/10.1038/ s41592-018-0171-3 4. Wu J, Abdelfattah AS, Zhou H et al (2018) Genetically encoded glutamate indicators with altered color and topology. ACS Chem Biol 13(7):1832–1837. https://doi.org/10.1021/ acschembio.7b01085 5. Negrean A, Mansvelder HD (2014) Optimal lens design and use in laser-scanning microscopy.

Biomed Opt Express 5(5):1588–1609. https:// doi.org/10.1364/BOE.5.001588 6. Pologruto TA, Sabatini BL, Svoboda K (2003) ScanImage: flexible software for operating laser scanning microscopes. Biomed Eng Online 2: 13. https://doi.org/10.1186/1475-925X2-13 7. Suter BA, O’Connor T, Iyer V et al (2010) Ephus: multipurpose data acquisition software for neuroscience experiments. Front Neural Circuits 4:100. https://doi.org/10.3389/fncir. 2010.00100 8. Gee CE, Ohmert I, Wiegert JS et al (2017) Preparation of slice cultures from rodent hippocampus. Cold Spring Harb Protoc 2017(2). https://doi.org/10.1101/pdb.prot094888 9. Helassa N, Durst CD, Coates C et al (2018) Ultrafast glutamate sensors resolve highfrequency release at Schaffer collateral synapses. Proc Natl Acad Sci U S A 115(21):5594–5599. https://doi.org/10.1073/pnas.1720648115

Chapter 17 Dynole 34-2 and Acrylo-Dyn 2-30, Novel Dynamin GTPase Chemical Biology Probes Jennifer R. Baker, Nicholas S. O’Brien, Kate L. Prichard, Phillip J. Robinson, Adam McCluskey, and Cecilia C. Russell Abstract This protocol describes the chemical synthesis of the dynamin inhibitors Dynole 34-2 and Acrylo-Dyn 2-30, and their chemical scaffold matched partner inactive compounds. The chosen active and inactive paired compounds represent potent dynamin inhibitors and very closely related dynamin-inactive compounds, with the synthesis of three of the four compounds readily possible via a common intermediate. Combined with the assay data provided, this allows the interrogation of dynamin in vitro and potentially in vivo. Key words Dynamin inhibitors, Endocytosis, Dynole, Dynole 34-2, Dynole-31-2, Acrylo-Dyn, Chemical synthesis, Molecular probes

1

Introduction Dynamin is a large GTPase. There are three dynamin genes, DNM1, 2 and 3, and the three protein families (DynI, II or III) have the same functional domains—GTPase or G-domain; a Bundle Signaling Element (BSE); a lipid binding pleckstrin homology domain (PH); a 4-helical bundle middle domain (MiD); and a proline/arginine-rich tail (PRD). The BSE/MiD interface is the main pivot point for major domain movements, being termed hinge 1 [1]. Via its PRD, dynamin is recruited by SH3 domaincontaining proteins to the plasma membrane, where it undergoes major conformational rearrangements from a tetramer to a helix at the neck of newly forming vesicles and brings about fission using lipid torsion within vesicle necks [2]. Dynamin’s role in actin dynamics is based on its ability to assemble as rings which bind short F-actin filaments and stimulate actin elongation [3]. Its actindependent cellular functions are diverse, e.g., vesicle fusion pores in

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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chromaffin cell exocytosis [4], HIV-1 internalization [5, 6], and actin repair in damaged kidney [7, 8]. The dynamin helix is induced by the tetramer binding to lipid membranes via the PH domain to oligomerize and drive membrane fission [9, 10]. This helix comprises thousands of monomers [11, 12], with helical turns of different pitch and diameters. This remarkable conformational flexibility is thought to represent transitions within the hydrolysis cycle of the G-domain [9, 13]. In contrast, dynamin rings are comprised of 26–32 monomers [14, 15], and their assembly can be induced by SH3 domain proteins [16–18] or short actin filaments [3]. The mechanisms of domain conformational changes are under intense investigation. In an effort to unravel some of the details of dynamin’s macromolecular assembly, we have developed an extensive dynamin chemical biology probe platform. Herein we describe the chemical synthesis of a selected series of analogs that target an unknown allosteric site within dynamin. We stress in the use of these protocols the benefits of (a) the use of a chemical scaffold match active and inactive control pair, and (b) the use of multiple chemical scaffolds in biological systems to increase the probability that the resultant phenotype is a direct function of dynamin inhibition in cells, and in some case in vivo. The Dynoles are a range of non-GTP competitive dynamin inhibitors that have been utilized as molecular probes for the understanding of the endocytic and non-endocytic roles of dynamin [18, 19]. The dynole series contains a number of potent inhibitors of both dyn I and II, exhibiting potent antimitotic effects, targeting CME and acting at the point of abscission to induce cytokinesis failure [20]. The dynoles were initially developed by structure simplification of a family of bisindolemaleimides, the dimethylamino variant of which exhibited modest dynamin I inhibition, IC50 ~ 100 μM (compound 1, Fig. 1). Further development of the indole-based analogs afforded Dynole®34–2 (compound 2), as a potent DynI inhibitor (IC50 ¼ 1.3  0.3 μM). Shortening of the octylamine tail to the

O

H N

N

O

N H N

1 DynI IC50 = ~100µM

Fig. 1 Bisindolemaleimide (compound 1), the lead dynole compound

Synthesis of Allosteric Dynamin Inhibitors O

H N

N

N

2 DynI IC50 = 1.3±0.3 µM

H N

O

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CN N

N

3 Inactive

Fig. 2 The active Dynole®34–2 (compound 2), and its inactive counterpart Dynole 31–2 (compound 3)

propyl variant removes dynamin inhibition, with Dynole®31–2 (compound 3) inactive at 300 μM drug concentration (active and inactive pair shown in Fig. 2). Synthesis of the active Dynole 34-2 (compound 2) is conducted via the preparation of the cyanoamide octylamine intermediate (compound 4), itself accessed by stirring ethylcyanoacetate (compound 5) with n-octylamine (compound 6) (Fig. 3a). The inactive compound 3 utilizes a propylamine cyanoamide (compound 7), afforded via the same protocol as compound 4 with Npropylamine (compound 8). These reactions are conducted under solvent-free conditions, with diethyl ether added after 1.5 h and the reaction mixture cooled at 18  C. Both analogs are isolated via a vacuum filtration. The formation of the common indole core (compound 11) is accomplished by N-alkylation of indole-3-carboxaldehyde (compound 9) with chloro-propyldimethylamine (compound 10). Coupling of this common core with the intermediates previously prepared, compound 5 or compound 6, via a Knoevenagel condensation, affords either the active compound 2 or inactive compound 3, respectively (Fig. 3b). Standard spectroscopic data for the final active and inactive compounds 2 and 3 is shown in Fig. 4. The Acrylo-Dyns were originally serendipitously discovered as part of an antifungal project [21]; routine testing against dynamin uncovered the dimethylamino-functionalised compound 12, which exhibited excellent inhibitory activity against dynamin I (IC50 ¼ 7.5  1.0 μM). Replacement of the phenyl dimethylamino with a benzyl-functionalised pyrrole afforded the inactive compound 13 (active and inactive pair shown in Fig. 5). The synthesis of these compounds is carried out via an Nalkylation and Knoevenagel condensation. The synthesis of active compound 12 is initiated via the same intermediate as the dynole reactions above: an N-alkylation of indole-3-carboxaldehyde (compound 9) with 3-dimethylamino-1-propyl chloride (compound 10). With intermediate compound 11 in hand, a base mediated Knoevenagel condensation with 3,4-dichlorophenylacetonitrile (compound 14) affords the desired active compound 12 (Fig. 6).

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Fig. 3 (a) Synthesis of the dimethylamino active intermediate compound 4 or the inactive compound 7. (b) Synthesis of the dimethylamino active compound 2 or the inactive compound 3

The inactive intermediate compound 15 is generated by an initial Knoevenagel condensation between 3,4-dichlorophenylacetonitrile (compound 14) and pyrrolecarboxaldehyde (compound 16); subsequent N-alkylation with benzyl bromide (compound 17) gives the desired compound 13 (Fig. 7). Standard spectroscopic data for the final active and inactive compounds 12 and 13 is shown in Fig. 8.

2

Materials Many of the chemicals used in these procedures are potentially hazardous, and therefore the material safety data sheets should be consulted for each chemical. Additionally, a lab coat, gloves, and eye protection should be used and, where possible, all operations must be carried out inside a fume hood. All solvents should be of reagent grade or higher.

Synthesis of Allosteric Dynamin Inhibitors

Compound (E)-2-cyano-3-(1-(3(dimethylamino)propyl)1H-indol-3-yl)-Noctylacrylamide (compound 2, Dynole active)

(E)-2-cyano-3-(1-(3(dimethylamino)propyl)1H-indol-3-yl)-Npropylacrylamide (compound 3, Dynole inactive)

1

H NMR (400 MHz, CDCl3) δ 8.64 (s, 1H), 8.40 (s, 1H), 7.86 (dd, J = 6.9, 1.5 Hz, 1H), 7.45 (d, J = 7.3 Hz, 1H), 7.35 – 7.29 (m, 2H), 6.15 (t, J = 5.1 Hz, 1H), 4.31 (t, J = 6.8 Hz, 2H), 3.42 (dd, J = 13.1, 7.1 Hz, 2H), 2.25 – 2.22 (m, 8H), 2.06 – 1.98 (m, 2H), 1.64 – 1.57 (m, 2H), 1.39 – 1.28 (m, 10H), 0.88 (t, J = 6.9 Hz, 3H); (400 MHz, DMSO-d6) δ 8.64 (s, 1H), 8.40 (s, 1H), 7.86 – 7.84 (m, 1H), 7.44 (d, J = 7.3 Hz, 1H), 7.30 (td, J = 7.1, 1.1 Hz, 2H), 6.20 (s, 1H, NH), 4.30 (t, J = 6.8 Hz, 2H), 3.39 (dd, J = 12.6, 6.0 Hz, 2H), 2.25 – 2.22 (m, 8H), 2.02 – 2.00 (m, 2H), 1.63 (dd, J = 14.5, 7.3 Hz, 2H), 0.98 (t, J = 7.4 Hz, 3H)

Fig. 4 Spectroscopic data for Dynole compounds

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C NMR (101 MHz, CDCl3) δ 162.0, 144.0, 136.4, 133.0, 128.6, 123.7, 122.4, 119.7, 119.0, 110.7, 110.2, 95.3, 55.9, 45.5 (2C), 45.0, 40.6, 31.9, 29.7, 29.4, 29.3, 27.8, 27.1, 22.8, 14.2

Melting point 95-97 °C

UPLC-MS LRMS: (ESI+) m/z: 409 (M+H, C25H37N4O, 100%); UPLC: Peak retention time: 1.581 mins; Area (%): 99

(101 MHz, DMSO-d6) δ 162.0, 144.0, 136.3, 133.0, 128.6, 123.7, 122.4, 119.6, 118.9, 110.6, 110.1, 95.2, 55.9, 45.5 (2C), 45.0, 42.1, 27.7, 23.0, 11.5

89-92 °C

LRMS: (ESI+) m/z: 339 (MH, C20H27N4O, 100%); UPLC: Peak retention time: 0.572 mins; Area (%): 100

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CN

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Fig. 5 Active propyldimethylamino (compound 12), inactive benzyl derivative (compound 13) Cl N

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Fig. 6 Synthesis of the dimethylamino active compound 12 Br Cl

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Fig. 7 Synthesis of the benzylated inactive compound 13 2.1

Reagents

1. n-Octylamine 99% [CAS no. 534-17-8]. 2. n-Propylamine >99% [CAS no. 107-10-8]. 3. Ethyl cyanoacetate >99% [CAS no. 105-56-6]. 4. Anhydrous magnesium sulfate (MgSO4) 99% [CAS no. 748788-9]. 5. 3-Dimethylamino-1-propyl chloride hydrochloride 96% [CAS no. 5407-04-5]. 6. Piperidine 99% [CAS no. 110-89-5]. 7. Cesium carbonate (Cs2CO3), 60–80 mesh, 99.9% [CAS no. 534-17-8]. 8. Potassium iodide (KI) >99.0% [CAS no. 7681-11-0]. 9. Sodium chloride >99% [CAS no. 7467-14-5]. 10. 3,4-Dichlorophenylacetonitrile 98% [CAS no. 3218-49-3]. 11. Pyrrole-2-carboxaldehyde 99% [CAS no. 1003-29-8].

Synthesis of Allosteric Dynamin Inhibitors

Compound (Z)-2-(3,4-Dichlorophenyl)3-(1-(3(dimethylamino)propyl)1H-indol-3-yl)acrylonitrile (Compound 12, AcryloDyn active)

(Z)-3-(1-Benzyl-1H-pyrrol2-yl)-2-(3,4dichlorophenyl)acrylonitrile (Compound 13, Acrylonitrile inactive)

1

H NMR (400 MHz, DMSO-d6) δ 8.40 (d, J = 19.1 Hz, 2H), 8.18 (d, J = 7.8 Hz, 1H), 8.10 (t, J = 1.2 Hz, 1H), 7.72 (d, J = 1.2 Hz, 2H), 7.62 (d, J = 8.1 Hz, 1H), 7.33 – 7.29 (m, 1H), 7.25 (dd, J = 11.0, 3.9 Hz, 1H), 4.37 (t, J = 6.7 Hz, 2H), 2.17 – 2.14 (m, 8H – 2xCH3 singlet overlaps with one of the CH2 peaks), 1.95 – 1.98 (m, 2H) (400 MHz, CDCl3) δ 7.57 (dd, J = 4.2, 0.8 Hz, 1H), 7.39 (d, J = 6.5 Hz, 2H), 7.38 – 7.36 (m, 2H), 7.32 (dd, J = 8.7, 5.8 Hz, 1H), 7.17 (d, J = 2.3 Hz, 2H), 7.08 (d, J = 7.0 Hz, 2H), 7.00 (dd, J = 2.4, 1.5 Hz, 1H), 6.41 (dd, J = 3.9, 2.7 Hz, 1H), 5.26 (s, 2H)

Fig. 8 Spectroscopic data for Acrylo-Dyn compounds

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C NMR (101 MHz, DMSO-d6) δ 135.8, 135.7, 135.2, 132.0, 131.0, 130.8, 129.9, 127.9, 126.2, 125.1, 123.1, 121.1, 119.4, 119.3, 110.8, 109.9, 99.6, 55.5, 45.1 (2C), 44.2, 27.3

Melting point 106-108 °C

UPLC-MS LRMS: (ESI+) m/z: 389 (M+H, C22H22Cl2N3, 100%); UPLC: Peak retention time: 1.860 mins; Area (%): 99

(101 MHz, CDCl3) δ 137.1, 135.2, 133.4, 132.1, 130.9, 129.8, 129.4 (2C), 128.4, 128.1, 127.7, 126.9, 126.4 (2C), 124.5, 118.4, 115.9, 111.0, 101.6, 51.5

115-117 °C

LRMS: (ESI+) m/z: 353 (MH, C20H15Cl2N2, 100%); UPLC: Peak retention time: 1.819 mins; Area (%): 100

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12. Benzyl bromide 98% [CAS no. 100-39-0]. 13. Benzyltrimethylammonium hydroxide (40% w/w aq. Sol.) [CAS no. 100-85-6]. 14. Transferrin, Alexa Fluor™ 488 conjugate (Tfn-A488) OR 594 conjugate (Tfn-A594), 4 μg/mL. 15. Dulbecco’s Modified Eagle’s Medium (DMEM). 16. High grade 100% DMSO (non-oxidized, ideally small sealed ampoules). 17. Paraformaldehyde, 4% in PBS. 18. 40 ,6-Diamine-20 -phenylindole dihydrochloride (DAPI). 2.2

Solvents

1. Ethanol (EtOH). 2. Acetonitrile (ACN). 3. Methanol (MeOH). 4. 10% hydrochloric acid solution (10% HCl). 5. Diethyl ether (ether, Et2O). 6. Ethyl acetate (EtOAc). 7. Dichloromethane (DCM). 8. Water. 9. Acidic wash solution for whole cell assay: 0.2 M acetic acid + 0.5 M NaCl, pH 2.8. 10. Phosphate-buffered saline (PBS).

2.3

Equipment

1. Weighing balance (e.g., Shimadzu AUW 220 D to 4 d.p.). 2. Magnetic stirrer with a temperature probe. 3. Rotary evaporator (e.g., Bu¨chi). 4. Vacuum system (e.g., Vacuubrand no. PC3001 VARIO). 5. Container for ice baths. 6. Teflon-coated magnetic stirrer bars. 7. Metal Drysyn heating blocks in various sizes (Asynt) or oil baths for heating. 8. Glassware: 50 mL round-bottomed flask, 100 mL roundbottomed flask, 250 mL round-bottomed flask, 10 mL conical flask, sintered glass funnel, Hirsch funnel, Bu¨chner flask, graduated cylinders, separatory funnel, and graduated pipettes. 9. Spatulas and tweezers. 10. Precoated silica gel 60F-254 plates (Merck, cat no. 1.05554.0001) and spotter for thin layer chromatography (TLC). 11. UV lamp (UVGL-58 handheld lamp, Australian Scientific) at 254 nm.

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12. Heat gun. 13. Flash chromatography system (Biotage Isolera) or a column for flash chromatography (supplied by custom blown glassware). 14. Empty silica columns (25 g Biotage, PN: BIOT-DLV-00000025) or silica 40–63 μM. 15. Biotage® Initiator+ microwave (optional). 16. Microwave vials for Biotage® microwave; 2–5 mL and 10–20 mL (optional). 17. Fibronectin-coated (5 μg/mL) 96-well glass plates.

3

Methods All procedures have been the subject to at least three independent trial by two researchers and the yields quoted are the average of three independent experiments. General spectroscopic details can be found in Note 1. The progress of reactions was monitored using a combination of LCMS and TLC. Subheading 3.1 refers to the Dynole compounds (Subheading 3.1.1, Dynole intermediates, Subheading 3.1.2 Dynole active compound 2 and Dynole inactive compound 3). Subheading 3.2 refers to the Acrylo-Dyn compounds (Subheading 3.2.1, Acrylo-Dyn active compound 12, and Subheading 3.2.2, Acrylo-Dyn inactive compound 13). Subheading 3.3 refers to the use of these probes in a typical whole cell assay.

3.1 Dynole Compounds 3.1.1 Synthesis of 2-Cyano-N-octylacetamide (Compound 4, Dynole Active Intermediate)

1. Weigh ethylcyanoacetate (compound 5, 0.94 mL, 8.8 mmol) and n-octylamine (compound 6, 1.46 mL, 8.8 mmol) and combine in a 10 mL conical flask equipped with a magnetic stirrer. 2. Allow to stir at room temperature for 1.5 h (see Note 2). 3. Add ether (25 mL) to redissolve precipitate and cool the flask in a freezer or ice bath for 30 min, or until a white precipitate forms. 4. Collect the white precipitate by filtration using a sintered funnel. Wash the solid with cold Et2O (2  5 mL, see Note 3). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. A second crop can usually be collected by evaporation of half the solvent and cooling (see Note 3). 5. Allow to dry under vacuum for at least 30 min. The product is usually obtained as a crystalline white solid (see Note 4).

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3.1.2 Synthesis of 2Cyano-N-Propylacetamide (Compound 7, Dynole Inactive Intermediate)

1. Weigh ethylcyanoacetate (compound 5, 5.33 mL, 50 mmol) and n-propylamine (compound 8, 4.11 mL, 50 mmol) and combine in a 25 mL conical flask equipped with a magnetic stirrer. 2. Allow to stir at room temperature for 1.5 h (see Note 5). 3. Add Et2O (5 mL) and cool the flask in a freezer or ice bath for 60 min, or until a white precipitate forms. 4. Collect the white precipitate by filtration using a sintered funnel. Wash the solid with cold Et2O (5 mL, see Note 6). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. A second crop can usually be collected by evaporation of half the solvent and cooling (see Note 6). 5. Allow to dry under vacuum for at least 30 min. The product is usually obtained as a crystalline white solid (see Note 7).

3.1.3 Synthesis of 1-(3(Dimethylamino)propyl)1H-indole-3-carbaldehyde (Compound 11, Dynole Common Intermediate)

Prepare as previously described [18]. 1. To a 250 mL round-bottomed flask fitted with a Teflon-coated stirrer bar, add the following reagents sequentially. Indole-3carboxaldehyde (compound 9, 2.000 g, 13.8 mmol), 3-dimethylamino-1-propyl chloride hydrochloride (compound 10, 2.40 g, 15.2 mmol, 1.1 eq.), potassium iodide (0.23 g, 1.38 mmol, 0.1 eq.), and cesium carbonate (11.24 g, 34.4 mmol, 2.5 eq.). 2. Add 100 mL acetonitrile and fit the flask with a reflux condenser. 3. Heat the reaction to reflux (100  C). 4. Stir the reaction at reflux for 18 h (it will turn from pale yellow/apricot to orange/red) (see Note 8). 5. Cool the reaction to room temperature, add Et2O (10 mL) and filter using a sintered funnel. 6. Wash the solid with acetonitrile (2  10 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. 7. Combine both the filtrate and washings and remove solvent via rotary evaporation. 8. Add EtOAc (30 mL) to the resultant dark oil and add to a separatory funnel. Add water (30 mL). Stopper the flask and gently invert a number of times. Ensure stopper is held in place, and pressure is vented periodically. 9. Drain the aqueous layer into a conical flask and repeat the washing with water (30 mL) twice more. Collect the organic layer in a conical flask.

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10. Dry the product with the use of MgSO4 (add dry MgSO4 until it is free-flowing, leave to sit for 5 min, then filter through a fluted filter paper into a round-bottomed flask). 11. Remove solvent via rotary evaporation and dry under high vacuum. Product is usually obtained as a golden brown oil (see Note 9). 3.1.4 Synthesis of (E)-2Cyano-3-(1-(3(dimethylamino)propyl)1H-indol-3-yl)-Noctylacrylamide (Compound 2, Dynole Active)

1. To a 100 mL round-bottomed flask fitted with a Teflon-coated stirrer bar, add 2-cyano-N-octylacetamide (compound 4, Dynole active intermediate, 5.713 g, 25.50 mmol) and 1-(3-(dimethylamino)propyl)-1H-indole-3-carbaldehyde (compound 11, Dynole active intermediate, 5.586 g, 24.29 mmol). 2. Add ethanol (50 mL) and piperidine (8 drops, catalytic). 3. Fit the round-bottomed flask with a reflux condenser and heat to reflux (100  C) for 18 h (see Note 10). 4. Once the reaction mixture has cooled to room temperature, evaporate the reaction mixture to dryness via rotary evaporation (see Note 11). 5. Load the oily solid onto silica by dissolving in dichloromethane (~100 mL) and adding silica (~1 tablespoon) and removing the solvent via rotary evaporation. 6. Purify the product via column chromatography: Load the product (now adsorbed onto silica) onto a bed of silica (approximately 10 times the thickness of the product/silica thickness). 7. The unreacted cyanoamide (compound 4) is eluted with 0–1% MeOH in DCM (the material will not fluoresce on a TLC plate under UV light, but can be visualized by dipping the plate in a permanganate stain). 8. Once all the cyanoamide intermediate has been eluted, increasing the MeOH concentration to 3% will elute the desired product (TLC r.f.: 0.33 in 10% MeOH:DCM). 9. Combine together all fractions containing the desired product and remove the solvent via rotary evaporation. 10. Dry the resulting yellow residue under high vacuum. The product is usually obtained as a yellow solid (see Note 12). 11. Conducting this reaction with 2-cyano-N-propylacetamide (compound 7) will afford the (E)-2-cyano-3(1-(3-(dimethylamino)propyl)-1H-indol-3-yl)-N-propylacrylamide (compound 3, Dynole inactive). This product is typically a yellow solid (see Note 13).

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3.2 Acrylo-Dyn Compounds 3.2.1 Synthesis of (Z)-2(3,4-Dichlorophenyl)-3-(1(3-(dimethylamino)propyl)1H-indol-3-yl)acrylonitrile (Compound 12, Acrylo-Dyn Active) Microwave Method

This reaction can be conducted via either microwave irradiation (Biotage® Initiator+) or batch methodology. The yield and purity are equivalent using either method.

1. To a 2–5 mL microwave vial fitted with a Teflon-coated stirrer bar, add 3,4-dichlorophenylacetonitrile (compound 14, 202 mg, 1.08 mmol) and 1-(3-(dimethylamino)propyl)-1Hindole-3-carbaldehyde (compound 11, 250 mg, 1.08 mmol). 2. Add ethanol (3 mL) and piperidine (4 drops, catalytic). 3. Seal the microwave vial and irradiate for 20 min at 120  C. 4. Once the vial has cooled to room temperature, cool at 0  C overnight (see Note 14). 5. Filter the reaction mixture through a sintered funnel. 6. Wash with cold EtOH (5 mL), then cold Et2O (5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. A second crop can usually be collected by evaporation of half the solvent and cooling (see Note 15). 7. Dry under vacuum for at least 30 min. Product is usually obtained as a bright yellow solid (see Note 16).

Batch Method

1. To a 50 mL round-bottomed flask fitted with a Teflon-coated stirrer bar, add 3,4-dichlorophenylacetonitrile (compound 14, 253 mg, 1.36 mmol) and 1-(3-(dimethylamino)propyl)-1Hindole-3-carbaldehyde (compound 11, 340 mg, 1.36 mmol). 2. Add ethanol (20 mL) and piperidine (4 drops, catalytic). 3. Fit the round-bottomed flask with a reflux condenser and heat to reflux (100  C) for 18 h (see Note 17). 4. Once the reaction mixture has cooled to room temperature, reduce the solvent by half and cool at 0  C overnight (see Note 18). 5. Filter the reaction mixture through a sintered funnel. 6. Wash with cold EtOH (5 mL), then cold Et2O (5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. A second crop can usually be collected by evaporation of half the solvent and cooling (see Note 15). 7. Dry under vacuum for at least 30 min. Product is usually obtained as a bright yellow solid (see Note 16).

Synthesis of Allosteric Dynamin Inhibitors 3.2.2 Synthesis of (Z)-2(3,4-ichlorophenyl)-3-(1Hpyrrol-2-yl)acrylonitrile (Compound 15, Acrylonitrile Inactive Intermediate)

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Prepare as previously described [4]. 1. To a 50 mL round-bottomed flask fitted with a Teflon-coated stirrer bar, add pyrrole-2-carboxaldehyde (compound 16, 2.010 g, 21.14 mmol, 1.05 eq.). 2. Add 10 mL water and heat the reaction mixture to 50  C while stirring vigorously. 3. Leave the reaction to stir at 50  C for 10 min, or until aldehyde dissolves. 4. Add 3,4-dichlorophenylacetonitrile (compound 14, 3.745 g, 20.13 mmol, 1 eq.) slowly to the vigorously stirring reaction mixture to form a suspension. 5. Stir the reaction at 50  C for a further 10 min. 6. Add benzyltrimethylammonium hydroxide (40 wt.% aq. solution) (7 mL, see caution) dropwise (see note). The reaction will change color from yellow/pale brown to bright yellow and a precipitate will form immediately (see Note 19). 7. Stir the reaction at 50  C for at least 4 h (see Note 20). 8. Filter the reaction mixture warm through a sintered funnel and wash with warm water (10 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. Dry under vacuum. 9. Recrystallize the solid from EtOH, by adding boiling EtOH to the solid until it just dissolves (see Note 21). 10. Place in the freezer overnight, or until crystals form. 11. Collect the resultant solid by filtering through a sintered funnel. Wash with cold EtOH (2  10 mL) or filtrate (2  10 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. A second crop can usually be collected by evaporation of half the solvent and cooling. 12. Dry under vacuum for at least 30 min. The product is usually obtained as a crystalline golden solid (see Note 22).

3.2.3 Synthesis of (Z)-3(1-Benzyl-1H-pyrrol-2-yl)2-(3,4-dichlorophenyl) acrylonitrile (Compound 13, Acrylonitrile Inactive)

1. To a 50 mL round-bottomed flask fitted with a Teflon-coated stirrer bar, add (Z)-2-(3,4-dichlorophenyl)-3-(1H-pyrrol-2-yl) acrylonitrile (compound 15, 435 mg, 1.65 mmol) and Cs2CO3 (3.06 g, 8.3 mmol, 5 eq.). 2. Add 20 mL ACN and stir at room temperature for 30 min. 3. Add benzyl bromide (compound 17, 500 μL, 4.12 mmol, 2.5 eq.) and KI (180 mg, 0.83 mmol, 0.5 eq.) 4. Fit the round-bottomed flask with a reflux condenser and heat to reflux (100  C) for 18 h (see Note 23).

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5. Allow the reaction to cool to room temperature and filter through a sintered funnel to remove the Cs2CO3. Wash with ACN (10 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred prior to filtering to enable adequate washing. 6. Transfer the filtrate to a round-bottomed flask and evaporate to dryness. 7. To the resulting dark green residue, add EtOAc (30 mL). Transfer to a separatory funnel and add water (30 mL). Stopper the flask and gently invert a number of times. Ensure stopper is held in place, and pressure is vented periodically. 8. Drain the aqueous layer into a conical flask and repeat the washing with water (50 mL) twice more. Wash the organic layer with brine (30 mL). Collect the organic layer in a conical flask. 9. Dry the product with the use of MgSO4 (add dry MgSO4 until it is free-flowing, leave to sit for 5 min, then filter through a fluted filter paper into a round-bottomed flask). 10. Remove solvent via rotary evaporation. Crude product is a green solid (see Note 24). 11. Recrystallize the solid from EtOH. 12. Collect the resultant solid by filtering through a sintered funnel. Wash with cold EtOH (2  10 mL). Dry under vacuum for at least 30 min. A second crop can be collected by evaporation of half the solvent and cooling. The product is usually obtained as a crystalline bottle green solid (see Note 25). 3.3 Biological Methods for Whole Cell Assay

1. Prepare stock solution of drug as a 30 mM solution in 100%, high grade DMSO (see Note 26). 2. U2OS cells are grown in fibronectin-coated (5 μg/mL) 96-well glass plates then serum-starved overnight (16 h) in serum-free DMEM (see Note 27). 3. Pre-incubate cells with different concentrations of (a) dynamin inhibitor, (b) inactive analogs, or (c) vehicle (DMSO) for 15–30 min prior to addition of 4 μg/mL Tfn-A488 or Tfn-A594 for a further 8 min at 37  C in the continued presence of the compound (see Note 28). 4. Cell surface-bound fluorescent Tfn is removed by incubating the cells with an ice-cold acidic wash solution (0.2 M acetic acid + 0.5 M NaCl, pH 2.8) for 10 min followed by its removal with an ice-cold PBS wash for 5 min (see Note 29). 5. Fix cells with 4% paraformaldehyde for 10 min at 37  C, and stain cell nuclei using DAPI.

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Notes 1. 1H NMR were recorded at either 400 or 600 MHz and 13C NMR were recorded at either 101 or 150 MHz, respectively, using a Bruker Avance 400 MHz or 600 MHz spectrometer in CDCl3 and DMSO-d6. LCMS was performed using an Agilent 6100 series single quadrupole LCMS using a mobile phase of 1:1 acetonitrile:H2O with 0.1% formic acid. The UPLC was fitted with a Zorbax SB-C18 Rapid Resolution HT 2.1  50 mm 1.8-Micron column. 2. The reaction mixture can be stirred longer or overnight without reduction in product purity. Follow the reaction progress by TLC or LCMS. 3. Over-washing with ether will result in a reduced yield due to loss of product to filtrate. 4. Average yield, 70%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 5. The reaction mixture can be stirred longer or overnight without reduction in product purity. Follow the reaction progress by TLC or LCMS. 6. A precipitate will not form with this product during the reaction, unlike with compound 4. This product is soluble in ether, so ensure the ether is cold, and use a minimal amount. Overwashing will result in a reduced yield due to loss of product to filtrate. 7. Average yield, 70%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 8. The reaction mixture can be stirred at reflux for several days without reduction in product purity. Follow the reaction progress by TLC or LCMS. 9. Average yield, 83%. This material should be stored at 0  C and used within a few days. 10. The reaction mixture can be left to stir at reflux for several days without reduction in product purity. 11. The reaction mixture can be left at 0  C for several weeks without reduction in product purity. 12. Average yield, 76%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 13. Average yield, 74%.

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14. The reaction mixture can be left at 0  C for several weeks without reduction in product purity. 15. Over-washing with EtOH and Et2O will result in a reduced yield due to loss of product to filtrate. 16. Average yield, 47%. 17. The reaction mixture can be left to stir at reflux for several days without reduction in product purity. 18. The reaction mixture can be left at 0  C for several weeks without reduction in product purity. 19. Adding the benzyltrimethylammonium too quickly will result in large lumps of solid. Adding the solution will allow a fine precipitate to form instead. If large lumps do form, they can be carefully broken up with a glass rod. Caution: The benzyltrimethylammonium hydroxide (40 wt.% aq. solution) is very hazardous upon eye contact (irritant), and can cause severe burns in case of skin contact. Ensure the SDS is consulted before using. It also has a noxious odor. Keep the reaction and all glassware that has come into contact with the liquid in the fumehood. The compound can be quenched with 10% HCl to remove all odor. 20. The reaction mixture can be stirred at 50  C for several days without reduction in product purity. 21. For recrystallization at 20 mmol scale, approximately 3–400 mL EtOH is required. 22. Average yield, 81%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 23. The reaction mixture can be left to stir at reflux for several days without reduction in product purity. 24. The crude solid can be left at 0  C for several weeks without reduction in product purity. 25. Average yield, 61%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 26. Sub-aliquots of the stock solution can be generated and frozen at 20  C for at least 6 months after at least two freeze-thaw cycles. Subsequent experiments can be performed using working solutions that are freshly prepared, immediately before use, by dilution of primary stocks into pre-warmed (37  C) cell culture medium such as DMEM (or 20 mM Tris–HCl, pH 7.4 for GTPase assay) to the desired final concentrations. Most cells can tolerate up to 1% DMSO, while dynamin GTPase assays in vitro can tolerate up to 3%. It is preferable

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to avoid serum or albumin in all drug solutions as most of these compounds bind albumin, reducing the compound access to cells. 27. This stage removes endogenous hormones and growth factors and reduces mitosis. A minimum of 2 h of serum-starvation is sufficient for sensitive CME assays, whereas overnight may be more operationally convenient. 28. We typically recommend use of 20–30 μM of each inhibitor, but a concentration curve for each active and inactive analog is the optimal experimental design, ranging from 1 μM to 60 μM. With many small molecules, there can be an issue of the compounds lifting the cells from the plates, confounding analysis by imaging microscopy. Lifting differs for each cell type we have investigated and can become pronounced at higher concentrations and should be monitored for each cell type. 29. This low temperature prevents further endocytosis and halts all known vesicle trafficking during this step. References 1. Faelber K, Posor Y, Gao S et al (2011) Crystal structure of nucleotide-free dynamin. Nature 477:556–560. https://doi.org/10.1038/ nature10369 2. Mattila JP, Shnyrova AV, Sundborger AC et al (2015) A hemi-fission intermediate links two mechanistically distinct stages of membrane fission. Nature 524:109–113. https://doi.org/ 10.1038/nature14509 3. Gu C, Yaddanapudi S, Weins A et al (2010) Direct dynamin-actin interactions regulate the actin skeleton. EMBO J 29:3593–3606. https://doi.org/10.1038/emboj.2010.249 4. Jackson J, Papadopulos A, Meunier FA et al (2015) Small molecules demonstrate the role of dynamin as a bi-directional regulator of the exocytosis fusion pore and vesicle release. Mol Psych 20:810–819. https://doi.org/10. 1038/mp.2015.56 5. Jones DM, Alvarez LA, Nolan R et al (2017) Dynamin-2 stabilises the HIV-1 fusion pore with a low oligomeric state. Cell Rep 18: 443–453. https://doi.org/10.1016/j.celrep. 2016.12.032 6. Aggarwal A, Hitchen TL, Ootes L et al (2017) HIV infection is influenced by dynamin at 3 independent points in the viral life cycle. Traffic 18:392–410. https://doi.org/10. 1111/tra.12481 7. Schiffer M, Teng B, Gu C et al (2015) Pharmacological targeting of actin-dependent dynamin oligomerisation ameliorates chronic kidney disease in diverse animal models. Nat

Med 21:601–609. https://doi.org/10.1038/ nm.3843 8. Sever S, Altintas MM, Nankoe SR et al (2007) Proteolytic processing of dynamin by cytoplasmic cathepsin L is a mechanism for proteinuric kidney disease. J Clin Invest 117:2095–2104. https://doi.org/10.1172/JCI32022 9. Sundborger AC, Fang S, Heymann JA et al (2014) A dynamin mutant defines a superconstricted pre-fission state. Cell Rep 8: 734–742. https://doi.org/10.1016/j.celrep. 2014.06.054 10. Kong L, Sochacki KA, Wang H et al (2018) Cryo-EM of the dynamin polymer assembled on lipid membrane. Nature 560:258–262. https://doi.org/10.1038/s41586-0180378-6 11. Stowell MH, Marks B, Wigge P et al (1999) Necleotide-dependent conformational changes in dynamin: evidence for a mechanochemical molecular spring. Nat Cell Biol 1:27–32. https://doi.org/10.1038/8997 12. Morlot S, Galli V, Klein M et al (2012) Membrane shape at the edge of the dynamin helix sets location and duration of the fission reaction. Cell 151:619–629. https://doi.org/10. 1016/j.cell.2012.09.017 13. Chen YJ, Zhang P, Egelman EH et al (2004) The stalk region of dynamin drives the constriction of dynamin tubes. Nat Struct Mol Biol 11:574–575. https://doi.org/10.1038/ nsmb762

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14. Hinshaw JE, Schmid SL (1995) Dynamin selfassembles into rings suggesting a mechanism for coated vesicle budding. Nature 374: 190–192. https://doi.org/10.1038/ 374190a0 15. Zhang P, Hinshaw JE (2001) Threedimensional reconstruction of dynamin in the constricted state. Nat Cell Biol 3:922–926. https://doi.org/10.1038/ncb1001-922 16. Ross JA, Chen Y, Muller J et al (2011) Dimeric endophilin A2 stimulates assembly and GTPase activity of dynamin 2. Biophys J 100:729–737. https://doi.org/10.1016/j.bpj.2010.12. 3717 17. Knezevic I, Predescu D, Bardita C et al (2011) Regulation of dynamin-2 assembly-disassembly and function through the SH3A domain of intersectin-1s. J Cell Mol Med 15: 2364–2376. https://doi.org/10.1111/j. 1582-4934.2010.01226.x 18. Robertson MJ, Deane F, Robinson PJ et al (2014) Synthesis of Dynole 34-2, Dynole

2-24 and Dyngo 4a for investigating dynamin GTPase. Nat Protoc 9:851–870. https://doi. org/10.1038/nprot.2014.046 19. Hill TA, Gordon CP, McGeachie AB et al (2009) Inhibition of dynamin mediated endocytosis by the dynoles™—synthesis and functional activity of a family of indoles. J Med Chem 52:3762–3773. https://doi.org/10. 1021/jm900036m 20. Chircop M, Perera S, Mariana A et al (2011) Inhibition of dynamin by Dynole 34-2 induces cell death following cytokinesis failure in cancer cells. Mol Cancer Ther 10:1553–1562. https://doi.org/10.1158/1535-7163.MCT11-0067 21. Tarleton M, Gilbert J, Robertson MJ et al (2011) Library synthesis and cytotoxicity of a family of 2-phenylacrylonitriles and discovery of an estrogen dependent breast cancer lead compound. Med Chem Commun 2:31–37. https://doi.org/10.1039/c0md00147c

Chapter 18 Synthesis of Phthaladyn-29 and Naphthalimide-10, GTP Site Directed Dynamin GTPase Inhibitors Cecilia C. Russell, Kate L. Prichard, Nicholas S. O’Brien, Adam McCluskey, Phillip J. Robinson, and Jennifer R. Baker Abstract Herein we describe the detailed synthesis of the dynamin inhibitors Phthaladyn-29 and Napthaladyn-10, and their chemical scaffold matched partner inactive compounds. Combined with the assay data provided, this allows the interrogation of dynamin in vitro and potentially in vivo. Key words Dynamin inhibitors, Endocytosis, Phthaladyn, Naphthalimide, Chemical synthesis, Chemical probe

1

Introduction Endocytosis is the process by which hormones, nutrients, and other extracellular materials enter the cell through budding into the plasma membrane. It is an intricate interplay requiring the orchestration of a wide array of proteins that assist in membrane invagination and the subsequent fission step that releases a new vesicle into the cell [1]. Dynamin GTPase is a protein critical to this process during clathrin mediated endocytosis (CME), a common endocytic mode using clathrin-coated vesicles. Dynamin’s key role occurs at a late stage of the process whereby it self-assembles into a helical collar or ring at the neck of the newly forming vesicle. Upon GTP hydrolysis a major conformational change in the dynamin helix is triggered resulting in bud fission and the generation of a free endocytic vesicle [1]. The pivotal role of dynamin in this process means that it is considered an attractive drug target across a myriad of human diseases including epilepsy [2–4], bone loss diseases [5, 6], kidney diseases [7, 8], and the spread of infectious diseases [9, 10].

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Dynamin is a large GTPase comprised of five domains: an amino-terminal G domain that binds and hydrolyzes GTP; a middle domain or a stalk region which is involved in self-assembly and oligomerization; a pleckstrin homology (PH) domain which interacts with the plasma membrane and induces the hemifission state; a GTPase effector domain (GED) which is also involved in selfassembly; and a proline-rich carboxy terminal domain (PRD) that interacts with SH3 domains in accessory proteins [11–15]. Mammals have three dynamins, dynI (found in neurons), dynII (ubiquitous), and dynIII (testes and lung) [13, 16]. Each gene can functionally replace the others for CME, but not for all other known cellular functions of dynamin [17–19]. We advise the use of dynamin inhibitors based on unrelated chemical scaffolds in the same experiment as this will potentially minimize the identification of off-target effects and reinforce conclusions that the observed effects are dynamin mediated. It is possible that some cell types may exhibit sensitivity towards the chosen dynamin inhibitor and show toxic effects [20]. Should this be the case, the use of another dynamin inhibitor, e.g., the allosteric compounds in the preceding protocol chapter: the Dynoles [21] and the Acrylo-Dyns. We have reported that dynamin inhibitors result in a cytokinesis block, but this requires at least 2 h to be effected, this is not an off-target effect. The use of confluent cell with minimal numbers of mitotic cells and short experiment durations, e.g., 30 min, mitigates such effects. Similar considerations apply to the best practice use of any small molecule enzyme inhibitor or siRNA studies [22]. In addition to the above advice, the use of inactive-active chemical scaffold matched pairs is strongly recommended when possible to mitigate any scaffold mediated off-target effects. All inactive compounds show no dynamin or Tfn CME activity in U2OS cells, but they have not been extensively examined in other cell lines. In the event of CME inhibition by the inactive analogs, we suggest the data from the active analogs should be excluded from any further analysis of the compound effects on dynamin inhibition. Further off-target effects can also be controlled for by the use of cells that do not express dynamin [23]. The Phthaladyns were developed from a virtual screening campaign using the ChemBridge and ChemDiversity library (800,000 compounds total) against an in-house developed homology model prior to the crystal structure of dynamin being solved [24]. The lead compound, 2-(2-([1,10 -biphenyl]-2-yl)-1,3-dioxoisoindoline5-carboxamido)-4-chlorobenzoic acid (compound 1) (Phthaladyn-1, Fig. 1) returned a Dynamin GTPase (Dyn) IC50 ca 400 μM, however the lead was amenable to a focused library design approach and a library of 34 compounds was prepared [24].

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Fig. 1 Lead compound for the first-generation Phthaladyns

Fig. 2 Structures of active (compound 2; also known as Phthaladyn-29) and inactive (compound 3) Phthaladyns

This library approach allowed rapid structure activity relationship data acquisition. Structure-based design, synthesis and subsequent optimization resulted in the development of compound 2 as the most active analog from this library synthesis (Dyn 1 IC50 4.58  0.06 μM; and known as Phthaladyn-29). Within the series of compounds analyzed, compound 3 was found to be inactive at concentrations up to 400 μM against Dyn 1. Thus compounds 2 and 3 are a structurally matched chemical biology probe active and inactive pair (Fig. 2). To access these compounds, a critical step is the synthesis of the common intermediate (compound 6). This intermediate is readily accessed from commercially available trimellitic anhydride (compound 4) and 2-amino-4-chlorobenzoic acid (compound 5) via an amide-forming reaction (Fig. 3a). Subsequent on-reaction with (4-aminophenyl)methanol (compound 7) and aniline (compound 8) yields the desired compounds 2 and 3, respectively (Fig. 3b) [24]. Standard spectroscopic data for the final compounds 2 and 3, as well as the intermediate compound 6, is shown in Fig. 4. The same virtual screening campaign that identified lead compound 1 was also responsible for the identification of 6-amino-2-(2-hydroxyethyl)-1,3-dioxo-2,3-dihydro-1H-benzo [de]isoquinoline-5-sulfonic acid (compound 9) with a Dyn IC50 ¼ 42  4.22 μM. Kinetic studies confirmed the GTP competitive nature of the lead analog in the naphthalimide series [25]. Structure-based design, synthesis and subsequent optimization resulted in the development of a library of 1,8-naphthalimide derivatives, which we call the Naphthaladyn™ series, with compound 10 (herein we call this Napthaladyn-10) being the most active (Dyn 1 IC50 13  6 μM). As with the phthaladyn series, we

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A

B

Fig. 3 Synthesis of the Phthaladyn common intermediate compound 6 (a), and synthesis of the Phthaladyn compounds 2 and 3 from the common intermediate compound 6 (b)

identified a structurally matched inactive control (compound 11, IC50 > 300 μM) for use alongside the dynamin active compound 10 (Fig. 5). Access to active and inactive naphthalimides is via two, one step syntheses, from commercially available starting materials. The active compound 10 is synthesized via a condensation between 3-hydroxy-1,8-naphthalimide (compound 12) and 4-aminobenzylamine (compound 13) (Fig. 6). The inactive naphthalimide (compound 11) is also afforded via a condensation reaction, this time between 1,8-naphthalimide (compound 14) and ethanolamine (compound 15) (Fig. 7). Both inactive and active naphthalimide compounds can be afforded via either an overnight reflux (see batch method in procedure, 3.2.1 and 3.2.2) or a microwave protocol (see batch method in procedure, 3.2.1 and 3.2.2). The product purity is comparable for both; however, reaction time is vastly reduced for the microwave method. As a laboratory microwave is not equipment that every lab has access to, we have reported both methods in this protocol. Standard spectroscopic data for the final compounds 10 and 11 is shown in Fig. 8.

Synthesis of Pthaladyn-29 and Napthalimide-10

Compound 4-Chloro-2-(1,3-dioxo1,3dihydroisobenzofuran-5carboxamido)benzoic acid (compound 6, phthaladyn intermediate) (see Note 2)

1

H NMR (400 MHz, DMSO-d6): δ 12.39 (s, 1H, NH), 8.75 (d, J = 2.1 Hz, 1H), 8.27 (d, J = 1.6 Hz, 1H), 8.14 (dd, J = 8.0, 1.7 Hz, 1H), 8.04 (d, J = 8.5 Hz, 1H), 7.85 (d, J = 8.0 Hz, 1H), 7.29 (dd, J = 8.5, 2.0 Hz, 1H) 4-Chloro-2-(2-(4(400 MHz, (hydroxymethyl)phenyl)- DMSO-d6): δ 12.51 (s, 1H, 1,3-dioxoisoindoline-5NH), 8.75 (s, carboxamido)benzoic 1H), 8.45 – 8.41 acid (compound 2, (m, 2H), 8.18 phthaladyn active) (d, J = 7.7 Hz, 1H), 8.08 (d, J = 8.4 Hz, 1H), 7.45 (dd, J = 23.4, 7.9 Hz, 4H), 7.35 (d, J = 8.4 Hz, 1H), 5.29 (s, 1H, OH), 4.57 (s, 2H) 4-Chloro-2-(1,3-dioxo-2- (400 MHz, phenylisoindoline-5DMSO-d6): δ 12.47 (s, 1H), carboxamido)benzoic 8.74 (d, J = 2.0 acid (compound 3, Hz, 1H), 8.44 – phthaladyn inactive) 8.40 (m, 2H), 8.18 (d, J = 7.7 Hz, 1H), 8.07 (d, J = 8.5 Hz, 1H), 7.57 – 7.54 (m, 2H), 7.49 – 7.45 (m, 3H), 7.33 (dd, J = 8.5, 2.1 Hz, 1H)

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C NMR (101 MHz, DMSO-d6): δ 169.5, 168.4, 167.6, 163.4, 141.8, 138.7, 136.9, 135.6, 132.9, 132.6, 129.9, 129.1, 127.2, 123.3, 119.5, 115.7

Melting point 253-254 °C

UPLC-MS LRMS: (ESI-) m/z: 344 (MH, C16H7ClNO6, 100%); UPLC: Peak retention time: 1.854 mins; Area (%): 100

(101 MHz, DMSO-d6): δ 169.4, 166.4, 163.3, 142.8, 141.5, 139.5, 138.6, 134.5, 133.8, 133.0, 132.4, 130.2, 127.1 (2C), 126.8 (2C), 124.2, 123.5, 121.3, 119.7, 116.3, 62.5

270-273 °C

LRMS: (ESI-) m/z: 449 (MH, C23H14ClN2O5, 100%); UPLC: Peak retention time: 0.839 mins; Area (%): 100

(101 MHz, DMSO-d6): δ 169.4, 166.29, 166.26, 163.3, 141.5, 139.5, 138.6, 134.5, 133.8, 133.0, 132.4, 131.8, 128.9 (2C), 128.3, 127.4 (2C), 124.2, 123.5, 121.3, 119.7, 116.2

273-276 °C

LRMS: (ESI+) m/z: 421 (M+H, C22H14ClN2O5, 100%); (ESI-) m/z: 419 (MH, C22H12ClN2O5, 100%); UPLC: Peak retention time: 1.766 mins; Area (%): 100

Fig. 4 Spectroscopic data for Phthaladyn compounds

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Fig. 5 Structures of lead naphthalimide (compound 9), the most active analog developed in this series (Naphthaladyn-10; compound 10) and inactive (compound 11)

Fig. 6 Synthesis of the active naphthalimide (compound 10)

Fig. 7 Synthesis of the inactive naphthalimide (compound 11)

2

Materials Many of the chemicals used in these procedures are potentially hazardous, and therefore the material safety data sheets should be consulted for each chemical. Additionally, a lab coat, gloves, and eye protection should be used and, where possible, all operations must be carried out inside a fume hood. All solvents should be of reagent grade or higher.

2.1

Reagents

1. Trimellitic anhydride chloride 98% [CAS no. 1204-28-0]. 2. 4-Chloro-2-aminobenzoic acid 97% [CAS no. 97–77-0]. 3. 4-Aminobenzyl alcohol 98% [CAS no. 623-04-1].

Synthesis of Pthaladyn-29 and Napthalimide-10

Compound 2-(4-Aminobenzyl)-5hydroxy-1Hbenzo[de]isoquinoline1,3(2H)-dione (compound 10, Naphthalimide active)

2-(2-Hydroxyethyl)-1Hbenzo[de]isoquinoline1,3(2H)-dione (compound 11, Naphthalimide inactive)

1

H NMR (400 MHz, DMSO-d6) δ 10.50 (s, 1H, OH), 8.22 (dd, J = 13.9, 7.7 Hz, 2H), 8.02 (d, J = 2.2 Hz, 1H), 7.72 (t, J = 7.8 Hz, 1H), 7.63 (d, J = 2.1 Hz, 1H), 7.07 (d, J = 8.3 Hz, 2H), 6.47 (d, J = 8.3 Hz, 2H), 5.04 (s, 2H), 4.98 (s, 2H) (400 MHz, DMSO-d6) δ 3.61 (m, 2H), 4.14 (t, J = 6.6 Hz, 2H), 4.81 (t, J = 6.0 Hz, OH), 7.85 (dd, J = 7.5, 8.1 Hz, 2H), 8.44 (dd, J = 0.9, 8.1 Hz, H2), 8.47 (dd, J = 0.9, 7.5 Hz, 2H)

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C NMR (101 MHz, DMSO-d6) δ 163.5, 163.2, 156.2, 147.9, 133.3, 132.6, 129.1 (2C), 127.6, 127.4, 124.4, 123.4, 122.1, 122.0, 121.9, 115.8, 113.6 (2C), 42.5

Melting point 255-257 °C

UPLC-MS LRMS: (ESI+) m/z: 319.1 (M+H, C19H15N2O3, 100%); (ESI-) m/z: 317.1 (MH, C19H13N3O3, 100%); UPLC: Peak retention time: 1.202 mins; Area (%): 100

(101 MHz, DMSO-d6) δ 163.5 (2C), 134.2 (2C), 131.3, 130.6 (2C), 127.4, 127.2 (2C), 122.2 (2C), 57.8, 41.8

174-175 °C

LRMS: (ESI+) m/z: 242 (M+H, C14H12NO3, 100%); UPLC: Peak retention time: 0.786 mins; Area (%): 100

Fig. 8 Spectroscopic data for naphthalimide compounds

4. Triethylamine (TEA) 99.5% [CAS no. 7087-68-5]. 5. Anhydrous magnesium sulfate (MgSO4) 99% [CAS no. 748788-9]. 6. Aniline 99.5% [CAS no. 62-53-3]. 7. 4-Aminobenzylamine 98% [CAS no. 4403-71-8]. 8. 1,8-Naphthalimide 99% [CAS no. 81-83-4]. 9. 3-Hydroxy-1,8-naphthalic anhydride 98% [CAS no. 2320436-6]. 10. Ethanolamine 98% [CAS no. 141-43-5]. 11. Piperidine 99% [CAS no. 110-89-4].

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12. Transferrin, Alexa Fluor™ 488 conjugate (Tfn-A488) OR 594 conjugate (Tfn-A594), 4 μg/mL. 13. Dulbecco’s Modified Eagle’s Medium (DMEM). 14. High grade 100% DMSO (non-oxidized, ideally small sealed ampoules). 15. Paraformaldehyde, 4% in PBS. 16. 40 ,6-Diamine-20 -phenylindole dihydrochloride (DAPI). 2.2

Solvents

1. Tetrahydrofuran (THF). 2. Dimethyl formamide (DMF). 3. Ethanol (EtOH). 4. Dichloromethane (DCM). 5. Concentrated hydrochloric acid (conc. HCl). 6. Toluene. 7. Methanol (MeOH). 8. Diethyl ether (ether, Et2O). 9. Hydrochloric acid: 1 M (1 M HCl). 10. Water. 11. Acidic wash solution for whole cell assay: 0.2 M acetic acid + 0.5 M NaCl, pH 2.8. 12. Phosphate-buffered saline (PBS).

2.3

Equipment

1. Weighing balance (e.g., Shimadzu AUW 220 D to 4 d.p.). 2. Magnetic stirrer with a temperature probe. 3. Rotary evaporator (e.g., Bu¨chi). 4. Vacuum system (e.g., Vacuubrand no. PC3001 VARIO). 5. Container for ice baths. 6. Teflon-coated magnetic stirrer bars. 7. Metal Drysyn heating blocks in various sizes (Asynt) or oil baths for heating. 8. Glassware: 25 and 50 mL round-bottom flasks, sintered glass funnel (porosity 3), Hirsch funnel, Bu¨chner flask, graduated cylinders, separatory funnel, and graduated pipettes. 9. Spatulas and tweezers. 10. Precoated silica gel 60F-254 plates (Merck, cat no. 1.05554.0001) and spotter for thin layer chromatography (TLC). 11. UV lamp (UVGL-58 handheld lamp, Australian Scientific) at 254 nm. 12. Heat gun.

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13. Biotage® Initiator+ microwave (optional). 14. Microwave vials for Biotage® microwave; 2–5 mL and 10–20 mL (optional). 15. Fibronectin-coated (5 μg/mL) 96-well glass plates.

3

Methods All procedures have been the subject to at least three independent trial by two researchers and the yields quoted are the average of three independent experiments. General spectroscopic details can be found in Note 1. The progress of reactions was monitored using a combination of LCMS and TLC. Subheading 3.1 refers to the Phthaladyn compounds (Subheading 3.1.1, Phthaladyn intermediate compound 6; Subheading 3.1.2, Phthaladyn active compound 2; and Subheading 3.1.3, Phthaladyn inactive compound 3). Subheading 3.2 refers to the Naphthalimide compounds (Subheading 3.2.1, Naphthalimide active compound 10 and Subheading 3.2.2, Naphthalimide inactive compound 11). Subheading 3.3 refers to the use of these probes in a typical whole cell assay.

3.1 Phthaladyn Compounds 3.1.1 Synthesis of 4-Chloro-2-(1,3-dioxo-1,3dihydroisobenzofuran-5carboxamido)benzoic Acid (Compound 6, Phthaladyn Intermediate)

1. Prepare an ice-water bath. 2. Weigh trimellitic anhydride chloride (600 mg, 0.35 mmol) and dissolve in 20 mL of THF in a 50 mL round-bottomed flask equipped with a Teflon-coated magnetic stirrer bar. 3. Weigh 4-chloro-2-aminobenzoic acid (482 mg, 0.35 mmol) and dissolve in 10 mL of THF. 4. Chill the trimellitic anhydride chloride solution for 20 min in the ice bath. 5. Add the benzoic acid solution dropwise. 6. Stir the reaction at 0  C for 3 h (see Note 2). 7. Collect the precipitate by filtration using a Hirsch or sintered funnel. 8. Wash the solid with 3  5 mL cold THF to give the intermediate (compound 6) as a white solid (see Note 3). NMR spectra in Fig. 9 (see Note 4).

3.1.2 Synthesis of 4-Chloro-2-(2-(4(hydroxymethyl)phenyl)1,3-dioxoisoindoline-5carboxamido)benzoic Acid (Compound 2, Phthaladyn Active)

1. Weigh intermediate (compound 6) (150 mg, 0.31 mmol). 2. Weigh 4-aminobenzyl alcohol (compound 7) (64 mg, 0.52 mmol). 3. Combine both reagents in 5 mL of toluene in a 25 mL roundbottomed flask fitted with a Teflon-coated magnetic stirrer bar. 4. Add 1 mL of DMF and 0.5 mL of TEA. 5. Heat the solution at 130  C for 18 h (see Note 5).

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Fig. 9 1H and 13C NMR spectra of 4-Chloro-2-(1,3-dioxo-1,3-dihydroisobenzofuran-5 carboxamido)benzoic acid (compound 6). Residual THF (solvent) present in spectra

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6. When the reaction is complete, cool to room temperature and add 1 M HCl (10 mL). 7. Add water (10 mL) and filter through a sintered glass funnel. 8. Collect the precipitate in a conical flask and add hot MeOH. The solid will not all dissolve. Cool to 0  C overnight (see Note 6). 9. Filter through a sintered funnel. Wash with cold EtOH (2  5 mL), then cold ether (2  5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. Allow the solid to dry under vacuum for at least 30 min (see Note 7). NMR spectra in Fig. 10. 3.1.3 Synthesis of 4Chloro-2-(1,3-dioxo-2phenylisoindoline-5carboxamido)benzoic Acid (3, Phthaladyn Inactive)

1. Weigh intermediate (compound 6) (150 mg, 0.31 mmol). 2. Dispense aniline (compound 8) (80 μL; 0.88 mmol). 3. Combine both reagents in 5 mL of toluene in a 25 mL roundbottomed flask fitted with a Teflon-coated magnetic stirrer bar. 4. Add 1 mL of DMF and 0.5 mL of TEA. 5. Heat the solution at 130  C for 18 h (see Note 8). 6. When the reaction is complete, cool to room temperature and add 10% HCl (10 mL). 7. Add water (10 mL) and filter through a sintered glass funnel. 8. Collect the precipitate in a conical flask and add hot MeOH. The solid will not all dissolve. 9. Cool to 0  C overnight (see Note 9). 10. Collect the product via filtration through a sintered glass funnel to give a brown solid (see Note 10). NMR spectra in Fig. 11.

3.2 Naphthalimide Compounds 3.2.1 Synthesis of 2-(4Aminobenzyl)-5-hydroxy1H-benzo[de]isoquinoline1,3(2H)-dione (Compound 10, Naphthalimide Active) Microwave Method

This reaction can be conducted via either microwave irradiation (Biotage® Initiator+) or batch methodology. The yield and purity are equivalent using either method.

1. To a 10–20 mL microwave vial fitted with a Teflon-coated stirrer bar, add 5-hydroxy-1,8-naphthalimide (compound 12) (200 mg, 0.93 mmol) and 4-aminobenzylamine (compound 13) (1.2 eq., 130 μL, 1.15 mmol). 2. Add ethanol (15 mL).

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Fig. 10 1H and 13C NMR spectra of 4-Chloro-2-(2-(4-(hydroxymethyl)phenyl)-1,3-dioxoisoindoline-5-carboxamido)benzoic acid (compound 2)

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Fig. 11 1H and (compound 3)

13

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C NMR spectra of 4-Chloro-2-(1,3-dioxo-2-phenylisoindoline-5-carboxamido)benzoic acid

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3. Seal the microwave vial and irradiate for 20 min at 120  C. 4. Once the vial has cooled to room temperature, cool at 0  C overnight (see Note 11). 5. Filter the reaction mixture through a sintered funnel. 6. Wash the collected solid with cold EtOH (2  5 mL), then cold ether (2  5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. 7. Allow to dry under vacuum for at least 30 min. The product is afforded as a crystalline tan solid (see Note 12). Batch Method

1. Weigh 5-hydroxy-1,8-naphthalimide (200 mg, 0.93 mmol).

(compound

12)

2. Dispense 4-aminobenzylamine (compound 13) (1.2 eq., 130 μL, 1.15 mmol). 3. Combine both reagents in 10 mL EtOH in a 35 mL roundbottomed flask fitted with a Teflon-coated magnetic stirrer bar. 4. Heat at reflux (100  C) for 18 h (see Note 13). 5. Cool the reaction to room temperature, and isolate the resulting yellow solid via filtration through a sintered glass funnel. 6. Wash the collected solid with cold EtOH (2  5 mL), then cold ether (2  5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. 7. Allow to dry under vacuum for at least 30 min. The product is afforded as a tan solid (see Note 14). NMR spectra in Fig. 12. 3.2.2 Synthesis of 2-(2Hydroxyethyl)-1H-benzo [de]isoquinoline-1,3(2H)dione (Compound 11, Naphthalimide Inactive) Microwave Method

This reaction can be conducted via either microwave irradiation (Biotage® Initiator+) or batch methodology. The yield and purity are equivalent using either method.

1. To a 10–20 mL microwave vial fitted with a Teflon-coated stirrer bar, add 1,8-naphthalimide (compound 14) (198 mg, 1 mmol) and ethanolamine (compound 15) (84 mL, 1.4 mmol). 2. Add ethanol (15 mL). 3. Seal the microwave vial and irradiate for 20 min at 120  C. 4. Once the vial has cooled to room temperature, cool at 0  C overnight (see Note 15). 5. Filter the reaction mixture through a sintered funnel.

Synthesis of Pthaladyn-29 and Napthalimide-10

Fig. 12 1H and (compound 10)

13

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C NMR spectra of 2-(4-Aminobenzyl)-5-hydroxy-1H-benzo[de]isoquinoline-1,3(2H)-dione

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6. Wash the collected solid with cold EtOH (2  5 mL), then cold ether (2  5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. 7. Allow to dry under vacuum for at least 30 min. The product is afforded as a crystalline tan solid (see Note 16). Batch Method

1. To a 50 mL round-bottomed flask fitted with a Teflon-coated stirrer bar, add 1,8-naphthalimide (compound 14) (198 mg, 1 mmol) and ethanolamine (compound 15) (84 mL, 1.4 mmol). 2. Add ethanol (20 mL) and piperidine (4 drops, catalytic). 3. Fit the round-bottomed flask with a reflux condenser and heat to reflux (100  C) overnight (see Note 17). 4. Once the reaction mixture has cooled to room temperature, reduce the solvent by half and cool at 0  C overnight (see Note 18). 5. Filter the reaction mixture through a sintered funnel. 6. Wash the collected solid with cold EtOH (2  5 mL), then cold ether (2  5 mL). Ensure the vacuum hose is removed from the flask and the solid is stirred with each solvent prior to filtering to enable adequate washing. 7. Allow to dry under vacuum for at least 30 min. The product is afforded as a tan solid (see Note 19). NMR spectra in Fig. 13.

3.3 Biological Methods for Whole Cell Assay

1. Prepare stock solution of drug as a 30 mM solution in 100%, high grade DMSO (see Note 20). 2. U2OS cells are grown in fibronectin-coated (5 μg/mL) 96-well glass plates then serum-starved overnight (16 h) in serum-free DMEM (see Note 21). 3. Pre-incubate cells with different concentrations of (a) dynamin inhibitor, (b) inactive analogs, or (c) vehicle (DMSO) for 15–30 min prior to addition of 4 μg/mL Tfn-A488 or Tfn-A594 for a further 8 min at 37  C in the continued presence of the compound (see Note 22). 4. Cell surface-bound fluorescent Tfn is removed by incubating the cells with an ice-cold acidic wash solution (0.2 M acetic acid + 0.5 M NaCl, pH 2.8) for 10 min followed by its removal with an ice-cold PBS wash for 5 min (see Note 23). 5. Fix cells with 4% paraformaldehyde for 10 min at 37  C, and stain cell nuclei using DAPI.

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Fig. 13 1H and [13] NMR spectra of 2-(2-Hydroxyethyl)-1H-benzo[de]isoquinoline-1,3(2H)-dione (compound 11)

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Notes 1. 1H NMR were recorded at either 400 or 600 MHz and 13C NMR were recorded at either 101 or 150 MHz, respectively, using a Bruker Avance 400 MHz or 600 MHz spectrometer in CDCl3 and DMSO-d6. LCMS was performed using an Agilent 6100 series single quadrupole LCMS using a mobile phase of 1:1 acetonitrile:H2O with 0.1% formic acid. The UPLC was fitted with a Zorbax SB-C18 Rapid Resolution HT 2.1  50 mm 1.8-Micron column. 2. Follow the reaction progress by TLC or LCMS. 3. Average yield, 61%. Over-washing the product with THF may result in a lower yield. This material is best stored at 0  C and used within a few days. 4. Pulse delay is required for NMR for this compound. 5. Follow the reaction progress by TLC or LCMS. 6. The residue can be stored at 0  C for several days at this point without loss in product purity or yield. 7. Average yield, 29%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 8. Follow the progress of the reaction by TLC (1:9 MeOH: DCM). 9. The residue can be stored at 0  C for several days at this point without loss in product purity or yield. 10. Average yield, 46%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 11. The reaction mixture can be left at 0  C for several weeks without reduction in product purity. 12. Average yield, 86%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 13. Follow the progress of the reaction by LCMS. The reaction mixture can be stirred at reflux over the weekend without loss in product purity or yield. 14. Average yield, 69%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 15. The reaction mixture can be left at 0  C for several weeks without reduction in product purity.

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16. Average yield, 86%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 17. The reaction mixture can be left to stir at reflux for several days without reduction in product purity or yield. 18. The reaction mixture can be left at 0  C for several weeks without reduction in product purity. 19. Average yield, 62%. This compound can be stored below 18  C in a container in the dark for more than 12 months without notable loss of purity. 20. Sub-aliquots of the stock solution can be generated and frozen at 20  C for at least 6 months after at least two freeze-thaw cycles. Subsequent experiments can be performed using working solutions that are freshly prepared, immediately before use, by dilution of primary stocks into pre-warmed (37  C) cell culture medium such as DMEM (or 20 mM Tris–HCl pH 7.4 for GTPase assay) to the desired final concentrations. Most cells can tolerate up to 1% DMSO, while dynamin GTPase assays in vitro can tolerate up to 3%. It is preferable to avoid serum or albumin in all drug solutions as most of these compounds bind albumin, reducing the compound access to cells. 21. This stage removes endogenous hormones and growth factors and reduces mitosis. A minimum of two hours of serumstarvation is sufficient for sensitive CME assays, whereas overnight may be more operationally convenient. 22. We typically recommend use of 20–30 μM of each inhibitor, but a concentration curve for each active and inactive analog is the optimal experimental design, ranging from 1 μM to 60 μM. With many small molecules, there can be an issue of the compounds lifting the cells from the plates, confounding analysis by imaging microscopy. Lifting differs for each cell type we have investigated and can become pronounced at higher concentrations and should be monitored for each cell type. 23. This low temperature prevents further endocytosis and halts all known vesicle trafficking during this step. References 1. Ferguson SM, De Camilli P (2012) Dynamin, a membrane-remodelling GTPase. Nat Rev Mol Cell Biol 13:75–88. https://doi.org/10. 1038/nrm3266 2. Boumil RM, Letts VA, Roberts MC et al (2010) A missense mutation in a highly conserved alternate exon of dynamin-1 causes epilepsy in fitful mice. PLoS Genet 6:e1001046.

https://doi.org/10.1371/journal.pgen. 1001046 3. Pathan SA, Jain GK, Akhter S et al (2010) Insights into the novel three ‘D’s of epilepsy treatment: drugs, delivery systems and devices. Drug Discov Today 15:717–732. https://doi. org/10.1016/j.drudis.2010.06.014

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4. Chaumont S, Andre´ C, Perrais D et al (2013) Agonist-dependent endocytosis of γ-aminobutyric acid type A (GABAA) receptors revealed by a γ2(R43Q) epilepsy mutation. J Biol Chem 288:28254–28265. https://doi. org/10.1074/jbc.M113.470807 5. Eleniste PP, Huang S, Wayakanon K et al (2014) Osteoblast differentiation and migration are regulated by Dynamin GTPase activity. Int J Biochem Cell Biol 46:9–18. https://doi. org/10.1016/j.biocel.2013.10.008 6. Bruzzaniti A, Neff L, Sanjay A et al (2005) Dynamin forms a Src kinase-sensitive complex with Cbl and regulates podosomes and osteoclast activity. Mol Biol Cell 16:3301–3313. https://doi.org/10.1091/mbc.E04-12-1117 7. Wolf MTF, Wu XR, Huang CL (2013) Uromodulin upregulates TRPV5 by impairing Caveolin-mediated endocytosis. Kidney Int 84:130–137. https://doi.org/10.1038/ki. 2013.63 8. Soda K, Balkin DM, Ferguson SM et al (2012) Role of dynamin, synaptojanin, and endophilin in podocyte foot processes. J Clin Invest 122: 4401–4411. https://doi.org/10.1172/ JCI65289 9. Harper CB, Popoff MR, McCluskey A et al (2013) Targeting membrane trafficking in infection prophylaxis: dynamin inhibitors. Trends Cell Biol 23:90–101. https://doi.org/ 10.1016/j.tcb.2012.10.007 10. Sun Y, Tien P (2013) From endocytosis to membrane fusion: emerging roles of dynamin in virus entry. Crit Rev Microbiol 39:166–179. https://doi.org/10.3109/1040841x.2012. 694412 11. Chappie JS, Mears JA, Fang S et al (2011) A pseudo-atomic model of the dynamin polymer identifies a hydrolysis-dependent powerstroke. Cell 147:209–222. https://doi.org/10.1016/ j.cell.2011.09.003 12. Marks B, Stowell MHB, Vallis Y et al (2001) GTPase activity of dynamin and resulting conformation change are essential for endocytosis. Nature 410:231–235. https://doi.org/10. 1038/35065645 13. Praefcke GJK, McMahon HT (2004) The dynamin superfamily: universal membrane tubulation and fission molecules? Nat Rev Mol Cell Biol 5:133–147. https://doi.org/ 10.1038/nrm1313 14. Sever S, Muhlberg AB, Schmid SL (1999) Impairment of dynamin’s GAP domain stimulates receptor-mediated endocytosis. Nature 398:481–486. https://doi.org/10.1038/ 19024

15. Schmid SL, Frolov VA (2011) Dynamin: functional design of a membrane fission catalyst. Annu Rev Cell Dev Biol 27:79–105. https:// doi.org/10.1146/annurev-cellbio100109-104016 16. Faelber K, Held M, Gao S et al (2012) Structural insights into dynamin-mediated membrane fission. Structure 20:1621–1628. https://doi.org/10.1016/j.str.2012.08.028 17. Ferguson SM, Brasnjo G, Hayashi M et al (2007) A selective activity-dependent requirement for dynamin 1 in synaptic vesicle endocytosis. Science 316:570–574. https://doi.org/ 10.1126/science.1140621 18. Liu YW, Surka MC, Schroeter T et al (2008) Isoform and splice-variant specific functions of dynamin-2 revealed by analysis of conditional knock-out cells. Mol Biol Cell 19:5347–5359. https://doi.org/10.1091/mbc.E08-08-0890 19. Raimondi A, Ferguson SM, Lou X et al (2011) Overlapping role of dynamin isoforms in synaptic vesicle endocytosis. Neuron 70: 1100–1114. https://doi.org/10.1016/j.neu ron.2011.04.031 20. Hill TA, Gordon CP, McGeachie AB et al (2009) Inhibition of dynamin mediated endocytosis by the dynoles™—synthesis and functional activity of a family of indoles. J Med Chem 52:3762–3773. https://doi.org/10. 1021/jm900036m 21. Luwor R, Morokoff AP, Amiridis S et al (2019) Targeting glioma stem cells by functional inhibition of dynamin 2: a novel treatment strategy for glioblastoma. Cancer Investig 37:144–155. https://doi.org/10.1080/07357907.2019. 1582060 22. Bain J, Plater L, Elliot M et al (2007) The selectivity of protein kinase inhibitors: a further update. Biochem J 408:297–315. https://doi. org/10.1042/BJ20070797 23. Park RJ, Shen H, Liu L et al (2013) Dynamin triple knockout cells reveal off target effects of commonly used dynamin inhibitors. J Cell Sci 126:5305–5312. https://doi.org/10.1242/ jcs.138578 24. Odell LR, Howan D, Gordon CP et al (2010) The pthaladyns: competitive inhibitors of dynamin I and Dynamin II GTPase derived from virtual screening. J Med Chem 53: 5267–5280. https://doi.org/10.1021/ jm100442u 25. Abdel-Hamid MK, Macgregor KA, Odell LR et al (2015) 1,8-Naphthalimide derivatives: new leads against dynamin I GTPase activity. Org Biomol Chem 13:8016–8028. https:// doi.org/10.1039/c5ob00751h

INDEX A Acetylation solution .......................................................... 3 Ammonium chloride buffer................................. 186, 189 AMPA receptors ............................................................ 107 Artificial cerebrospinal fluid (ACSF).......... 114, 115, 208

B Basket cell (BC).................................................... 115–119 Blocking buffer........................... 3, 10, 11, 16, 91, 94–96

C Calcium imaging ..........................................194, 196–199 Calcium-phosphate transfection................................... 188 cDNA.............................8, 34, 46, 60, 61, 63, 64, 66, 67 Cell culture .................................... 21, 31–36, 39, 46, 53, 56, 75, 92, 122, 187, 236, 257 Cell detection ............................................................11, 12 Chemical scaffolds................................................ 222, 240 Chemical synthesis ........................................................ 222 Concanamycin A .................................182, 186, 188, 189 Crosscorrelation ............................................................ 149 Cryo-grinding ...................................................... 121–129 Culture medium................................................22, 23, 25, 31–35, 47, 48, 50, 184, 206, 214, 236, 257 Cutting solution................................................... 114, 115 Cytometry by Time-Of-Flight mass spectrometry (CyTOF) ......................................... 70, 72, 76, 78, 80, 81, 85, 86

D Dense core vesicle (DCV) ..................193–195, 201, 202 Digestion solution......................................................... 185 Drosophila melanogaster.................................................. 20 DsRed ............................................................................ 213 Dynamin-inactive compounds ........................... 222, 223, 226, 229, 240, 254 Dynamins............................................. 221–237, 239–257 Dynoles .............................. 222, 223, 225, 229–231, 240

E Electrically-triggered release................................ 194, 202 Electrotransfer ..............................................89, 91, 93–95 Endocytosis .............. 107, 114, 132, 181, 237, 239, 257

Endosomes ................................................................29–43 Exocytosis ........................................ 1, 56, 102, 104, 105, 114, 132, 182, 186, 188, 202, 222

F Fluorescence cross correlation spectroscopy (FCCS)................. 147, 167–179 in situ hybridization (FISH) ................................2, 12 western Blotting .................................................. 89–98 Fly strains.............................................................. 196, 197 Fo¨rster resonance energy transfer (FRET) .................151, 168, 178 Functional crosstalk .................................... 102, 103, 108

G GABA.................................................29, 46, 53, 113–119 GAL4 ............................................................................. 194 Genetically encoded .................................... 182, 193, 206 Genetically-encoded glutamate indicators (GEGIs) ................. 206, 207, 210, 213, 214, 217 Glutamate ...................................... 29, 30, 103–105, 107, 114, 127, 140, 195–197, 201, 202, 205–219 GLYT type 1 (GLYT1)................................................. 105 Gradient ultracentrifugation...............123, 125, 128, 129

H HEK293T.................................31, 34, 39, 41, 47, 48, 56 Henderson-Hasselbalch .................................................. 54 Hippocampus .................................... 1, 4, 14, 61, 62, 67, 83, 187, 206, 210–212 Hippocampus culture .......................................... 183–188

I IGluSnFR .......................... 206, 211–213, 215, 217, 218 ImageJ.......................................................... 197, 199, 201 Imaging buffer ....................................183, 186, 188, 189 In situ hybridization ................................................... 1–17 Intracellular photolysis.................................................. 113

L Lentiviral transfer plasmid ........................................47, 48 Live-cell fluorescence imaging .............................. 53, 186

Jana Dahlmanns and Marc Dahlmanns (eds.), Synaptic Vesicles: Methods and Protocols, Methods in Molecular Biology, vol. 2417, https://doi.org/10.1007/978-1-0716-1916-2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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260 Index M

Master mix.......................................................... 3, 6, 8, 85 MATLAB ........................... 166, 177, 182, 187, 197, 210 Membrane fusion ................................................. 167–179 Membrane transporters ..................................... 30, 32, 33 Methanesulfonate................................................. 115, 116 MicroRNAs (miRNA) ............................ 1–10, 12, 14, 16 miRNA RT-qPCR......................................................... 2, 8 Mixer Mill........................................................................ 61 Molecular probes .......................................................... 222 Morange2 ....................................... 46, 47, 51, 52, 55, 57 Motor neurons ..................................................... 194–202 Multiplexing ..............................................................70, 96 Mushroom body (MB) ................................................... 21

N Nanodrop ............................................................. 5, 14, 63 Napthalimide-10 .................................................. 239–257 Neuromuscular junctions .................................... 193–202 Neuropeptides ...................................................... 193–202 Nitrocellulose membranes ........................ 91, 94, 96, 135 NMDA receptors ................................................. 103–108

O Organelles................. 19, 30, 45, 46, 100, 122, 132, 133 Organotypic slice cultures .................. 205–213, 215–219

P Patch-clamp ..........................................30, 38, 39, 42, 43, 115, 116, 210, 213 Patch pipette perfusion ........................................ 113–119 pH ................................................................ 221, 222, 240 Phosphoprotein detection .............................................. 96 Phosphorylation .......................................... 105, 108, 205 Photolysis.............................................................. 113–119 pH-sensitive.................................... 46, 51, 142, 144, 182 Pipette perfusion ......................................... 115, 116, 118 Pipette solution ..............................................30, 115, 117 pka ............................................................... 46, 51, 52, 54 Plasmids ................................................31, 32, 34, 35, 47, 48, 186, 212, 213, 216, 217 PolyJet ................................................................ 31, 34, 35 Polymerase.................................................................59, 64 Presynaptic receptors ...................................102, 107–108 Presynaptic terminals .................105, 113, 122, 132, 215 Primary hippocampus cultures ....................183, 186–188 Proteoliposomes........................... 30, 169–173, 175, 177 Pthaladyn-29 ................................................................. 241

Pupae .................................................................. 22, 25, 26 Pyramidal cells ...................................................... 206, 211

R Rat.....................................................59–67, 96, 114, 139, 187, 190, 194, 206, 210, 211 Readily releasable pool (RRP) ............................ 181–184, 189–191, 200 Real-time qPCR ........................................................59–67 Receptor trafficking ...................................................... 108 Recycling pool (RP)............................ 181–184, 189–191 Release reporters .................................................. 193–202 Reverse transcription............................................ 2, 3, 6, 7 RNA ...................................................... 2, 4–6, 10, 13, 14, 27, 60–65, 96, 122 RNase.................................................................... 2, 60, 62

S ScanImage ............................................................ 210, 218 SDS polyacrylamide gel electrophoresis ..................90, 92 Single-cell electroporation ...........................207–208, 210 Size exclusion liquid chromatography ............... 121–129, 132, 133, 135, 139, 141, 169 Slice cultures........................................206, 208, 212–214 Slice preparations ................................................. 113–119 SNAREs ....................................................... 167, 168, 179 Stereomicroscope ................................................... 24, 213 SYBR Green ................................. 6, 8, 15, 59, 60, 64, 67 Synaptic transmission ................................. 108, 114, 121, 181, 208–210, 213 Synapto-pHluorin (spH) ..................................... 181–191

T Taqman ...........................................................2, 3, 6, 8, 59 TaqMan miRNA reverse transcription kit ....................... 2 Thermal Cycler................................................................ 61 Trizol .................................................................. 2, 4, 5, 13

V Vesicle pools ............................... 182, 183, 187, 189–191

W Western blotting..................................62, 67, 89–98, 105 Wistar/ST rats ...................................................... 114, 115

Z Z-stack .................................................................. 199, 201