Fluorescence In Situ Hybridization (FISH): Methods and Protocols (Methods in Molecular Biology, 2784) 1071637657, 9781071637654

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Methods in Molecular Biology 2784

Gal Haimovich  Editor

Fluorescence In Situ Hybridization (FISH) 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-by step 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.

Fluorescence In Situ Hybridization (FISH) Methods and Protocols

Edited by

Gal Haimovich Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel

Editor Gal Haimovich Department of Molecular Genetics Weizmann Institute of Science Rehovot, Israel

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3765-4 ISBN 978-1-0716-3766-1 (eBook) https://doi.org/10.1007/978-1-0716-3766-1 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 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. Cover Caption: The cover image shows a single-molecule FISH of a cell-line. Image courtesy of Gal Haimovich. 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. Paper in this product is recyclable.

Preface In situ hybridization methods allow the detection, quantification, and localization of specific nucleic acid sequences (DNA and RNA) by microscopic analysis. The method stems from the fact that two strands in a DNA double helix or the RNA secondary structures can be separated by denaturation and then reannealed (hybridized) with labeled complementary strands under conditions which favor specific duplex formation. A labeled DNA sequence (probe) is used to localize the gene sequences of interest (the target sequence). The technique was originally developed by three groups independently: Gall and Pardue (PNAS), John et al. (Nature), and Buongiorno-Nardelli and Amaldi (Nature) in 1969– 1970 using radioactive-labeled probes for the in situ detection of DNA sequences but later was switched to the safer and more sensitive fluorescent probes. Since its invention, multiple Fluorescent In Situ Hybridization (FISH) protocols were developed, combining state-ofthe-art probe design, microscopy methods, and computational tools to study unique aspects of nucleic acids biology and cell biology. These methods allow detection of single mRNA molecules or distinct DNA loci at remarkable precision. Multiplexing FISH allows the detection of several and up to thousands of distinct sequences in a single experiment. Furthermore, FISH provides quantitative measurements at the single cell, single organelle, and single molecule level with high accuracy. This book aims to offer up-do-date FISH approaches and protocols, which would benefit the broader scientific community. The protocols provide detailed steps from probe design to imaging and image analysis, including tips that are typically absent in research papers but can sometimes make or break an experiment. This volume is divided into two parts. The first part is dedicated to RNA FISH protocols and the second part, with fewer protocols, to DNA FISH protocols. MiMB have previously published volumes on DNA FISH (Ed. Bridger and Volpi, 2010) and Imaging Gene Expression (Ed. Shav-Tal, 2019), so for this volume I tried to collect protocols that focus on unique biological questions, model organisms not typically studied by FISH, protocols combining FISH with immunofluorescence (FISH-IF) and protocols for high-throughput experiments. In the first part, RNA FISH and FISH-IF protocols provide methods for designing OligoPaint probes and for studying distinct aspects of RNA biology such as transcription and splicing dynamics, mRNA and small RNA expression and localization, RNA folding, and RNA modifications. These protocols cover a wide variety of organisms, including bacteria, plants, yeast, hydra, sea anemone, and mammalian cells and tissues. In the second part, we transition with a combined DNA and RNA FISH-IF protocol, followed by DNA FISH protocols that are used to study DNA repair dynamics, gene compaction, chromatin conformation, and gene rearrangements in plant, insect, and mammalian cells. For the last protocol, I included a different approach to study unique DNA sequences, not by FISH, but rather by CRISPR, for targeting of a luminescent sensor to specific DNA sequences. I am grateful to the many scientists who contributed to this book. Thank you for taking the time to write these detailed protocols for the benefit of your colleagues. Rehovot, Israel

Gal Haimovich

v

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

PART I

v ix

RNA FISH

1 Single-Molecule Fluorescent In Situ Hybridization (smFISH) for RNA Detection in Bacteria. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Camilla Ciolli Mattioli and Roi Avraham 2 Single-Molecule Fluorescent In Situ Hybridization (smFISH) for RNA Detection in the Fungal Pathogen Candida albicans . . . . . . . . . . . . . . . . Sander van Otterdijk, Maryam Motealleh, Zixu Wang, Thomas D. Visser, Philipp Savakis, and Evelina Tutucci 3 RNA and Protein Detection by Single-Molecule Fluorescent in Situ Hybridization (smFISH) Combined with Immunofluorescence in the Budding Yeast S. cerevisiae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anna Maekiniemi and Robert H. Singer 4 Fluorescence In Situ Hybridization as a Tool for Studying the Specification and Differentiation of Cell Types in Nematostella vectensis . . . . . . . . . . . . . . . . . . . Oce´ane Tournie`re, Henriette Busengdal, James M. Gahan, and Fabian Rentzsch 5 SABER-FISH in Hydractinia. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Miguel Salinas-Saavedra 6 smFISH for Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sahar Hani, Caroline Mercier, Pascale David, Thierry Desnos, Jean-Marc Escudier, Edouard Bertrand, and Laurent Nussaume 7 Fluorescent In Situ Detection of Small RNAs in Plants Using sRNA-FISH . . . . Kun Huang, Blake C. Meyers, and Jeffrey L. Caplan 8 hcHCR: High-Throughput Single-Cell Imaging of RNA in Human Primary Immune Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Manasi Gadkari, Jing Sun, Adrian Carcamo, Iain Fraser, Luis M. Franco, and Gianluca Pegoraro 9 High-Throughput RNA-HCR-FISH Detection of Endogenous Pre-mRNA Splice Variants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Asaf Shilo, Gianluca Pegoraro, and Tom Misteli 10 Simultaneous In Situ Detection of m6A-Modified and Unmodified RNAs Using DART-FISH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Charles J. Sheehan and Kate D. Meyer 11 Multiplexed Immunofluorescence and Single-Molecule RNA Fluorescence In Situ Hybridization in Mouse Skeletal Myofibers . . . . . . . . . . . . . Lance T. Denes, Chase P. Kelley, and Eric T. Wang

vii

3

25

45

59

77 87

101

113

133

147

163

viii

Contents

12

Designing Oligonucleotide-Based FISH Probe Sets with PaintSHOP . . . . . . . . . 177 Monika W. Perez, Conor K. Camplisson, and Brian J. Beliveau 13 Efficient Prediction Model of mRNA End-to-End Distance and Conformation: Three-Dimensional RNA Illustration Program (TRIP) . . . . 191 Jiayun Ma and Tatsuhisa Tsuboi

PART II

DNA FISH

14

Combined 3D DNA FISH, Single-Molecule RNA FISH, and Immunofluorescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Souvik Sen, Shivnarayan Dhuppar, and Aprotim Mazumder 15 Determining the Compaction State of Genes Using DNA FISH. . . . . . . . . . . . . . Masako Narita, Ioana Olan, and Masashi Narita 16 Hi-M: A Multiplex Oligopaint FISH Method to Capture Chromatin Conformations In Situ and Accompanying Open-Source Acquisition Software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jean-Bernard Fiche, Marie Schaeffer, Christophe Houbron, Christel Elkhoury Youhanna, Olivier Messina, Franziska Barho, and Marcelo Nollmann 17 Rapid DNA-FISH in Arabidopsis thaliana Somatic Cells . . . . . . . . . . . . . . . . . . . . Olga Raskina and Ofir Hakim 18 DBD-FISH Using Specific Chromosomal Region Probes for the Study of Cervical Carcinoma Progression . . . . . . . . . . . . . . . . . . . . . . . . . . . Catalina Garcı´a-Vielma, Elva I. Corte´s-Gutie´rrez, Jose´ L. Ferna´ndez, Martha I. Da´vila-Rodrı´guez, and Jaime Gosa´lvez 19 CRISPR-Based Split Luciferase as a Biosensor for Unique DNA Sequences In Situ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nicholas G. Heath and David J. Segal Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

203 215

227

259

271

285 301

Contributors ROI AVRAHAM • Department of Immunology and Regenerative Biology, Weizmann Institute of Science, Rehovot, Israel FRANZISKA BARHO • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France BRIAN J. BELIVEAU • Department of Genome Sciences, University of Washington, Seattle, WA, USA; Brotman Baty Institute for Precision Medicine, Seattle, WA, USA; Institute for Stem Cell and Regenerative Medicine, University of Washington, Seattle, WA, USA EDOUARD BERTRAND • Institut de Ge´ne´tique Humaine, CNRS, UMR9002, Montpellier, France HENRIETTE BUSENGDAL • Michael Sars Centre, University of Bergen, Bergen, Norway CONOR K. CAMPLISSON • Department of Genome Sciences, University of Washington, Seattle, WA, USA JEFFREY L. CAPLAN • Department of Plant and Soil Sciences, University of Delaware, Newark, DE, USA; Bio-Imaging Center, Delaware Biotechnology Institute, University of Delaware, Newark, DE, USA ADRIAN CARCAMO • High-Throughput Imaging Facility (HiTIF), Center for Cancer Research (CCR), NCI/NIH, Bethesda, MD, USA CAMILLA CIOLLI MATTIOLI • Department of Immunology and Regenerative Biology, Weizmann Institute of Science, Rehovot, Israel ELVA I. CORTE´S-GUTIE´RREZ • Faculty of Biological Sciences, Universidad Autonoma de Nuevo Leon, Monterrey, Mexico PASCALE DAVID • Aix Marseille Univ, CEA, CNRS, BIAM, UMR7265, Saint-Paul lez Durance, France MARTHA I. DA´VILA-RODRI´GUEZ • Faculty of Public Health and Nutrition, Universidad Autonoma de Nuevo Leon, Monterrey, Mexico LANCE T. DENES • Institute for Systems Genetics, NYU Grossman School of Medicine, New York, NY, USA THIERRY DESNOS • Aix Marseille Univ, CEA, CNRS, BIAM, UMR7265, Saint-Paul lez Durance, France SHIVNARAYAN DHUPPAR • Tata Institute of Fundamental Research Hyderabad, Hyderabad, Telangana, India; Ann Romney Center for Neurologic Diseases, Department of Neurology, Brigham and Women’s Hospital and Harvard Medical School, Boston, MA, USA CHRISTEL ELKHOURY YOUHANNA • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France JEAN-MARC ESCUDIER • Laboratoire Synthe`se et Physico-Chimie de Mole´cules d’inte´reˆt Biologique, Universite´ Paul Sabatier, CNRS, Toulouse, France JOSE´ L. FERNA´NDEZ • Genetics Unit, INIBIC, Complejo Hospitalario Universitario A Corun˜a, La Corun˜a, Spain; Laboratorio de Gene´tica Molecular y Radiobiologı´a Centro Oncologico de Galicia, La Corun ˜ a, Spain JEAN-BERNARD FICHE • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France LUIS M. FRANCO • Functional Immunogenomics Section, NIAMS/NIH, Bethesda, MD, USA

ix

x

Contributors

IAIN FRASER • Signaling Systems Section, Laboratory of Immune System Biology, NIAID/NIH, Bethesda, MD, USA MANASI GADKARI • Functional Immunogenomics Section, NIAMS/NIH, Bethesda, MD, USA JAMES M. GAHAN • Michael Sars Centre, University of Bergen, Bergen, Norway; Department of Biochemistry, University of Oxford, Oxford, UK CATALINA GARCI´A-VIELMA • Department of Genetics, Centro de Investigacion Biome´dica del Noreste, Instituto Mexicano del Seguro Social (IMSS), Monterrey, Nuevo Leon, Mexico JAIME GOSA´LVEZ • Unit of Genetics, Department of Biology, Universidad Autonoma de Madrid, Madrid, Spain OFIR HAKIM • The Mina and Everard Goodman Faculty of Life Sciences, Bar-Ilan University, Ramat-Gan, Israel SAHAR HANI • Aix Marseille Univ, CEA, CNRS, BIAM, UMR7265, Saint-Paul lez Durance, France; Center for Integrative Genomics, University of Lausanne, Lausanne, Switzerland NICHOLAS G. HEATH • Genome Center and Department of Biochemistry and Molecular Medicine, University of California, Davis, Davis, CA, USA; Integrative Genetics and Genomics, University of California, Davis, Davis, CA, USA; Innovative Genomics Institute, University of California, Berkeley, CA, USA CHRISTOPHE HOUBRON • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France KUN HUANG • Department of Plant and Soil Sciences, University of Delaware, Newark, DE, USA; Bio-Imaging Center, Delaware Biotechnology Institute, University of Delaware, Newark, DE, USA; Molecular Imaging Core, Dana-Farber Cancer Institute, Boston, USA CHASE P. KELLEY • Department of Molecular Genetics and Microbiology, University of Florida, Gainesville, FL, USA JIAYUN MA • Institute of Biopharmaceutical and Health Engineering, Tsinghua Shenzhen International Graduate School, University Town of Shenzhen, Shenzhen, Guangdong, China ANNA MAEKINIEMI • Department of Cell Biology, Albert Einstein College of Medicine, Bronx, NY, USA APROTIM MAZUMDER • Tata Institute of Fundamental Research Hyderabad, Hyderabad, Telangana, India CAROLINE MERCIER • Aix Marseille Univ, CEA, CNRS, BIAM, UMR7265, Saint-Paul lez Durance, France; Biochimie et Physiologie Mole´culaire des Plantes, Univesite´ de Montpellier, CNRS, INRAE, Institut Agro, Montpellier, France OLIVIER MESSINA • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France KATE D. MEYER • Department of Biochemistry, Duke University School of Medicine, Durham, NC, USA; Department of Neurobiology, Duke University School of Medicine, Durham, NC, USA BLAKE C. MEYERS • Department of Plant and Soil Sciences, University of Delaware, Newark, DE, USA; Donald Danforth Plant Science Center, St. Louis, MO, USA; Division of Plant Science and Technology, University of Missouri – Columbia, Columbia, MO, USA TOM MISTELI • Cell Biology of Genomes, Center for Cancer Research (CCR), National Cancer Institute, NIH, Bethesda, MD, USA

Contributors

xi

MARYAM MOTEALLEH • Systems Biology Lab, A-LIFE department, Amsterdam Institute of Molecular and Life Sciences (AIMMS), Vrije Universiteit Amsterdam, Amsterdam, The Netherlands MASAKO NARITA • Cancer Research UK Cambridge Institute, University of Cambridge, Cambridge, UK MASASHI NARITA • Cancer Research UK Cambridge Institute, University of Cambridge, Cambridge, UK MARCELO NOLLMANN • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France LAURENT NUSSAUME • Aix Marseille Univ, CEA, CNRS, BIAM, UMR7265, Saint-Paul lez Durance, France IOANA OLAN • Cancer Research UK Cambridge Institute, University of Cambridge, Cambridge, UK GIANLUCA PEGORARO • High-Throughput Imaging Facility (HiTIF), Center for Cancer Research (CCR), National Cancer Institute, NIH, Bethesda, MD, USA MONIKA W. PEREZ • Department of Genome Sciences, University of Washington, Seattle, WA, USA OLGA RASKINA • Institute of Evolution, University of Haifa, Haifa, Israel FABIAN RENTZSCH • Department of Biological Sciences, University of Bergen, Bergen, Norway MIGUEL SALINAS-SAAVEDRA • University of Galway, Galway, Ireland PHILIPP SAVAKIS • Systems Biology Lab, A-LIFE department, Amsterdam Institute of Molecular and Life Sciences (AIMMS), Vrije Universiteit Amsterdam, Amsterdam, The Netherlands MARIE SCHAEFFER • Centre de Biologie Structurale, Univ Montpellier, CNRS UMR5048, INSERM U1054, Montpellier, France DAVID J. SEGAL • Genome Center and Department of Biochemistry and Molecular Medicine, University of California, Davis, Davis, CA, USA; Integrative Genetics and Genomics, University of California, Davis, Davis, CA, USA; Innovative Genomics Institute, University of California, Berkeley, CA, USA SOUVIK SEN • Tata Institute of Fundamental Research Hyderabad, Hyderabad, Telangana, India CHARLES J. SHEEHAN • Department of Biochemistry, Duke University School of Medicine, Durham, NC, USA ASAF SHILO • Cell Biology of Genomes, Center for Cancer Research (CCR), National Cancer Institute, NIH, Bethesda, MD, USA ROBERT H. SINGER • Department of Cell Biology, Albert Einstein College of Medicine, Bronx, NY, USA JING SUN • Signaling Systems Section, Laboratory of Immune System Biology, NIAID/NIH, Bethesda, MD, USA OCE´ANE TOURNIE`RE • Michael Sars Centre, University of Bergen, Bergen, Norway; Institut de Biologie Valrose, Universite´ Coˆte d’Azur, CNRS, INSERM, Nice, France TATSUHISA TSUBOI • Institute of Biopharmaceutical and Health Engineering, Tsinghua Shenzhen International Graduate School, University Town of Shenzhen, Shenzhen, Guangdong, China EVELINA TUTUCCI • Systems Biology Lab, A-LIFE department, Amsterdam Institute of Molecular and Life Sciences (AIMMS), Vrije Universiteit Amsterdam, Amsterdam, The Netherlands

xii

Contributors

SANDER VAN OTTERDIJK • Systems Biology Lab, A-LIFE department, Amsterdam Institute of Molecular and Life Sciences (AIMMS), Vrije Universiteit Amsterdam, Amsterdam, The Netherlands THOMAS D. VISSER • Systems Biology Lab, A-LIFE department, Amsterdam Institute of Molecular and Life Sciences (AIMMS), Vrije Universiteit Amsterdam, Amsterdam, The Netherlands; TNW-BT-IMB, Delft University of Technology, Delft, The Netherlands ERIC T. WANG • Department of Molecular Genetics and Microbiology, University of Florida, Gainesville, FL, USA ZIXU WANG • Systems Biology Lab, A-LIFE department, Amsterdam Institute of Molecular and Life Sciences (AIMMS), Vrije Universiteit Amsterdam, Amsterdam, The Netherlands

Part I RNA FISH

Chapter 1 Single-Molecule Fluorescent In Situ Hybridization (smFISH) for RNA Detection in Bacteria Camilla Ciolli Mattioli and Roi Avraham Abstract In this chapter, we describe in detail how to perform a successful smFISH experiment and how to quantify mRNA transcripts in bacterial cells. The flexibility of the method allows for straightforward adaptation to different bacterial species and experimental conditions. Thanks to the feasibility of the approach, the method can easily be adapted by other laboratories. Finally, we believe that this method has a great potential to generate insights into the complicated life of bacteria. Key words smFISH, Bacteria

1

Introduction In recent years, it has become clear that bacteria are intrinsically characterized by a high degree of phenotypic heterogeneity within otherwise isogenic populations [1–4], i.e., they can show a reversible bimodal gene expression, termed bistability, where two distinct subpopulations coexist, expressing genes at discrete and different levels. The causes for these substantial differences are thought to be stochastic gene expression and variable environmental factors, and the consequences of such phenotypic variability can result in a difference in fitness among the cells composing the population, a phenomenon called “bet-hedging,” or in a division of labor where specialized functions are undertaken by subpopulations of cells [5–7]. In this scenario, techniques that allow the bacterial interrogation at the single-cell level are pivotal. The advantages that a microscopy technique like single-molecule fluorescence in situ hybridization (smFISH) grants are several: the single-cell resolution, the maintenance of the spatial dimension, and the detection of lowly transcribed mRNAs. In fact, even with the recent development in scRNA-seq, dropout events are a major drawback: only

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Camilla Ciolli Mattioli and Roi Avraham

transcripts with more than ten copies per cell can be detected in scRNA-seq of pure microorganism cultures [8]. FISH is a powerful technique which allows the visualization of mRNA molecules at the subcellular level in fixed samples, via probing with fluorescent oligos using widefield fluorescence microscopy. With the occurrence of single-molecule FISH (smFISH) [9, 10], this technique became quantitative: the visualization of individual transcripts as diffraction-limited spots is enabled by the use of 10–50 DNA probes labeled with fluorescent dyes complementary to the target RNA that yields high signal-tonoise ratio, providing an estimate of mRNA copy number in the cell. Different techniques stemmed from the original smFISH (see Table 1 and Fig. 1), with alterations in three main directions: lower the costs, amplify the signal and improve the contrast, and move Table 1 FISH methodologies Name

Concept

Multiplexing

References

smFISH

10–50 probes per mRNA; probes are labeled

Lowthroughput

[9]

smiFISH

Primary probes are unlabeled and carry a common extra sequence as readout for the labeled secondary probe

Lowthroughput

[11]

FISH-STICs Based on amplifier, detectors, and three round of hybridization

Lowthroughput

[33]

bDNA

Based on preamplifier, amplifier, detectors, and four rounds of hybridization

Lowthroughput

[34]

HCR

Based on metastable hairpins that upon the presence of the Lowthroughput initiator (primary probe) will start a hybridization chain reaction

[35]

Padlock probes

Enzyme-based: Hybridized padlock probes can circularize Lowthrough ligation and be template for rolling-circle throughput amplification

[12]

ClampFISH Click-chemistry-based: Hybridized padlock probes can Low1.0 circularize through click-chemistry and be template for throughput the next round of padlock hybridization

[36]

LowClampFISH Click-chemistry-based: Hybridized padlock probes can throughput 2.0 circularize through click-chemistry and be template for the next round of padlock hybridization

[37]

seqFISH

25,000 genes visualized in eight sequential rounds of hybridization. Error prone

Highthroughput

[13]

MERFISH

Allows for error detection or detection and correction. High1001 genes are visualized in 14 rounds of hybridization throughput

[14]

seqFISH+

60 pseudocolors allow the detection of 24,000 genes in 4 sequential rounds of hybridization

[15]

Highthroughput

smFISH in Bacteria

5

Fig. 1 Variations in smFISH technique. (a) Classic smFISH diagram. 10–50 fluorescent probes bind the target mRNA. (b) Inexpensive smFISH (smiFISH) diagram. mRNA-specific primary probes hybridize to the targeted mRNA; labeled probes can be built on the readout sequence of the primary probes. (c), FISH with sequential tethered and intertwined oligodeoxynucleotides complexes (FISH-STICs) diagram. An mRNA-specific primary probe hybridizes to the targeted mRNA; amplifiers and labeled probes can be built onto the pair, leading to signal amplification. (d) Branched DNA (bDNA) diagram. A pair of mRNA-specific primary probes hybridizes to the targeted mRNA; preamplifiers, amplifiers, and labeled probes can be built onto the pair, leading to signal amplification. (e) Hybridization chain reaction (HCR) diagram. mRNA-specific primary probe hybridizes to the targeted mRNA and serves as initiator for the HCR. Metastable fluorescent RNA hairpins self-assemble into fluorescent amplification polymers upon detection of the initiator. (f) Padlock probes diagram. Locked nucleic

6

Camilla Ciolli Mattioli and Roi Avraham

from low-throughput imaging of up to five mRNAs together to high-throughput where thousands of mRNAs can be visualized by sequential rounds of hybridization. The first category aimed at lowering the costs of smFISH is based on indirect labelling schemes, where the primary probes are unlabeled and composed of two parts: an mRNA-specific sequence that hybridizes to the target and a readout sequence where a secondary labeled probe is hybridized. This allows the synthesis of primary probes at lower costs [11]. The second category aimed at amplifying smFISH signal enables a 10- to 100-fold signal enhancement and encompasses several different techniques that relies either on successive hybridizations of preamplifier, amplifier and detectors to increment the number of detection probes that contribute to the signal (FISHSTICs, bDNA, HCR, and ClampFISH), or enzymatic reactions (ligation and rolling-circle amplification) to produce more template for the hybridization of the detection probes (padlock probes). These techniques come particularly handy when the target RNAs are very short (i.e., bacterial transcripts); in fact, padlock probes have been successfully employed even for the detection of single micro RNA (miRNA) molecules 22 nt long [12]. Multiplexing and high-throughput smFISH analysis are enabled by a method called sequential FISH (seqFISH), based on sequential barcoding [13]. In this approach, during each round of hybridization, the previous probes are removed by DNAse digestion, freeing the previously assigned color channels. With this method, as little as four dyes and eight rounds of hybridization achieve the visualization of the whole transcriptome of eukaryotic cells (48 = 65,536). A drawback of such method, however, is that detection errors are not negligible, and they grow exponentially with the number of imaging rounds. A method directly targeted to detect and correct such detection errors is MERFISH [14], where an error-robust encoding scheme is employed, based on the Hamming code used in digital electronics, that allows to detect and correct the erroneously assigned mRNAs. These high-throughput methods are not trivial to implement, as they require extensive automation of the microscopy setup, which might not be feasible to achieve in many laboratories. Nevertheless, commercial solutions are now accessible: i.e., MERFISH ä Fig. 1 (continued) acid (LNA)-modified primers are used for cDNA synthesis, RNA is removed by RNase H, and cDNA is probed with padlock probes, followed by ligation and rolling-circle amplification (RCA). Fluorescent detection probes can then hybridize on the rolling-circle products. (g) Click-amplifying FISH (ClampFISH) diagram. Similar to the padlock method, it replaces the ligation step with a click-chemistry reaction between the alkyne and azide moieties present at the 5′ and 3′ ends of the padlock probes. To achieve exponential amplification, secondary padlock probes are designed to hybridize to two readout sequences present on the primary probe. The amplification can be repeated n times

smFISH in Bacteria

7

is available via the company Vizgen (https://vizgen.com/), and Xenium, based on padlock probes, is available at 10× Genomics (https://www.10xgenomics.com/platforms/xenium). However, the commercial solutions reach a multiplexing level of
500–1000 mRNAs only. In fact, an important issue that highthroughput smFISH faces is mRNAs crowdedness. This issue has been addressed with 2 different approaches: an analogical one, where the samples are physically expanded to achieve greater spatial resolution (expansion microscopy – ExFISH) [14], and a digital one, where 60 pseudocolors are used instead of 4 barcoding colors, reducing the chance of problematic overlapping signals [15]. The implementation of expansion microscopy allows to bring molecules in a diffraction-limited region to greater distances in an isotropic fashion, gaining higher spatial resolution. In ExFISH, RNA is covalently functionalized with a free-radical polymerizable group, LabelX [16], or through acrydite-modified poly(dT) locked nucleic acid (LNA) probe hybridized to the poly(A) tail of mRNAs [17]. Only the LabelX method is adaptable to bacteria [14], where the retention of RNA is independent of the polyadenylation status, opening up opportunities for studying bacterial transcriptomes at the single-cell level in a quantitative and high-throughput manner. smFISH has been successfully applied in several bacterial species, in low- and high-throughput formats. That is, a genome-wide study in E. coli showed that mRNA is spatially organized, where mRNAs encoding inner membrane proteins are localized at the membrane [18]. In another genome-wide study focusing on P. aeruginosa biofilms, it was found that oxygen availability shaped the metabolism at a spatial scale of microns [19]. These are only a few examples on how this technique can provide compelling insights in deciphering bacterial transcriptome regulation and population heterogeneity. However, the experimental implementation of smFISH is not trivial. In this chapter, we describe how to conduct a standard smFISH experiment [9] to study bacterial transcripts, meticulously describing each step required, from bacterial growth to image analysis.

2

Materials When performing smFISH, it is imperative to limit RNA degradation. Ensure that all consumables and reagents are RNase-free. The reagents and instruments needed for the whole protocol are listed below. Depending on the method of choice (ready-to-use or unlabeled probes), the dyes to perform the labelling reaction must be purchased separately (listed below).

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Camilla Ciolli Mattioli and Roi Avraham

Table 2 LGC Biosearch dyes

2.1

Probes Ordering

2.2 smFISH Probe Labelling

Dye

Absorbance (nm)

Emission (nm)

FAM

495

520

Cal Fluor Orange 560

538

599

Quasar 570

548

566

TAMRA

546

579

Cal Fluor Red 590

569

591

Cal Fluor Red 610

590

610

Cal Fluor Red 635

618

637

Quasar 670

647

670

Unlabeled probes: Probes are ordered from Biosearch Technologies with a 3′ C7 amine group at a synthesis scale of 5 nmol (see Note 1). Probes are shipped in a 96-well plate format, dissolved at 100 μM concentration. Let the plate thaw and spin it down. Per set of probes, pool the same volume from each well in a reservoir, mix well, and make 480 μL aliquots in 1.5 mL low binding Eppendorf for long-term storage at -20 °C. Each aliquot contains 48 nmols. Alternatively, probes can be ordered already labeled and ready-touse from Biosearch Technologies. The available dyes are listed in Table 2 (see Note 2). 1. Reaction buffer: 0.1 M sodium bicarbonate pH 9. Dissolve 0.42 gr of sodium bicarbonate in 50 mL of distilled water. Adjust the pH to 9 with 1 M sodium hydroxide. For 50 mL of solution, around 400 μL of 1 M sodium hydroxide are needed to reach pH 9. To avoid RNase contamination, use an aliquot of the solution to confirm the pH with a pH meter; adjust if necessary (see Note 3). Sterilize the solution by passing it through a 0.22 μm filter. 2. Dyes: Suggested dyes include Alexa Fluor 555, 594, or 647 NHS Ester (Succinimidyl Ester), 6-TAMRA SE (6-carboxytetramethylrhodamine, succinimidyl ester) single isomer, and Cy3 or Cy5 Mono-Reactive Dye. All dyes can be directly resuspended in the reaction buffer (120 μL per mg of dye, to divide among four labelling reactions), with the exception of TAMRA. 1 mg of TAMRA is first resuspended in 10 μL of DMSO, to which 230 μL of reaction buffer are added (to divide among eight labelling reactions). Once they have dissolved, the dyes quickly lose reactivity; promptly proceed to the labelling reaction. As an alternative, because of some

smFISH in Bacteria

9

photophysical shortcoming of organic dyes (signal deterioration during photoexcitation and low multiplexing capability due to spectra overlap), quantum dots have been optimized in order to allow their use in FISH [20]. They provide exceptional photostability and more robust transcript quantification, thanks to enhanced brightness, particularly significant for 3D biological specimen where acquisition of a full z-stack might take up to tens of seconds. Moreover, they allow a broader level of multiplexing, as they are characterized by a greater multispectral tenability compared to organic dyes. See [20] for quantum dot synthesis and conjugation. 2.3 Fixation and Permeabilization

1. 10× phosphate buffered saline (PBS). 2. 16% paraformaldehyde (PFA). 3. Fixation solution: 3.7% PFA in 1× PBS (see Note 4). 4. 100% ethanol (see Note 5).

2.4

smFISH Staining

1. Vanadyl ribonucleoside complex (VRC) stock solution: 200 mM vanadyl ribonucleoside complex solution comes as a solution. Aliquot the solution in PCR tubes (100 μL per tube) and store at -20 °C (see Note 6). 2. E. coli tRNA stock solution: Dissolve the powder in deionized nuclease-free water to a concentration of 20 mg/mL, and aliquot in 500 μL vials. Store at -20 °C. 3. BSA stock solution: Dissolve the powder in deionized nucleasefree water to a concentration of 50 mg/mL. Filter-sterilize and aliquot in 40 μL aliquots. Store at -20 °C. 4. 20× saline sodium citrate (SSC) buffer: 3 M NaCl, 300 mM sodium citrate in water. Adjust the pH to 7.4, and store at room temperature (RT). 5. Hybridization buffer: 10% dextran sulfate, 25% formamide (see Note 7 and Note 8, Fig. 2), 1 mg/mL E. coli tRNA, 2× SSC, 2 mM VCR, and 0.02% BSA (50 mg/mL). Dextran sulfate is difficult to dissolve; see Note 9 for instructions. Store the hybridization buffer at -20 °C in 250 μL aliquots. 6. Wash buffer: 2×SSC, 25% formamide (see Note 10). 7. DAPI stock solution: Dissolve the powder in DMSO to a concentration of 10 mg/mL (1000× stock). Store at -20 °C in 50 μL aliquots in amber tubes. 8. Wash buffer with DAPI: wash buffer supplemented with 10 μg/mL DAPI. 9. 3 M sodium acetate, pH 5.5. 10. TE buffer: 10 mM TRIS 1 mM EDTA, pH 8.

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Camilla Ciolli Mattioli and Roi Avraham

Fig. 2 Formamide allows relaxation of RNA secondary structures. (a) Formamide favors the relaxation of RNA secondary structures by destabilizing non-covalent bonds. (b) Optimal formamide concentration results in the highest number of probes specifically bound to the mRNA target

11. Glucose oxidase stock: Dissolve the glucose oxidase in 50 mM sodium acetate pH 5.2 to a concentration of 3.7 mg/mL at 37 °C for 1 h. Divide the solution in 5 μL aliquots and store them at -20 °C. 12. Catalase from Aspergillus niger stock solution: 13.8 mg/mL in ammonium sulfate suspension. Keep at 4 °C; do not freeze. 13. Mounting buffer without enzymes: 10 mM Tris pH 8, 2× SSC, 0.4% glucose (see Note 11). Divide the solution in 100 μL aliquots and store them at -20 °C. 14. Mounting buffer (GLOX anti-fade buffer): Thaw a 100 μL aliquot, and add 1 μL of glucose oxidase and 1 μL of catalase. 15. Coverslips of thickness no. 1 (see Note 12). 16. Polylysine coated microscope slides.

smFISH in Bacteria

11

17. Thermomixer. 18. SpeedVac concentrator. 19. Nanodrop. 20. Microcentrifuge. 2.5 Microscope Setup

Imaging is performed on an inverted epifluorescence microscope (e.g., Eclipse Ti2-E, Nikon), equipped with a ×100 NA 1.45 oil-immersion phase-contrast objective, suitable filter sets (DAPI, GFP, Cy3, mCherry, Cy5/Cy5.5), and a CMOS camera (e.g., Prime 95C, Photometrics).

2.6 Computer Requirements

Bacterial cell segmentation is performed using the machinelearning software Ilastik [21]. Ilastik requires a 64-bit machine, with at least 8 GB of RAM. Bacterial mRNA quantification is performed using the software Sp€atzcell [22]. To run Sp€atzcells, MATLAB (The MathWorks) and the MATLAB Image Processing and Optimization toolboxes are required. Sp€atzcells was developed using MATLAB versions 7.10 (R2010a) and 7.13 (2011b) and run on Intel machines with 3GHz Core 2 Quad CPUs and 4–16 GB of RAM.

3

Methods To perform a successful smFISH experiment and be able to troubleshoot, it is helpful to include a positive and a negative control. A positive control is a strain of bacteria engineered to carry a plasmid where the expression of the gene of interest (GOI) is inducible. When the GOI endogenous expression levels are unknown, the inclusion of this type of positive control will allow to trust any lack of signal in the sample of interest. A negative control can either be a sample not hybridized with the probe set, or a strain of bacteria deleted in the GOI, when the GOI is not essential. This second type is the preferred one, since it will also allow to test for unspecificity of binding of the probe set. Another type of control that allows to test for the binding specificity of the probe set consists in having the probe set divided in two sets, labeled with different fluorophores. That is, if the probe set is composed of 50 oligos, divide the oligos in two groups of 25 oligos each, and label the two groups with two different fluorophores. If the oligos are specific to the target, these two sets will produce co-localized signal in the two channels; if the oligos are binding to mRNAs other than the target, the signal will not co-localize.

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Camilla Ciolli Mattioli and Roi Avraham

Fig. 3 Formamide optimization. (a) A formamide concentration of 25% reaches the maximum of corrected total cell fluorescence (CTCF). (b) Depending on the total amount of probes per mRNA, the signal of real mRNA peaks will separate from the background at different folds. The dotted line indicates the 99% percentile of the Δ control. (c), mRNA quantification per cell ( = mean, σ = standard deviation) using three different formamide concentrations shows that above 25% formamide the annealing of the probes is hindered 3.1 smFISH Probe Design

Our protocol has been optimized for oligos that are 20 nucleotides in length and have a GC content
45%. With these parameters, we found that the optimal formamide percentage to use is 25% (Fig. 3a, c). Adjust the stringency of the hybridization and wash solutions by changing the concentration of formamide if the probe set diverges from this standard. The minimum amount of probes we successfully tested via microscopy is 15. Depending on the size of the probe set, the true positive signal will separate from the background by several folds. For example, for a probe set composed of 26 oligos, the true mRNA peaks showed a 5-fold increase in fluorescence compared to background peaks, and for a probe set composed of 37 oligos, the fold difference reached 10 (Fig. 3b). Probes are designed using the stellaris probe designer available at https://www.biosearchtech. com/support/tools/design-software/stellaris-probe-designer. For a few model organisms (human, mouse, rat, D. melanogaster, C. elegans), the software includes a parameter called “Masking” that improves probe specificity by using genome information. For bacteria, assessment of probe specificity requires an additional step (see below). 1. When possible, for each target mRNA design 48 oligos. 2. Input the sequence for your gene of interest in FASTA format, and use the default parameters to start:

smFISH in Bacteria

13

(a) Max number of probes = 48 (b) Oligo length = 20 (c) Min spacing length = 2 (d) Masking level = 2 (this level of masking only avoids problematic RNA structures) 3. If the target gene is too short or the sequence is problematic to provide enough probes, default settings can be tuned to reduce the length of the oligos and/or reduce the spacer (oligos shorter than 18 nt are not recommended as specificity drops). 4. Once the set of probes has been generated, make a FASTA file with all the probes and BLAST against your model organism genomic sequence. Eliminate the probes that have off-targets on other transcripts (with 20-nt probe, eliminate all the probes with >17 nt match to off-targets). If the sample of interest includes eukaryotic cells (i.e., intracellular bacteria inside macrophages), use masking level = 5 in the previous step during probe generation via stellaris, selecting the correct eukaryotic model organism. 3.2 smFISH Probe Labelling 3.2.1 Oligo Dehydration (
5 h)

Aliquots of pooled oligos as prepared in Subheading 2.1 are dehydrated as follows: 1. Place the Eppendorfs containing the 480 μL oligo pools in a SpeedVac concentrator with the following settings: (a) V-AQ for aqueous solutions (b) High temperature 2. Evaporate the solution completely. 3. Reconstitute the oligos in 70 μL of 0.1 M sodium bicarbonate.

3.2.2 Probe Labelling Reaction (Overnight)

The reaction between the carboxyl acid of the dye and the amine of the oligos will result in an amide bound, with the loss of a water molecule. Considering the nmol of the oligo library (48 nmol), and a
10-fold excess of dye per reaction, 1 mg of dye is sufficient for four labelling reactions, with the exception of TAMRA where 1 mg is sufficient for eight labelling reactions 1. Resuspend the dyes as indicated in Subheading 2.2. 2. Add the resuspended dye solution to the oligo pool (final volume 100 μL). 3. Incubate at 37 °C overnight (O/N) on a thermoblock with gentle shaking.

3.2.3 Probe Precipitation (2 h to O/N)

1. Stop the labelling reaction by adding 1:10 volume of 3 M sodium acetate pH 5.2. Mix the solution thoroughly by pipetting.

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Camilla Ciolli Mattioli and Roi Avraham

2. Add 3× volumes of 100% ethanol. Mix the solution thoroughly by inversion. 3. Place the mixture at -80 °C for at least 3 h, up to O/N. SAFE STOPPING POINT. 4. Centrifuge the tube at max speed (20,000 × g) for 30 min at 4 °C. 5. Decant the supernatant. Remove the remaining liquid using a Kimwipe, while avoiding touching the pellet. 6. Wash 3× with freshly prepared 70% ethanol. 7. Dissolve the pellet in 50 μL of TE to obtain a 1 mM solution (1000× stock). The working concentration is 1 μM. Therefore, dilute 1:100 in TE to have a 10 μM stock. From the 10 μM stock, 5 μL of probe will be used per reaction in 50 μL hybridization buffer. 8. Aliquot in small volumes to avoid thawing/freezing cycles. 3.2.4 Efficiency of Labelling Estimation (1 h)

The relative efficiency of a labelling reaction can be evaluated by calculating the approximate ratio of bases to dye molecules. This ratio can be determined, as described below, by measuring the absorbance of the nucleic acid at 260 nm and the absorbance of the dye at its absorbance maximum. The calculations are based on the Beer-Lambert law (Eq. 1): A = ϵbC

ð1Þ

where A is the absorbance, ϵ is the molar absorptivity, b is the length of light path, and C is the concentration. To calculate the molarity for oligos and dyes, use the microarray application of nanodrop. The application reports nucleic acid concentration, an A260/A280 ratio, and the concentrations and measured absorbance values of the dye(s), allowing detection of dye concentrations as low as 0.2 picomole per microliter. 1. Blank with TE buffer, and measure the absorbance of the nucleic acid and dyes at 260 nm and at the maximal absorbance for the dye. The maximal absorbance values for the fluorophores are given in Table 3. To obtain an accurate absorbance measurement for the nucleic acid, it is necessary to account for the dye absorbance using a correction factor (CF260), as illustrated below. The CF260 values per dye are given in Table 3. 2. Calculate the accurate absorbance measurement for the nucleic acid with the following equation: A DNA ðcorrectedÞ = A DNA ðA Dye  CF260 Þ

ð2Þ

smFISH in Bacteria

15

Table 3 Properties of the dyes Dye

MW

nmol in 0.5 mg

Extinction coefficient

Absorbance max (nm)

CF260

Alexa 555

1250

400

155,000

555

0.04

Alexa 594

819.8

610

92,000

590

0.43

Alexa 647

1250

400

155,000

650

0

TAMRA

527.5

950

90,000

555

0.04

Cy5

1050

476

250,000

650

0.08

Where ADNA is the measured ssDNA absorbance at 260 nm, ADye is the dye absorbance at its maximal absorbance, and CF260 is the correction factor of the dye. 3. Calculate the μM of the probes with Eqs. 3 and 4: DNA ðμg=mLÞ =

A DNA ðcorrectedÞ × MW ssDNA × DF × 1000 ð3Þ εssDNA

where MWssDNA is the molecular weight of ssDNA (308.97 g/ mol/nt), DF is the dilution factor used for the measurement, ϵ ssDNA is the extinction coefficient of the oligos at 260 nm (10,750 for oligos with equal ratios of bases), and 1000 factor converts from g/L to μg/mL: DNA ðμMÞ =

DNA ðμg=mLÞ × 1000 len × MW ssDNA

ð4Þ

where len is the size of the oligos and 1000 factor converts from M to μM. 4. Calculate the μM of the dye with Eqs. 5 and 6: Dye ðμg=mLÞ =

A Dye × MW Dye × DF × 1000 εDye

ð5Þ

where MWDye is the molecular weight of the dye, DF is the dilution factor used for the measurement, and ϵ Dye is the extinction coefficient of the dye (see Table 3): Dye ðμMÞ =

Dye ðμg=mLÞ × 1000 MW ssDNA

ð6Þ

where 1000 factor converts from M to μM. 5. Calculate the labelling efficiency (LE) with Eq. 7: LE =

DNA ðμMÞ × 100 Dye ðμMÞ

ð7Þ

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Camilla Ciolli Mattioli and Roi Avraham

A labelling efficiency of at least 90% is expected. Lower efficiencies (
70%) also allow to perform successful experiments (especially with a probe set composed of more than 20 oligos), although the signal intensity will be lower and the separation from the background smaller. 3.3

Bacteria Culture

To keep the experiment conditions consistent among experiments, it is important to take the same amount of bacteria. To do so, bacteria numbers are normalized by measuring the OD600 and taking a volume corresponding to the following: Vol ðmLÞ =

3.4 Fixation and Permeabilization (2 h to O/N)

2 OD600

ð8Þ

Fixation can be performed with a standard method or a direct method. The standard method is as follows: 1. Collect a volume of sample calculated according to Eq. 8. 2. Pellet the cells for 5 min at 5000 × g, and remove the supernatant. 3. Resuspend each pellet in 1 mL of fixation solution, and incubate for 30 min at RT (see Note 13). 4. Pellet the cells for 5 min at 1000 × g, and remove the supernatant. 5. Wash with 1× PBS. 6. Resuspend the cells, first in 300 μL of water, and then add 700 μL of 100% ethanol. Dissolving in water first prevents the formation of cell aggregates. Incubate 1 h at RT or O/N at 4 ° C to permeabilize the cells. SAFE STOPPING POINT. Samples can be left in 70% ethanol for a limited amount of time (up to 2 weeks) at 4 °C. Alternatively, the direct method is as follows: 1. PFA can be added directly to the liquid cultures (volumes corresponding to an equal number of cells as per Eq. 8) to reach a final concentration of 3.7%. 2. Incubate 30 min at RT. 3. Pellet the cells for 5 min at 1000 × g, and remove the supernatant. 4. Wash with 1× PBS. 5. Resuspend the cells first in 300 μL of water, and then add 700 μL of 100% ethanol. Dissolving in water first prevents the formation of cell aggregates. Incubate for 1 h at RT or O/N at 4 °C to permeabilize the cells. SAFE STOPPING POINT.

smFISH in Bacteria

17

Table 4 Multiplexing of dyes Dye

Absorbance (nm)

Emission

555

580

Alexa 594

590

617

Alexa 647 or Cy5

650

665

Alexa

3.5 smFISH Staining (O/N)

555

or TAMRA

1. Pellet the permeabilized cells in 70% ethanol for 5 min at 1000 × g; remove the supernatant. 2. Rehydrate the cells in 2× SSC for 10 min at RT. 3. Per sample, prepare the hybridization buffer by adding 5 μL of each 10 μM probe in 50 μL hybridization buffer (see Note 14 and Table 4). 4. Pellet the cells for 5 min at 1000 × g; remove the 2× SSC buffer. 5. Slowly resuspend the cells in the hybridization buffer containing the probes; avoid forming bubbles. 6. Incubate O/N at 37 °C. After O/N incubation, stained cells can be placed at 4 °C for long-term storage (up to a month). SAFE STOPPING POINT.

3.6

Mounting (
2 h)

1. Transfer 5 μL of the hybridized samples to a 0.5 mL tube. The rest of the sample can be stored at 4 °C for months in case more imaging sessions are required. 2. Wash 1× with 200 μL of wash buffer, mix thoroughly, pellet for 5 min at 1000 × g, and discard the supernatant. 3. Wash 2× with 200 μ× of wash buffer, mix thoroughly, incubate for 30 min at 37 °C, pellet for 5 min at 1000 × g, and discard the supernatant. 4. Wash 1× with 200 μL of wash buffer with DAPI, mix thoroughly, incubate for 30 min at 37 °C, pellet for 5 min at 1000 × g, and discard the supernatant. 5. Wash 1× with 200 μL of 2×SSC, mix thoroughly, pellet for 5 min at 1000 × g, and discard the supernatant. 6. Resuspend the cells in 20 μL of mounting buffer. Before assembling the slides for imaging, pipette the cell suspension up and down multiple times to separate cell aggregates. 7. Place 2 μL on a slide and cover it with a coverslip. Let the samples sit for 1 h at 4 °C and then proceed to imaging.

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Camilla Ciolli Mattioli and Roi Avraham

Table 5 smFISH analysis software Name

Concept

Organism

Sp€atzcell

The copy number of the target mRNA is estimated from the total intensity of fluorescent foci in the cell, rather than from counting discrete spots

Prokaryotic Matlab

[22]

FISH-quant

1 peak = 1 mRNA

Eukaryotic Matlab

[38]

Rajlabimagetools 1 peak = 1 mRNA

Eukaryotic Matlab

[39]

ImageM

1 peak = 1 mRNA

Eukaryotic Matlab

[40]

TransQuant

1 peak = 1 mRNA

Eukaryotic Matlab

[41]

3.7 Microscopy Setup and Recording (
3 h)

Language References

1. Determine the optimal imaging settings using the sample with the highest expression level (see Note 15). 2. Confirm that with the chosen parameters dim foci are also detected in the negative control sample; otherwise, adjust the imaging parameters. 3. Per image, acquire z-slices at 200 nm spacing, for a total of
10 slices covering 2 μm. 4. Using automatic acquisition mode (to avoid repeated scanning of the same areas), acquire different field of views to image
1000 cells. Typically,
10–30 image positions would suffice, depending on the density. Avoid areas where bacteria are too dense, as these are problematic for cell segmentation.

3.8 Image Analysis (2–3 Days)

Most of the smFISH quantification software developed so far are tailored to mRNA quantification in eukaryotes, where mRNA crowdness is not critical, and quantification is based on the assumption “one diffraction-limited spot = one mRNA” (see Table 5). This concept however, cannot be assumed as true for bacterial system, where mRNA crowdness is an issue. For these reasons, Skinner and colleagues [22] developed a new method for bacterial mRNA quantification that is not based on counting discrete spots. In their method, the diffraction-limited spots – additionally to being identified – are also quantified in order to estimate the mRNA copy number of the target mRNA from the total fluorescent intensity of the foci. Briefly, in each z focal plane of an image stack, the local maxima are identified above a user-defined threshold based on the negative control (deletion (Δ) mutant control). Local maxima from different z-planes that correspond to the same spot are joined together, and the plane with the highest intensity is defined as the in-focus plane. In this plane, the fluorescence intensity profile of each spot is fit to a 2D Gaussian to estimate position, area, peak height, and spot intensity. Spots are assigned to cells using segmentation masks. False positive spots are discarded using

smFISH in Bacteria

19

Fig. 4 Example of yfia smFISH imaging in Salmonella typhimurium. Representative large field and close-up microscopy images of three samples (Δ control, medium expression, and high expression samples) are shown. mRNA, white; DAPI, blue. Single channels are shown in black. Scale bar, 10 μm. On the right side of each sample, distributions of total (per cell) mRNA

the negative control sample (Δ mutant control), setting a cutoff so that only spots brighter than the 99% of the spots detected in the negative control are considered true positives. Fluorescence corresponding to single RNA molecules is calculated using as reference the sample with the lowest transcriptional level. An example of FISH images and analysis is shown in Fig. 4. For detailed instructions, see [22]. The requirements to use this software are the following: 1. Δ mutant control for estimation of false positive foci. 2. Low-expression control to estimate the fluorescence of a single mRNA. To obtain such control, it is useful to perform RT-qPCR experiments prior to smFISH, in order to establish the range of expression of the GOI in different growth phases of the bacteria. For Salmonella typhimurium, the SalCom online tool (http://bioinf.gen.tcd.ie/cgi-bin/salcom_v2.pl?_HL) can be used instead. 3. Cell masks Bacterial cells can be segmented using the machinelearning software Ilastik [21], following the “pixel and object” classification workflow. As input for Ilastik, the phase-contrast

20

Camilla Ciolli Mattioli and Roi Avraham

channel is used. The Ilastik binary masks can be further processed with a custom script aimed at removing spurious objects (small and big objects) and at curating the segmentation of high-density regions via a watershed algorithm. Alternatively, cells can be segmented using [23].

4

Notes 1. It is recommended to order the probes with reversed phase cartridge (RPC) purification method to exclude any probe missing the 3′ C7 amino group, necessary for the labelling reaction with the NHS ester group present on the dye. 2. Note that due to the variable autofluorescence in the green channel inherent to some cells and tissues, it is recommended to not use FAM. 3. The labelling reaction is extremely sensitive to pH (optimal range 8.5–9.3); therefore, the reaction buffer (0.1 M sodium bicarbonate) should be freshly prepared before use, as pH changes upon long-term storage. 4. This type of fixation, involving aldehydes, belongs to the additive fixation category (also called cross-linking fixation) and involves the creation of covalent chemical bonds with proteins: the aldehyde group of the fixative reacts with nearby proteins to form methylene bridge adducts, ultimately forming a matrix where membrane and lipids are trapped and the cell morphology is preserved. Alternative fixatives are alcohols (methanol and ethanol), belonging to the denaturing or precipitating category, that work by reducing protein solubility and by disrupting protein hydrophobic interactions, thus modifying their tertiary structures. Methanol fixation is used in Turbo FISH [24]; however, due to the serious cell shrinkage it causes [25], it is not recommended to use in bacteria. 5. This combination of fixation and permeabilization is efficient with several Gram-negative and some Gram-positive bacteria. However, this step should be optimized experimentally especially when working with Bacteroidetes and/or Gram-positive bacteria. In fact, a standardization of the fixation and permeabilization procedures is lacking for microorganisms. Depending on the characteristics of the membrane and cell wall, optimal fixation length and permeabilization conditions should be tested empirically [26]. A reference guide is provided in [27], where it was found that the best working condition for most of the tested bacteria (Gram-positive and Gram-negative) was a combination of PFA, acting both as a fixative and as a weak detergent [28], and ethanol, achieving permeabilization by promoting the solubilization of cell envelope components

smFISH in Bacteria

21

[26]. Modifications from this standard include the use of an enzymatic digestion of the peptidoglycan wall by the lysozyme and digestion of cell wall proteins by proteases [29, 30] (applied for the Gram-positive Listeria monocytogenes), the use of detergents like Triton ×-100 inducing a channel-forming effect via the interaction and substitution of cell envelope lipid molecules [31] (applied for the Gram-positive Bacillus pumilus spores), and short-term incubation in lactic acid or hydrochloric acid inducing the release of lipopolysaccharide from the outer membrane [32] (applied for the Gram-negative Pseudomonas aeruginosa, E. coli, and Salmonella enterica). 6. 200 mM vanadyl ribonucleoside complex stock solution is reconstituted to a green-black clear solution by incubating the sealed vial at 65 °C in a water bath for 10 min. Aliquot the solution in PCR tubes (100 μL per tube). The solution can be stored at -20 °C. 7. The hybridization buffer can contain different percentages of formamide, to be determined experimentally based on probe design. Formamide relaxes the secondary structures of RNA, and its concentration in the hybridization and wash solutions is the main parameter that controls the stringency of probe binding (Fig. 2). Low formamide concentration (low stringency) favors nonspecific binding of probes and does not relax secondary structures. Increasing the formamide concentration (high stringency) reduces nonspecific binding but, eventually, also reduces the binding of probes to the target mRNA. The correct percentage of formamide to be used depends on the GC content and length of the probes and should be determined experimentally. 8. Formamide is toxic and teratogenic. Handle it in a fume hood while wearing protective gloves and a lab coat. 9. To prepare the hybridization buffer, first dissolve the dextran sulfate in water in a falcon on a roller for about 30 min. Once the dextran sulfate is in solution, add all the remaining components. 10. Prepare fresh; adjust to the desired formamide concentration. 11. The mounting buffer can be prepared without the glucose oxidase and catalase enzymes and stored at -20 °C in 100 μL aliquots. 12. Coverslips of thickness no. 1 are recommended to use, as other commonly used coverslips (e.g., thickness no. 1.5) show a higher fluorescent background. 13. Formaldehyde is highly toxic and a known carcinogen. Handle it under a fume hood while wearing a lab coat and protective gloves, and dispose of according to proper safety and environmental regulations.

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14. The choice of the dye for probe labelling depends on the microscopy setup and the level of multiplexing that such setup allows. With the settings reported in 2.5, a combination of dyes as in Table 4 can be used altogether. As quality control, to exclude the presence of cross talk, prepare single-color labeled samples and image under all the different filter sets. 15. A good rule-of-thumb is to choose exposure times that produce foci pixel values around 30% of the maximum pixel value of the camera (maximum value of 65,535 for a 16-bit camera). At the same time, exposure times should not reach above 300 ms to avoid photobleaching. References 1. Ackermann M (2015) A functional perspective on phenotypic heterogeneity in microorganisms. Nat Rev Microbiol 13(8):497–508 2. Evans CR, Kempes CP, Price-Whelan A, Dietrich LE (2020) Metabolic heterogeneity and cross-feeding in bacterial multicellular systems. Trends Microbiol 28(9):732–743 3. Kopf SH, McGlynn SE, Green-Saxena A, Guan Y, Newman DK, Orphan VJ (2015) Heavy water and (15) N labelling with NanoSIMS analysis reveals growth rate-dependent metabolic heterogeneity in chemostats. Environ Microbiol 17(7):2542–2556 4. Schreiber F, Littmann S, Lavik G, Escrig S, Meibom A, Kuypers MM, Ackermann M (2016) Phenotypic heterogeneity driven by nutrient limitation promotes growth in fluctuating environments. Nat Microbiol 1(6): 16055 5. Rosenthal AZ, Qi Y, Hormoz S, Park J, Li SHJ, Elowitz MB (2018) Metabolic interactions between dynamic bacterial subpopulations. elife 7:e33099 6. Armbruster CR, Lee CK, Parker-Gilham J, de Anda J, Xia A, Zhao K, Murakami K, Tseng BS, Hoffman LR, Jin F, Harwood CS (2019) Heterogeneity in surface sensing suggests a division of labor in Pseudomonas aeruginosa populations. elife 8:e45084 7. Diard M, Garcia V, Maier L, RemusEmsermann MN, Regoes RR, Ackermann M, Hardt WD (2013) Stabilization of cooperative virulence by the expression of an avirulent phenotype. Nature 494(7437):353–356 8. Zhang Y, Gao J, Huang Y, Wang J (2018) Recent developments in single-cell RNA-Seq of microorganisms. Biophys J 115(2):173–180 9. Femino AM, Fay FS, Fogarty K, Singer RH (1998) Visualization of single RNA transcripts in situ. Science 280(5363):585–590

10. Raj A, van den Bogaard P, Rifkin SA, van Oudenaarden A, Tyagi S (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5:877–879 11. Tsanov N, Samacoits A, Chouaib R, Traboulsi AM, Gostan T, Weber C, Zimmer C, Zibara K, Walter T, Peter M, Bertrand E, Mueller F (2016) smiFISH and FISH-quant – a flexible single RNA detection approach with superresolution capability. Nucleic Acids Res 44(22): e165–e165 12. Larsson C, Grundberg I, So¨derberg O, Nilsson M (2010) In situ detection and genotyping of individual mRNA molecules. Nat Methods 7(5):395–397 13. Lubeck E, Coskun AF, Zhiyentayev T, Ahmad M, Cai L (2014) Single-cell in situ RNA profiling by sequential hybridization. Nat Methods 11(4):360–361 14. Chen F, Tillberg PW, Boyden ES (2015) Expansion microscopy. Science 347(6221): 543–548 15. Eng CHL, Lawson M, Zhu Q, Dries R, Koulena N, Takei Y, Yun J, Cronin C, Karp C, Yuan GC, Cai L (2019) Transcriptome-scale super-resolved imaging in tissues by RNA seqFISH+. Nature 568(7751): 235–239 16. Chen F, Wassie AT, Cote AJ, Sinha A, Alon S, Asano S, Daugharthy ER, Chang JB, Marblestone A, Church GM, Raj A, Boyden ES (2016) Nanoscale imaging of RNA with expansion microscopy. Nat Methods 13(8): 679–684 17. Wang G, Moffitt JR, Zhuang X (2018) Multiplexed imaging of high-density libraries of RNAs with MERFISH and expansion microscopy. Sci Rep 8(1):1–13 18. Moffitt JR, Pandey S, Boettiger AN, Wang S, Zhuang X (2016) Spatial organization shapes

smFISH in Bacteria the turnover of a bacterial transcriptome. elife 5:e13065 19. Dar D, Dar N, Cai L, Newman DK (2021) Spatial transcriptomics of planktonic and sessile bacterial populations at single-cell resolution. Science 373(6556):eabi4882 20. Liu Y, Le P, Lim SJ, Ma L, Sarkar S, Han Z, Murphy SJ, Kosari F, Vasmatzis G, Cheville JC, Smith AM (2018) Enhanced mRNA FISH with compact quantum dots. Nat Commun 9: 4461 21. Berg S, Kutra D, Kroeger T, Straehle CN, Kausler BX, Haubold C, Kreshuk A (2019) Ilastik: interactive machine learning for (bio) image analysis. Nat Methods 16(12): 1226–1232 22. Skinner SO, Sepu´lveda LA, Xu H, Golding I (2013) Measuring mRNA copy number in individual Escherichia coli cells using singlemolecule fluorescent in situ hybridization. Nat Protoc 8(6):1100–1113 23. Young JW, Locke JC, Altinok A, Rosenfeld N, Bacarian T, Swain PS, Mjolsness E, Elowitz MB (2012) Measuring single-cell gene expression dynamics in bacteria using fluorescence timelapse microscopy. Nat Protoc 7:80–88 24. Shaffer SM, Wu MT, Levesque MJ, Raj A (2013) Turbo FISH: a method for rapid single molecule RNA FISH. PLoS One 8(9):e75120 25. Chao Y, Zhang T (2011) Optimization of fixation methods for observation of bacterial cell morphology and surface ultrastructures by atomic force microscopy. Appl Microbiol Biotechnol 92(2):381–392 26. Felix H (1982) Permeabilized cells. Anal Biochem 120(2):211–234 27. Rocha R, Almeida C, Azevedo NF (2018) Influence of the fixation/permeabilization step on peptide nucleic acid fluorescence in situ hybridization (PNA-FISH) for the detection of bacteria. PLoS One 13(5):e0196522 28. Cheng R, Zhang F, Li M, Wo X, Su YW, Wang W (2019) Influence of fixation and permeabilization on the mass density of single cells: a surface Plasmon resonance imaging study. Front Chem 7:588 29. Wagner M, Schmid M, Juretschko S, Trebesius K-H, Bubert A, Goebel W, Schleifer K-H (1998) In situ detection of a virulence factor mRNA and 16S rRNA in listeria monocytogenes. FEMS Microbiol Lett 160(1):159–168 30. Furukawa K, Hoshino T, Tsuneda S, Inamori Y (2006) Comprehensive analysis of cell wallpermeabilizing conditions for highly sensitive

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fluorescence in situ hybridization. Microbes Environ 21(4):227–234 31. Mohapatra BR, La Duc MT (2012) Evaluation of fluorescence in situ hybridization to detect encapsulated Bacillus pumilus SAFR-032 spores released from poly(methylmethacrylate). Microbiol Immunol 56(1):40–47 32. Alakomi HL, Skytt€a E, Saarela M, MattilaSandholm T, Latva-Kala K, Helander IM (2000) Lactic acid permeabilizes gramnegative bacteria by disrupting the outer membrane. Appl Environ Microbiol 66(5): 2001–2005 33. Sinnamon JR, Czaplinski K (2014) RNA detection in situ with FISH-STICs. RNA 20(2): 260–266 34. Battich N, Stoeger T, Pelkmans L (2013) Image-based transcriptomics in thousands of single human cells at single-molecule resolution. Nat Methods 10(11):1127–1133 35. Choi HM, Chang JY, Trinh LA, Padilla JE, Fraser SE, Pierce NA (2010) Programmable in situ amplification for multiplexed imaging of mRNA expression. Nat Biotechnol 28(11): 1208–1212 36. Rouhanifard SH, Mellis IA, Dunagin M, Bayatpour S, Jiang CL, Dardani I, Symmons O, Emert B, Torre E, Cote A, Sullivan A, Stamatoyannopoulos JA, Raj A (2018) ClampFISH detects individual nucleic acid molecules using click chemistry–based amplification. Nat Biotechnol 37(1):84–89 37. Dardani I, Emert BL, Goyal Y, Jiang CL, Kaur A, Lee J, Rouhanifard SH, Alicea GM, Fane ME, Xiao M, Herlyn M, Weeraratna AT, Raj A (2022) ClampFISH 2.0 enables rapid, scalable amplified RNA detection in situ. Nat Methods 19(11):1403–1410 38. Mueller F, Senecal A, Tantale K, Marie-NellyH, Ly N, Collin O, Basyuk E, Bertrand E, Darzacq X, Zimmer C (2013) FISH-quant: automatic counting of transcripts in 3D FISH images. Nat Methods 10(4):277–278 39. Raj A, Peskin CS, Tranchina D, Vargas DY, Tyagi S (2006) Stochastic mRNA synthesis in mammalian cells. PLoS Biol 4(10):1707–1719 40. Lyubimova A, Itzkovitz S, Junker JP, Fan ZP, Wu X, Van Oudenaarden A (2013) Singlemolecule mRNA detection and counting in mammalian tissue. Nat Protoc 8(9): 1743–1758 41. Halpern KB, Itzkovitz S (2016) Single molecule approaches for quantifying transcription and degradation rates in intact mammalian tissues. Methods 98:134–142

Chapter 2 Single-Molecule Fluorescent In Situ Hybridization (smFISH) for RNA Detection in the Fungal Pathogen Candida albicans Sander van Otterdijk, Maryam Motealleh, Zixu Wang, Thomas D. Visser, Philipp Savakis, and Evelina Tutucci Abstract Candida albicans is the most prevalent human fungal pathogen. Its pathogenicity is linked to the ability of C. albicans to reversibly change morphology and to grow as yeast, pseudohyphae, or hyphal cells in response to environmental stimuli. Understanding the molecular regulation controlling those morphological switches remains a challenge that, if solved, could help eradicate C. albicans infections. While numerous studies investigated gene expression changes occurring during C. albicans morphological switches using bulk approaches (e.g., RNA sequencing), here we describe a single-cell and singlemolecule RNA imaging and analysis protocol to measure absolute mRNA counts in morphologically intact cells. To detect endogenous mRNAs in single fixed cells, we optimized a single-molecule fluorescent in situ hybridization (smFISH) protocol for C. albicans, which allows one to quantify the differential expression of mRNAs in yeast, pseudohyphae, or hyphal cells. We quantified the expression of two mRNAs, a cell cyclecontrolled mRNA (CLB2) and a transcription factor (EFG1), which show expression changes in the different morphological cell types and nutrient conditions. In this protocol, we described in detail the major steps of this approach: growth and fixation, hybridization, imaging, cell segmentation, and mRNA spot analysis. Raw data is provided with the protocol to favor reproducibility. This approach could benefit the molecular characterization of C. albicans and other filamentous fungi, pathogenic or nonpathogenic. Key words smFISH, Candida albicans, Single-molecule RNA FISH, RNA localization, Single-cell imaging, Fungal pathogen

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Introduction Single RNA molecule fluorescence in situ hybridization (smFISH) is a method that enables the detection and localization of mRNAs in fixed cells [1–3], reviewed in [4, 5]. This approach provides absolute quantifications of mRNA molecules in their cellular context, and it has been used, from prokaryotic to multicellular

Maryam Motealleh, Zixu Wang, and Thomas D. Visser have equally contributed to this chapter Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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eukaryotic organisms, to investigate various aspects of gene regulation including transcription, mRNA nuclear export, subcellular mRNA localization, and mRNA decay [6]. The ability to localize and count mRNAs in intact cells unraveled an extensive and, at times, unexpected degree of cell-to-cell heterogeneity in isogenic populations of cells that could not be inferred by using bulk mRNA measurement approaches. The latest and more widespread smFISH protocols use a mix of 48 fluorescently labelled 20-mer DNA oligonucleotides that hybridize to the target mRNA molecule. The accumulation of tens of fluorophores on the mRNA enables its visualization as a diffraction-limited spot using widefield or confocal microscopy. The use of DNA probes labelled with different fluorophores (e.g., Cyanine, Quasars, Alexa, CalFluors) allows the simultaneous detection of up to four mRNA species in single cells [7–9]. In addition, smFISH can be combined to immunofluorescence to detect mRNAs and proteins within the same cell [10–14]. In recent years, smFISH has been improved to the point that thousands of mRNA species per cell can be visualized in single cells, opening the era of spatial transcriptomics [4, 7–9, 15]. Those improvements included the development of brighter fluorophores and the combination of fluorescence microscopes with microfluidic systems that allowed automating protocols combining multiple fluorophore-labelled probes and many rounds of hybridization to high-resolution imaging. Progress in the field of single-molecule RNA imaging was also driven by the advancement of fast, interactive, multidimensional image viewing systems (e.g. Napari [16]), automated cell segmentation (e.g., Cellpose [17, 18], Startdist [19, 20], Omnipose [21]), and the quantification of RNA spots in thousands of cells (e.g., FISH-quant [22–24], StarFISH [25]). Various RNA FISH protocols are available to measure gene expression changes in fungal cells, but besides extensive work done in the model organism S. cerevisiae (e.g [11, 12, 26, 27].), few examples have been reported in other fungal species such as Schizosaccharomyces pombe [28], Neurospora crassa [29], Ashbya gossypii [30], and Ustilago maydis [31]. Another fungal species for which RNA FISH has been reported is Candida albicans [32, 33], the most common human fungal pathogen [34, 35]. The pathogenicity of C. albicans is linked to its ability to change morphology and switch from yeast to pseudohyphal or hyphal growth depending on the environmental conditions (e.g., nutrients, temperature, cell density) [36–38] (Fig. 1a). While C. albicans morphological changes have been extensively studied, limited tools are available to correlate C. albicans mRNA expression and morphological changes (e.g., cell elongation). Thus, we set out to design an optimized smFISH hybridization and quantification protocol taking advantage of the latest improvements in the field of microscopy and imaging analysis.

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Fig. 1 Summary of the smFISH procedure in C. albicans. (a) Schematic representation of C. albicans cell types. (b) Schematic of the C. albicans growth, fixation, and permeabilization steps. (c) Schematic of the smFISH hybridization steps

The smFISH protocol (i.e., cell growth, probes hybridization and microscopy) is performed over 3 days, followed by data analysis (Fig. 1b, c). The major steps are: Day 1, (i) coverslip coating (1 h); (ii) growth, fixation, and permeabilization of the cells (overnight growth +10 hs’ processing time); Day 2, (iii) smFISH hybridization and washes (6 h); Day 3, (iv) fluorescence microscopy; and (v) imaging and data analysis (time depends on the aim of the experiment, usually few days). This approach can be applied to different biological questions and model organisms. The Notes section includes considerations that may be useful to develop smFISH protocols for other filamentous fungi.

2 Materials Prepare all solutions using sterile double distilled ultrapure RNAsefree water (DDW). Use RNAse-free glassware and plastic

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containers. Paraformaldehyde and deionized formamide are hazardous solutions: wear protective gloves and handle them under a fume hood. 1. 0.1 N HCl. 2. 70% ethanol. 3. Non-coated coverslips: 0.17 mm thick (#1.5H); diameter 16 mm. 4. 12-well culture dish. 5. 0.01% poly-L-lysine. 6. Refrigerated centrifuges for 1.5 mL and 50 mL tubes. 7. Incubators: for Candia albicans cultures, 30–37 °C; for smFISH hybridization and washes, 37 °C. 8. Candida albicans strain used in this study: SC5314 (wild-type isolate). 9. Tryptic Soy Broth (TSB) medium, for 1 L: 17 g Tryptone (Pancreatic Digest of Casein), 3 g Soytone (Peptic Digest of Soybean), 2.5 g glucose, 5 g sodium chloride, and 2.5 g dipotassium phosphate. Adjust to pH 7.3 ± 0.2. Sterilize by autoclaving. 10. Spider medium, for 1 L: 3 g beef extract, 20 g Bacto Peptone, 10 g mannitol, 2 g dipotassium phosphate (K2HPO4), 50 μg arginine, 10 μg histidine, and 50 μg tryptophan. Sterilize by autoclaving. 11. Tryptic Soy Broth (TSB) agar plates, for 1 L: 17 g Tryptone (Pancreatic Digest of Casein), 3 g Soytone (Peptic Digest of Soybean), 2.5 g glucose, 5 g sodium chloride, and 2.5 g dipotassium phosphate. Adjust to pH 7.3 ± 0.2, and then add 15 g agar. Sterilize by autoclaving and pour plates when cooled to ~50 °C. 12. 32% (w/v) paraformaldehyde. Store at room temperature (RT) and protect from light. 13. 3 M D-sorbitol. Filter-sterilize and store at 4 °C. 14. 1 M K2HPO4. 15. 1 M KH2PO4. 16. 1 M potassium phosphate buffer pH 7.5, for 1 L: mix 834 mL of 1 M K2HPO4 with 166 mL of 1 M KH2PO4. 17. Buffer B: 1.2 M sorbitol, 100 mM potassium phosphate buffer pH 7.5, double distilled sterile water. Store at 4 °C. 18. 200 mM vanadyl-ribonucleoside complexes (VRC) stock solution. Dissolve the powder in the vial at 65 °C for 10 min and store aliquots at -20 °C. 19. Lyticase solution: resuspend 25 K units in 1 mL 50% glycerol, 1×PBS. Store 100 μL aliquots at -20 °C.

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20. Spheroplast buffer: 1.2 M sorbitol, 100 mM potassium phosphate buffer pH 7.5, 20 mM VRC, 20 mM β-mercaptoethanol. 21. Tris-EDTA buffer (TE buffer): 10 mM Tris-Cl (pH 8), 1 mM ethylenediaminetetraacetic acid (EDTA) (pH 8). 22. Competitor DNA/RNA: 10 mg/mL sheared salmon sperm DNA, 10 mg/mL E. coli tRNA (both RNAse-/DNAse-free). Store at -20 °C. 23. Vacufuge: to lyophilize smFISH probes. 24. 10 mm and 15 mm petri dishes. 25. Kimtech tissues. 26. 20× saline-sodium citrate (SSC) buffer: 3 M NaCl, 0.3 M sodium citrate-HCl, pH 7.0. 27. smFISH pre-hybridization solution: 10% formamide, 2× SSC. Prepare fresh. 28. 1 M Na2HPO4. 29. 1 M NaH2PO4. 30. 200 mM sodium phosphate buffer pH 7.5, for 1 L, mix 168 mL of 1 M Na2HPO4 and 32 mL of 1 M NaH2PO4. Next, add sterilized water to final volume of 1 L. 31. Solution F: 20% formamide, 10 mM sodium phosphate buffer pH 7.5. Prepare fresh. 32. Solution H: 4× SSC, 2 mg/mL BSA, 10 mM VRC. Prepare fresh. 33. 2× SSC: dilute 20× SSC 1:10 with DDW. Store at room temperature. 34. 1× SSC: dilute 20× SSC 1:20 with DDW. Store at room temperature. 35. 2× SSC/0.1% Triton ×-100 solution. 36. 1× phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4. 37. smFISH probes stock: resuspend lyophilized probes in TE buffer to a final concentration of 25 μM. Probe sequences are listed in Subheading 4 (see Note 1). 38. Mounting solution with the DNA staining agent 4′,6-diamidin-2-phenylindol (DAPI). 39. Transparent nail polish. 40. Widefield epifluorescence microscope (see Note 2). 41. Image-processing software Fiji (Java software for imageprocessing analysis; https://fiji.sc/) [39]. 42. Image viewer: Napari (https://napari.org/stable/) [16], which runs in Python (https://www.python.org/).

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43. Cell segmentation software: Cellpose2 (https://www.cellpose. org/) [18]. 44. smFISH spot quantification: FISH-quant for Python (https:// fish-quant.github.io/) [22, 24]. 45. Analysis tools (Jupyter notebooks) and raw data associated to this protocol can be found on Zenodo (https://zenodo.org/ records/10613839) in the repository called “C_albicans_smFISH.”

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Method

3.1 Coverslip Washing and Coating

1. Use a 1 L beaker to boil three packages (~300 pieces) of microscope coverslips in 500 mL of 0.1 N HCl for 15–20 min. Gently stir every 5 min to separate and wash the coverslips. 2. Rinse the coverslips extensively (~10 times) with DDW water, dry in a stove overnight, autoclave, and keep at 4 °C in 70% ethanol for up to a year. 3. On the day of the smFISH, place the coverslips on a clean chromatography paper, air-dry the ethanol and rinse with water once. Aspirate the excess water, and air-dry. Treat the coverslips for 20 min at room temperature (RT) with 200 μL of 0.01% (w/v) poly-L-lysine. Aspirate the poly-L-lysine solution and let the covers air-dry. Wash one time with DDW, aspirate the excess, and allow to air-dry. 4. Use forceps to place each coverslip, with the poly-L-lysine coated side up, into a single well of a 12-well culture dish, and store the dish at RT. Coverslips need to be completely dry before seeding the cells.

3.2 Growth, Fixation, and Permeabilization of C. albicans Cells

1. Thaw C. albicans cells from the -80 °C on a rich medium plate (TSB), and incubate overnight at 30 °C. 2. Grow a single C. albicans colony overnight (~15 h) in 10 mL of TSB medium at 30 °C. 3. In the morning, dilute cells in 1× PBS at a ratio of 1:100. Use a counting chamber with a Bu¨rker pattern to count the cells. 4. Dilute cells in fresh TSB medium or filamentous-inducing Spider medium to a final concentration of 105 cells/mL. Grow cells to mid-log phase. The time may differ depending on the medium and the biological question. In this protocol, cells were grown in TSB or Spider medium for 6 h (see Note 3). 5. After the desired time has passed, use a counting chamber with a Bu¨rker pattern to count the cells. 6. Fix the cells by adding paraformaldehyde at a final concentration of 4% (w/v). Mix gently and incubate at RT for 45 min with constant shaking (see Note 4).

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7. Centrifuge the cells at 2400 g for 3 min at 4 °C, and wash once with 10 mL of ice-cold buffer B. 8. Resuspend cells in 1 mL of ice-cold buffer B and transfer to a 1.5 mL tube. 9. Centrifuge cells at 2400 g for 3 min at 4 °C, remove all buffer B, and resuspend cells by pipetting in 480 μL of spheroplast buffer. Keep cells on ice. 10. Once cells are resuspended, add lyticase. Use 12.5 U of enzyme per 107 cells. Adjust the lyticase volume based on the counting performed at point 5. 11. Incubate the cells in a water bath at 30 °C for 3–5 min, inverting gently and frequently (see Note 5). 12. Every 2 min, during the lyticase treatment, take 5 μL sample on a slide, cover it with a coverslip, and observe the cells with bright field illumination. Prevent overdigestion (Fig. 2), by stopping the incubation earlier if necessary. 13. Centrifuge the cells for 4 min at 1300 g at 4 °C, and wash them once with 500 μL of ice-cold buffer B. Do not vortex but

Fig. 2 Lyticase digestion effect on cell morphology of C. albicans. (a) Phase contrast time-lapse montage during lyticase digestion. Full lyticase digestion causes darkening of cells (12 hours). For smFISH, digestion was stopped between 3 and 5 min, to avoid over-digestion. (b) Single plane DIC images (gray) of cells exposed to different degrees of lyticase digestion. Left panel, cells in spheroplast buffer before adding lyticase. Middle panel, cells post-digestion mounted on a slide after hybridization is completed with intact cell morphology. Right panel, cells post-digestion mounted on a slide after hybridization is completed after overdigestion. Scale bars, 20 μm

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resuspend the cells by gently pipetting. Cells are fragile after the lyticase treatment. 14. Resuspend the cells in 900 μL of ice-cold buffer B. Drop 150 μL on each poly-L-lysine treated coverslip. Incubate for 30–60 min at 4 °C to allow the cells to adhere to the coverslips (see Note 6). 15. Wash once each well with 2 mL ice-cold buffer B. Gently aspirate the buffer and add 2 mL of cold 70% ethanol. Seal the 12-well plate with Parafilm and store at -20 °C (see Note 7). 3.3

Hybridization

1. Move the coverslips for hybridization into a new 12-well plate. Rehydrate the cells with two washes of 2 mL of 2×SSC, each for 5 min at RT (see Note 8). 2. Incubate the coverslips in 2 mL of pre-hybridization solution for 30 min to 1 h at RT (see Note 9). 3. For each coverslip, combine 0.125 μL of the original stock of probes for the mRNA of interest (25 μM) with 5 μL of the ssDNA/tRNA competitor. Lyophilize in a Vacufuge at 45 °C (see Note 10). Probes are lyophilized at ~1 μL/min rate. Be careful to prevent overdrying as this makes it more difficult to resuspend the probemix. 4. Add 12.5 μL of solution F per coverslip (e.g., if you hybridize two coverslips, add 25 μL of solution F). Heat at 95 °C for 2 min. Let the solution cool at RT for about 5 min. Keep probes in the dark to prevent photobleaching. 5. Add 12.5 μL of solution H per coverslip (e.g., if you hybridize two coverslips, add 25 μL of solution H). The resulting hybridization solution (25 μL per coverslip) now contains 125 nM probe mixture and 10% formamide. 6. Cover the bottom of a 150 mm petri dish with Parafilm, and tape it to the bottom to keep the surface flat. 7. Transfer 23 μL of the hybridization solution from step 5 onto the Parafilm; avoid bubbles. One drop for each coverslip. By using the forceps, take the coverslip from each well, remove from each coverslip the leftover pre-hybridization solution by touching the edge to a Kimtech tissue, and place each coverslip face down onto the prepared hybridization drop. Place a small container (e.g., the cap of a 15 mL plastic tube) toward the edge of the Petri dish and fill it with DDW. Seal the petri dish with Parafilm, and cover it with aluminum foil to create a dark hybridization chamber. Incubate in the dark at 37 °C for 3 h. Incubate at 37 °C the pre-hybridization solution that you will use for the following washes (step 8). 8. Use forceps to place the coverslips, facing up, back into a 12-well plate containing 2 mL of pre-warmed

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pre-hybridization solution. Cover the 12-well plate with aluminum foil. Incubate for 15 min at 37 °C. 9. Aspirate the solution carefully and replace it with 2 mL of pre-warmed pre-hybridization buffer, and incubate again for 15 min at 37 °C. Wash once with 2 mL of 2× SSC for 5 min at RT. 10. Wash once with 2 mL 2× SSC/0.1% Triton ×-100 solution for exactly 5 mins at RT (see Note 11). 11. Wash once with 2 mL of 1× SSC for 5 min at RT. 12. Wash once with 2 mL of 1× PBS for 5 min at RT. 13. Before mounting, dip coverslip in 100% EtOH, and let them dry at RT covered from the light. 14. Add a drop of mounting solution with DAPI (~20 μL) on a glass slide. Be careful to avoid bubbles. Place the dry coverslip with the cells facing down onto the drop of mounting solution. Allow the mounting solution to polymerize at least overnight at RT in the dark. 15. Seal coverslips with transparent nail polish. Let nail polish dry completely before imaging to avoid damaging the microscope objective. 16. Go to the microscope and enjoy your images. Slides can be stored in the dark at 4 °C for a few days and at -20 °C for months. 3.4 Image Acquisition

1. Use a widefield epifluorescence microscope with a motorized scanning stage, a high numerical aperture objective ~1.4NA, and narrow band-pass filters (see Note 12). 2. Optically section cells using 200 nm Z steps, spanning 12–15 μm Z depth to encompass the entire cell thickness. 3. For smFISH performed using Quasar 670 (CY5 filter), Calfluor610 (CY3.5 filter), or Quasar 570 (CY3 filter), use an exposure time around 750 ms to acquire each Z plane and 100% light power (LED lamp). 4. For the DAPI channel, use 25–50 ms exposure and 12.5% light source power. 5. Acquire the differential interference contrast (DIC) image for 15–30 ms on a single plane. 6. To visualize the smFISH results, in Fiji, create a MAX projection of the Z-stacks where cells are in focus, both for the smFISH and the DAPI channels. Merge the MAX projected images to the corrected DIC image (see Subheading 3.5.1) (Fig. 3).

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Fig. 3 smFISH for the CLB2 and EFG1 mRNAs in C. albicans. (a) Merge maximally projected image: CLB2 mRNA smFISH Quasar 670 (yellow) and DAPI (blue) merged to a single plane DIC image (gray). (b) Merge maximally projected image: EFG1 mRNA smFISH Quasar 670 (green) and DAPI (blue) merged to a single plane DIC image (gray). For both (a) and (b), left panels, cells were grown at 37 °C in TSB medium for 6 h. Right panels, cells were grown at 37 °C in Spider medium for 6 h. Representative cell types, yeast (Y), and pseudohyphal (P) or hyphal (H) cells are indicated on the images. A representative single mRNA and site of mRNA transcription (transcription site, Txs) are indicated on the images. Scale bars, 5 μm 3.5

Imaging Analysis

3.5.1 Drift Correction of DIC Images

Cell segmentation can be performed using DIC images. In this case, a correction of the DIC image should be applied in order to align the fluorescence channels to the DIC image and correct the drift introduced by the DIC prism (Fig. 4). To this end: 1. Open Fiji. 2. Split the original fluorescence images into different fluorescence channels, and apply the Fiji function of the in-focus Z-stacks to all channels. 3. Open the DIC image.

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Fig. 4 Image segmentation and spot analysis for a representative C. albicans CLB2 smFISH picture. (a) Translated DIC picture used for cell segmentation using Cellpose. (b) Overlay of the translated DIC picture in (a) with the cell mask generated and corrected with Cellpose. Note: manual corrections were necessary to outline hyphal cells. Scale bar, 20 μm. (c) Enlargements of image crop (dashed yellow box) from (b) showing the original CLB2 smFISH channel, the overlay of the mask generated with Cellpose and the spots identified using Big-FISH, overlay of the spots (red circles) onto the CLB2 smFISH picture. Scale bars, 20 μm

4. Correct the drift of the DIC image by using the function path: Image> Transform > Translate. 5. Set the correct x and y offset (e.g., x,y 15, -15 pixels). 6. Verify the correct offset using the function path: Image > Color > Merge channels. 7. Examine the composite image to ensure that the smFISH and DAPI signals are correctly located within the cell. 8. If the composite image is not aligned correctly, repeat steps 4–7. 9. Save the corrected DIC image. 10. Use the function Process> Batch> Macro in Fiji to batchtranslate multiple DIC images.

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3.5.2 Cell and Nuclear Segmentation and Mask Generation

The corrected DIC image can now be used for cell segmentation using Cellpose [17, 18]. Cellpose can be run with its own graphical user interface (GUI) or can be run as a plug-in in Napari. Here, we describe the steps using Cellpose GUI, because this option allows to train models for creating cell masks. 1. Launch Cellpose GUI and import the DIC image by navigating to File > Load Image. 2. Specify the cell diameter manually or “calibration” feature, which estimates the cell diameter based on the input image (a good starting value is, e.g., 80 pixels). 3. Select the appropriate model for cell segmentation. It is recommended to utilize the “cyto” or “cyto2” models from the “model zoo.” The users can also apply a custom-trained model to perform cell segmentation. 4. Once the segmentation process is complete, the Cellpose GUI generates cell masks corresponding to the identified cells. These masks are binary images. If necessary, manual modifications can be applied in Cellpose (see Cellpose “Help” menu for shortcuts). 5. Once the cell masks are correct, select File > Save masks as “seg.npy.” This automatically saves the mask file in the same folder where the drifted DIC picture is located. 6. Repeat the segmentation process for the Nuclei mask. Import in Cellpose a MAX projection generated in Fiji of the in-focus Z-stacks from the DAPI channel. Select the “nuc” model from the “model zoo.” Correct the nuclei outlines if necessary. 7. Use provided Jupyter Notebook “01-npy_to_tif_conversion” to convert the saved masks from the “.npy” format to “.tiff” format. (see Note 13).

3.5.3 smFISH Spot Detection

The detection of mRNA spots is performed using FISH-quant for Python [24], using the library called “Big-FISH,” while images and the intermediate steps of spot identification are visualized using Napari. This allows to integrate spot detection with a fast, interactive, and multidimensional image viewer. 1. Start the Jupyter Notebook “02-spot_detection.” 2. Load the images and preview them in the Notebook. 3. Perform spot detection on filtered images. This step includes the following sub-steps: (a) Scaling the image size (b) Filtering the image (c) Performing spot detection

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(d) Inspecting the quality of the filtering and spot detection in the Napari viewer and adjusting the parameters in the Notebook if necessary 4. If satisfied, return to Jupyter Notebook “02-spot_detection” and save the spot data. They will be automatically saved in a new folder called “Spots.” 5. If you have multiple images, and you are satisfied with the parameters set for your first image, you can proceed to batchprocess your dataset. 3.5.4 Spot Decomposition

To measure the number of nascent RNAs at the sites of transcription (Txs) in nuclei, we make use of the Big-FISH function “spot decomposition.” This step allows us to estimate the number of RNAs in dense regions of fluorescence by first calculating the average intensity of mRNAs across cells and second to fit the 3D image of an average single mRNA into the dense fluorescence region. Spot decomposition is also used to estimate the number of nascent RNAs within transcription sites in the cell’s nucleus. For more documentation of the Big-FISH library, see https://big-fish. readthedocs.io/en/stable/index.html. 1. Open the Jupyter Notebook “03-spot_decomposition.” 2. Follow the steps to test the spot decomposition on a single image. 3. Use standard parameters set in Notebook “03-spot_decomposition” to decompose the fluorescence of dense spots regions. 4. Inspect the detected dense RNA regions in Napari. Make sure all Txs in the nuclei are correctly identified. 5. If satisfied with the parameters, proceed with the detection of nascent RNAs at transcription sites via the batch-processing function. The results will be automatically saved in a new folder called “spot decomposition.”

3.5.5 Assignment of mRNA Spots to Cell Masks

The following steps describe how to assign spots (mRNAs and nascent RNAs) to cell masks, how to extract relevant cell data for Candida albicans morphological classification and prepare the data for statistical analysis. 1. Load the mask and spot data using the provided Jupyter Notebook “04_spot_assignment,” and follow the instructions. 2. Create a dataframe and extract cell parameters from cell masks. During this step, filter the data, and select parameters to extract from cell masks that are useful to describe cells’ morphology. Cell parameters such as cell area, cell centroid, and cell eccentricity can be then correlated to the mRNA spot counts (Fig. 5).

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Fig. 5 Quantifications of C. albicans CLB2 and EFG1 smFISH. (a) Dot plots of CLB2 (blue) and EFG1 (green) number of mRNAs per cell measured for cells grown either in Spider or TSB medium at 37 °C. Plots include expressing cells; black bar represents the mean. (b) Dot plots of CLB2 (blue) and EFG1 (green) number of nascent RNAs per nuclei measured for cells grown either in Spider or TSB medium at 37 °C. Plots include cells with transcription site; black bar represents the mean. (c) Histogram of CLB2 (left) and EFG1 (right) number of mRNAs per cell measured for cells grown either in Spider (blue) or TSB (orange) medium at 37 °C.

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3. Assign mRNAs and nascent RNAs present at transcription sites to individual cells. 4. Count the number of Nuclei per cell. This parameter is particularly interesting for hyphal cells which are multinucleated (Fig. 3). 5. Save the extracted cell data in the CSV format, by executing the Notebook section “Transfer data to dataframe structure and save.” 6. If satisfied with the results, you can proceed with the batch assignment of all the data. 7. Ensure that the data is properly saved for further statistical analysis. 8. smFISH spot data (i.e., mRNA spots per cell, nascent RNAs per transcription site) and cell parameters such as eccentricity (i.e., measure of cell roundness) can be plotted as illustrated in Fig. 5. The results can be plotted using the Jupyter Notebook “05-plotting.”

4

Probe Sequences CLB2-Q670 probe set: smFISH probes mixed in an equimolar ratio and hybridizing to different positions along the CLB2 mRNA (see Note 1) (gttttagtgacttgtggcat tgctgtacttttgatctagt gatggtacttatcgactct tgagaattggatatggtggt gacttgaggtggtgatgaaa actccttcagatgatgttac attgacgtgtattgacatgt ttgatttgaaacatcaccca gttggtttggcatttgttat cacccaatggttttctttta gtttttgtagaggggcattg ttggtctagatggtctatgc gttgttatcactggcaatgg gttactactgctactggtac ctgccaaagatgctagtcta cgtttttggggtaatcgaga tttgtcgacgattcagtagc tctggttgtggtactcttaa ggtatgactgacttctttcc ccaaatcctgccattcataa accattaattggtcgtcact attcgtgtttctaattcgta aaagatattgcggatcaggt tcgatctcattcttggtttt gcatttcaacaagccaatca tatccattacattgactgcc ttgaaccacttcaacagaca agctgctgtagccaataatt cttcatatttggcagcagta gcataatttttaaccagggg cttctggagtatatgaacca gcatgtatttttctgcttgt cattggattggggtaattca agctttagaaattctcctca tattttcctagcgttcttga cataaagatggtctcatacc ttgctaaatacatggccagg tggcaatttgcccaatatta tgaatcaaattcccattcca ctgattctataacctccact attcgatacattctctcata catcatgttctattggagca tctcatggcatattttttga agagtacttgctctcataaa attttttagcccaatttcga acaaatctcttcctgatgct tagcctatgggtcgataatc tactcttctgcttctgctac).

ä Fig. 5 (continued) (d) Histogram of cell eccentricity values measured for cells grown either in Spider (blue) or TSB (orange) medium at 37 °C. (e) Scatterplot with marginal KDEs (y-axis, kernel density estimate) and histplot (y-axis, cell counts) plots of CLB2 (left) and EFG1 (right) number of mRNAs per cell correlated to cell eccentricity (i.e., cells with eccentricity close to 0 are round, while cells with eccentricity close to 1 are elongated) measured for cells grown either in Spider or TSB medium at 37 °C

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EFG1-Q670 probe set: smFISH probes mixed in an equimolar ratio and hybridizing to different positions along the EFG1 mRNA (see Note 1) (gggtatagaatacgttgaca atttccgttcatttgattgt agttgtttgctggggcatac ggaaaagcctgttgattggc gttgctgttggctagcattg ttataaggctgttggactgc gtccttgttgctgatagaac tgtctgtccagtctgttgac gtaatcatattgttgctgct ggcaggatactgatatctgt gttggtaatagtttccttga ggctgtgacaattgattagg atccattgtaatgctgaggt cactaggagcacttgtgtaa tagtggtggctgttgtgaag ggataggtactgcttgttga gttgcattgtcgatacatgt acgtatcctgaacaggagtt cacagtggaagtgctcgagg tggtagttgttactcgtggt ccgacacattattggcatca attattatctgctcttctga cattgagcaatttggttccg cttctaccacgtgtcatttg tttctgatttcaaaatccca tccgattttcacaacgtgtc cagactcctttcaaatgcat gccaatgctctttcaaatgg caatttgttcacgttgagcc cggtttgaatcactcgttta agcagctgcattaggagtta aagcagaagtggcagtggca ccattaccgctagaactttt ccactggtagcagatatact aagcaccagacacattactg actagtggtggaacctgcac agtggcagccttggtattta taattaccttgaggggtacc caggagcattatactgacca atgcaggtgtattttggttc attggttgtagaacctggtt cccatacatttgttgttgtt agtaactactattagcagca gttgatctgactgttgttgt caccacttggtgtagaagtt ttggtgcacagatctagttc gtcaatgactgaacttgggg ggtgaagggtgaactgaacc).

5

Notes 1. smFISH probes can be custom designed using freely available software such as the Stellaris RNA FISH Probe Designer (Biosearch Technologies), or ProbeDealer [40]. Each smFISH probe mix is shipped as a dried set containing 5 nmoles of up to 48 individual smFISH probes that are 20-nucleotides long, mixed in an equimolar ratio. The probes are selected to target the mRNA coding sequence, but if the coding region is not long enough to be targeted by 48 probes, more probes can be added targeting the 5′ or 3′ untranslated regions. By performing a BLAST analysis, eliminate probes that have more than 85% homology with other mRNAs to avoid high background signal in the smFISH channel. Common probes vendors are LGC Biosearch Technologies and Synbio Technologies. 2. For smFISH imaging, we use an Olympus BX-63 epifluorescence microscope equipped with Ultrasonic stage and UPLSAPO 100× 1.4NA oil-immersion objective (Olympus). Lumencore SOLA FISH light source, a Hamamatsu ORCAFusion sCMOS camera (6.5 μm pixel size) mounted using U-CMT C-Mount Adapter, and zero-pixel shift filter sets: F36-500 DAPI HC Brightline Bandpass Filter, F36-502 FITC HC BrightLine Filter, F36-542 Cy3 HC BrightLine Filter, AHF-LED-FISH-R Filter for Cy3.5 and F36-523 Cy5 HC BrightLine Filter. Images are acquired across 61–81 optical

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sections (depending on the sample thickness) with a z-step size of 0.2 μm. The CellSens software (Olympus) is used for instrument control and image acquisition. 3. To obtain filamentous C. albicans cells, it is important to grow cells at low density. Keep the cells growing in exponential phase. At this density, the number of dead cells, which are autofluorescent, is minimal. 4. Excessive fixation will reduce the efficiency of yeast cell wall digestion during the lyticase treatment. 5. Duration of the lyticase treatment varies with fungal strains and treatment. Over-digestion will damage cell morphology and will affect accurate counting of single molecules. Because we observed lyticase batch-to-batch variability, we test each new batch digestion efficiency and adjust the duration of the lyticase treatment accordingly. For phase contrast images of lyticase digestion shown in Fig. 2a, cells were imaged at 30 °C using a Nikon Ti-eclipse widefield fluorescence microscope (Nikon, Minato, Tokio, Japan) equipped with an Andor Zyla 5.5 sCMOS camera and with a Plan Apo λ 100× 1.45 NA oil-immersion Ph3 objective and the Ph3 condenser annulus. 6. Usually 6*106 cells are seeded per coverslip. This provides high cell density during imaging. 7. Seventy percent ethanol allows to store the cells for several months. It also contributes to cell permeabilization, crucial for smFISH. For C. albicans, it is recommended at least an overnight incubation in 70% ethanol at -20 °C. We noticed that with prolonged incubation in 70% ethanol cellular background fluorescence is reduced. 8. At this stage and before starting the hybridization, use a phase contrast microscope to check if the cells are well attached to the coverslip. If cells detach at this step from the coverslips, the coverslip may not have been completely dry after poly-L-lysine treatment. 9. The pre-hybridization and the hybridization solution contain 10% (vol/vol) formamide in 2×SSC. This formamide concentration is optimal for ~20-nt-long probes. If using longer probes, increase the formamide concentration (e.g., for 35 nt probes, use 15–20% formamide in the pre-hybridization and the hybridization solutions). 10. To perform smFISH for two mRNAs simultaneously, add for each coverslip an equal amount of the two probe mixes in 5 μL of the ssDNA/tRNA competitor and lyophilize. Continue the protocol from Subeading 3.3.4. If only one smFISH probe set is used, we recommend the use of probes labeled with Quasar 670/Cy5 fluorophore. If a two color smFISH is performed, we

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recommend the use of probes labeled with Quasar 670/Cy5 and CalFluor610/Cy3.5 dyes. 11. Do not extend the time of this wash; otherwise, the smFISH signal will reduce significantly. 12. While acquiring the Z-stacks for the different channels, image from the longest (CY5) to the shortest (DAPI) wavelength and finish with DIC. Prolonged exposure times can cause photobleaching. To increase the signal of the smFISH, it is better to increase the number of probes specific for each mRNA rather than increasing the exposure time or light source power. 13. Representative images, Jupyter notebooks, and smFISH analysis files are available for download here: https://zenodo.org/ records/10613839.

Acknowledgments We would like to thank Prof. Bastiaan Krom (ACTA, The Netherlands) for providing the C. albicans strain SC5314 and the members of the Systems Biology section for helpful discussion. This work was supported by the Vrije Universiteit and by the Dutch Research Council (NWO) (OCENW.M.21.011) to E.T. Contributions S.O. designed the original protocol, generated scripts, and analyzed the data; M.M. optimized the smFISH protocol and generated the smFISH data. Z.W. generated scripts and analyzed the data; T.D.V. optimized the smFISH protocol and generated scripts for data analysis. P.S. optimized the analysis scripts and supervised the analysis. ET designed the protocol, analyzed the data, wrote the manuscript draft, and supervised the research. All authors read and approved the final version of the manuscript.

References 1. Femino AM, Fay FS, Fogarty K et al (1998) Visualization of single RNA transcripts in situ. Science 280:585–590 2. Zenklusen D, Larson DR, Singer RH (2008) Single-RNA counting reveals alternative modes of gene expression in yeast. Nat Struct Mol Biol 15:1263–1271 3. Raj A, van den Bogaard P, Rifkin SA et al (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5:877–879 4. Gerber A, van Otterdijk S, Bruggeman FJ et al (2023) Understanding spatiotemporal coupling of gene expression using single molecule

RNA imaging technologies. Transcription 0:1– 22 5. Das S, Vera M, Gandin V et al (2021) Intracellular mRNA transport and localized translation. Nat Rev Mol Cell Biol 22:483–504 6. Tutucci E, Livingston NM, Singer RH et al (2018) Imaging mRNA in vivo, from birth to death. Annu Rev Biophys 20(47):85–106 7. Lubeck E, Cai L (2012) Single-cell systems biology by super-resolution imaging and combinatorial labeling. Nat Methods 9:743–748 8. Chen KH, Boettiger AN, Moffitt JR et al (2015) RNA imaging. Spatially resolved,

Detection of Single mRNAs in Candida albicans Cells by smFISH highly multiplexed RNA profiling in single cells. Science 348:6233 9. Eng C-HL, Lawson M, Zhu Q et al (2019) Transcriptome-scale super-resolved imaging in tissues by RNA seqFISH+. Nature 568:235– 239 10. Eliscovich C, Shenoy SM, Singer RH (2017) Imaging mRNA and protein interactions within neurons. Proc Natl Acad Sci U S A 114:E1875–E1884 11. Tutucci E, Singer RH (2020) Simultaneous detection of mRNA and protein in S. cerevisiae by single-molecule FISH and immunofluorescence. Methods Mol Biol 2166:51–69 12. Maekiniemi A, Singer RH, Tutucci E (2020) Single molecule mRNA fluorescent in situ hybridization combined with immunofluorescence in S. cerevisiae: dataset and quantification. Data Brief 30:105511 13. Bayer LV, Batish M, Formel SK et al (2015) Single-molecule RNA in situ hybridization (smFISH) and immunofluorescence (IF) in the drosophila egg chamber. Methods Mol Biol 1328:125–136 14. Zhao L, Fonseca A, Meschichi A et al (2023) Whole-mount smFISH allows combining RNA and protein quantification at cellular and subcellular resolution. Nat Plants 9:1–9 15. Moffitt JR, Lundberg E, Heyn H (2023) The emerging landscape of spatial profiling technologies. Nat Rev Genet 23:741–759 16. Sofroniew N, Lambert T, Evans K, et al (2022) napari: a multi-dimensional image viewer for Python (Internet). Zenodo; cited 2023 Jun 18). Available from: https://zenodo.org/ record/7276432 17. Stringer C, Wang T, Michaelos M et al (2021) Cellpose: a generalist algorithm for cellular segmentation. Nat Methods 18:100–106 18. Pachitariu M, Stringer C (2022) Cellpose 2.0: how to train your own model. Nat Methods 19:1634–1641 19. Schmidt U, Weigert M, Broaddus C, et al (2018) Cell detection with star-convex polygons, p 265–273. Available from: http:// arxiv.org/abs/1806.03535 20. Weigert M, Schmidt U, Haase R, et al (2020) Star-convex polyhedra for 3D object detection and segmentation in microscopy. 2020 IEEE winter conference on applications of computer vision (WACV) (Internet). Snowmass Village, CO, USA: IEEE; p 3655–3662. Availablefrom: https://ieeexplore-ieee-org.vu-nl.idm. oclc.org/document/9093435/ 21. Cutler KJ, Stringer C, Lo TW et al (2022) Omnipose: a high-precision morphology-

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independent solution for bacterial cell segmentation. Nat Methods 19:1438–1448 22. Mueller F, Senecal A, Tantale K et al (2013) FISH-quant: automatic counting of transcripts in 3D FISH images. Nat Methods 10:277–278 23. Tsanov N, Samacoits A, Chouaib R et al (2016) smiFISH and FISH-quant – a flexible single RNA detection approach with super-resolution capability. Nucleic Acids Res 44:e165 24. Imbert A, Ouyang W, Safieddine A et al (2022) FISH-quant v2: a scalable and modular analysis tool for smFISH image analysis. RNA 28:786– 795 25. Perkel JM (2019) Starfish enterprise: finding RNA patterns in single cells. Nature 572:549– 551 26. Rahman S, Zenklusen D (2013) Singlemolecule resolution fluorescent in situ hybridization (smFISH) in the yeast S. cerevisiae. Methods Mol Biol 1042:33–46 27. Patel HP, Brouwer I, Lenstra TL (2021) Optimized protocol for single-molecule RNA FISH to visualize gene expression in S. cerevisiae. STAR Protocols 2:100647 28. Heinrich S, Geissen E-M, Kamenz J et al (2013) Determinants of robustness in spindle assembly checkpoint signalling. Nat Cell Biol 15:1328–1339 29. Bartholomai BM, Gladfelter AS, Loros JJ et al (2021) Quantitative single molecule RNA-FISH and RNase-free cell wall digestion in Neurospora crassa. Fungal Genet Biol 156: 103615 30. Lee C, Roberts SE, Gladfelter AS (2016) Quantitative spatial analysis of transcripts in multinucleate cells using single-molecule FISH. Methods 98:124–133 31. Baumann S, Pohlmann T, Jungbluth M et al (2012) Kinesin-3 and dynein mediate microtubule-dependent co-transport of mRNPs and endosomes. J Cell Sci 125:2740– 2752 32. Moreno-Vela´squez SD, Pe´rez JC (2021) Imaging and quantification of mRNA molecules at single-cell resolution in the human fungal pathogen Candida albicans. mSphere 6:4 33. Elson SL, Noble SM, Solis NV et al (2009) An RNA transport system in Candida albicans regulates hyphal morphology and invasive growth. PLoS Genet 5(9):e1000664 34. Pappas PG, Lionakis MS, Arendrup MC et al (2018) Invasive candidiasis. Nat Rev Dis Primers 4:18026 35. Bongomin F, Gago S, Oladele RO et al (2017) Global and multi-national prevalence of fungal diseases-estimate precision. J Fungi (Basel) 3: E57

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36. Min K, Neiman AM, Konopka JB (2020) Fungal pathogens: shape-shifting invaders. Trends Microbiol 28:922–933 37. Lohse MB, Gulati M, Johnson AD et al (2018) Development and regulation of single- and multi-species Candida albicans biofilms. Nat Rev Microbiol 16:19–31 38. Kadosh D (2019) Regulatory mechanisms controlling morphology and pathogenesis in

Candida albicans. Curr Opin Microbiol 52: 27–34 39. Schindelin J, Arganda-Carreras I, Frise E et al (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9: 676–682 40. Hu M, Yang B, Cheng Y et al (2020) ProbeDealer is a convenient tool for designing probes for highly multiplexed fluorescence in situ hybridization. Sci Rep 10:22031

Chapter 3 RNA and Protein Detection by Single-Molecule Fluorescent in Situ Hybridization (smFISH) Combined with Immunofluorescence in the Budding Yeast S. cerevisiae Anna Maekiniemi and Robert H. Singer Abstract The inherent stochastic processes governing gene expression give rise to heterogeneity across individual cells, highlighting the importance of single-cell studies. The emergence of single-molecule fluorescent in situ hybridization (smFISH) enabled gene expression analysis at the single-cell level while including the spatial dimension through the visualization and quantification of mRNAs in intact fixed cells. By combining smFISH with immunofluorescence (IF), a comprehensive approach takes shape facilitating the study of mRNAs and proteins to correlate gene expression profiles to different cellular states. This chapter serves as a comprehensive guide to a smFISH-IF protocol optimized for gene expression analysis in the budding yeast S. cerevisiae. We utilize smFISH to visualize the mRNA localization pattern of the CLB2 cyclin over the course of the cell cycle inferred by alpha-tubulin IF. Key words smFISH, Immunofluorescence, smFISH-IF, Single-molecule, RNA localization, Gene expression, Cell cycle, Microscopy

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Introduction It is widely recognized that cells possessing identical genetic codes do not manifest identical gene expression profiles, a phenomenon playing a pivotal role in the differentiation of cells and tissues [1]. This heterogeneity arises from inherently stochastic processes involved in mRNA and protein synthesis and degradation [2]. Extensive efforts have been dedicated to studying these processes to enhance our understanding of cellular functions and disease mechanisms. The repertoire of techniques designed to study gene expression is constantly growing and evolving, with RNA sequencing methods swiftly maturing to become a standard approach to investigate how genes behave, from single cells to

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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whole organisms. However, a limitation of these methods is the loss of the spatial dimension as cells are broken up to extract the RNA. Single-molecule fluorescence in situ hybridization (smFISH) conquers this limitation. Emerging in 1998 as the pioneering method to visualize mRNA molecules at single-molecule resolution in intact fixed cells [3], smFISH remains the gold standard for visualizing single mRNAs. This robust technique utilizes multiple dye-labeled DNA oligonucleotides designed to be complementary to the mRNA of interest. Upon hybridization of the probes along the transcript, single mRNA molecules can be detected as diffraction-limited spots using fluorescence microscopy, providing a snapshot of transcriptional activity, mRNA abundance, and their subcellular localization within intact fixed cells [4, 5]. Since its inception, smFISH has been adopted to study gene expression in a wide range of organisms, from single cells to whole tissue samples [6–10]. More often than not, the expression of a gene is studied in the context of various cellular processes or diseases regulated by multiple genes. In such cases, the importance of multiplexing becomes apparent. By designing different sets of smFISH probes targeting multiple genes, each labeled with distinct fluorophores, researchers can simultaneously characterize the mRNA expression profiles of multiple transcripts in a single experiment [11]. Moreover, smFISH can be effectively combined with immunofluorescence (IF). By correlating the expression of one or more mRNAs and proteins in space and time, researchers can study the expression and spatial distribution of a transcript in the context of different cellular states, such as during the cell cycle, by targeting specific protein markers [12–14]. This powerful approach allows for a comprehensive understanding of gene expression dynamics within complex biological contexts and provides valuable insights into the interplay between gene regulation and cellular processes and diseases. This chapter presents our smFISH-IF protocol for studying gene expression in the budding yeast S. cerevisiae. The visualized transcript here is the CLB2 mRNA, the major mitotic cyclin in budding yeast, whose mRNA localization to the bud compartment has recently been shown to play a crucial role in cell cycle progression [14, 15]. To analyze CLB2 gene expression throughout the cell cycle, we combine CLB2 smFISH with IF targeting the microtubule protein alpha-tubulin. Combining visualization of cell morphology, nuclear (DAPI) staining, and the alpha-tubulin IF enables the researcher to infer the approximate cell cycle phase of individual cells, evading the need for synchronization using drugs that might affect other cellular processes [16]. The current protocol is based on previously published protocols with some modifications [13, 17]. The procedure starts on day 1 with the setup of overnight cell cultures and concludes with imaging analysis on day 4 or 5, depending on the extent of the

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Fig. 1 Schematic overview of experimental procedure for smFISH Subheading 3.1–3.2). (a). Cells are incubated at 26 °C overnight under continuous shaking. (b). Coating of coverslips is done fresh for each experiment by adding 0.1% polyL-lysine to acid cleaned coverslips, and incubate at RT for 30–60 minbefore washing and drying. (c). Cell fixation is done in a 4% PFA solution at RT for 45 min under continuous shaking. (d). Cells are digested by the lyticase enzyme in spheroplast buffer at 30 °C for 7–10 min Digestion progression is monitored using a phase contrast microscope. Cells appear darker when the cell wall is digested. (e). Digested and washed cells are plated on poly-L-lysine coated coverslips placed in a 12-well culture plate. Once cells have settled, they are washed and 70% EtOH is added. Cells are stored at -20 °C

experiment. On day 2, approximately 5–6 h are required for fixing, permeabilizing, and plating the cells on coverslips. Day 3 requires approximately 10 h, including 1.5 h for preparing the smFISH probe hybridization, 4 h for hybridization itself, 1.5 h for posthybridization washing, and 3 h for IF. Figures 1, 2, and 3 provides an overview of the primary steps of the protocol.

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2. 0.1 N HCl. 3. Double-distilled ultrapure RNase-free water (DDW). 4. 70% ethanol (EtOH). 5. 0.1% poly-L-lysine. 6. 12-well culture plate.

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Fig. 2 Schematic overview of experimental procedure for smFISH Subheading 3.3) hybridization. (a). Cells stored in -20 °C are washed and pre-hybridized with 10% formamide in 2× SSC for 30 min at RT. (b). 20 μL hybridization buffer prepared during the pre-hybridization incubation is added to parafilm placed in a 15 cm petri dish wrapped in aluminum foil (hybridization chamber) to protect the samples from light. Coverslips are placed on top of the hybridization mix, cells facing down. A DDW filled cap is placed inside the hybridization chamber to maintain a humid environment. (c). The hybridization chamber is closed and sealed using parafilm to avoid evaporation. Allow hybridization to occur during incubation at 37 °C for 3 h

Fig. 3 Schematic overview of experimental procedure – immunofluorescence Subheading 3.4). (a). Extensively wash coverslips after hybridization. Wash twice with preheated pre-hybridization buffer for 15 min, once with permeabilization buffer for 5 minutes, and once with 1× SSC for 10 min. Fix cells with IF fixation buffer. (b). Put coverslips on top of the 1.5 mL tube caps, cell side up, in the IF chamber containing DDW at the bottom. Carefully add (1) blocking buffer and (2) primary and (3) secondary antibodies to the coverslips. Wash extensively after steps 2 and 3. (c). Mount coverslips using DAPI containing mounting media. Let polymerize overnight 2.2

smFISH

1. Synthetic complete (SC) medium: 2 g/L SC powder, 6.7 g/L yeast nitrogen base with ammonium sulfate, 20 g/L glucose. 2. 32% paraformaldehyde (PFA), EM grade. 3. Buffer B: 1.2 M sorbitol and 100 mM KHPO4 pH 7.5. Store at 4 °C. 4. Spheroplast buffer: 1.2 M sorbitol, 100 mM KHPO4 pH 7.5, 20 mM ribonucleoside-vanadyl complex (VRC) (see Note 1), 20 mM β-mercaptoethanol (see Note 2). Prepare fresh. 5. Lyticase buffer: 25,000 U/mL lyticase in 1x PBS to. Aliquot and store at -20 °C. 6. 70% ice-cold EtOH. 7. 20× saline-sodium citrate (SSC). 8. FISH pre-hybridization buffer: 10% formamide, 2× SSC. Prepare fresh.

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9. tRNA/ssDNA mix: 10 mg/mL E. coli tRNA, 10 mg/mL single-stranded salmon testes DNA. Aliquot and store at -20 °C. 10. Dye conjugated DNA probes (see Note 3). 11. Solution F: 10% formamide, 2× SSC, 10 mM NaHPO4 pH 7.5. Prepare fresh. 12. Solution H: 2× SSC, 2 mg/mL bovine serum albumin (BSA), 10 mM VRC. Prepare fresh. 13. Permeabilization buffer: 2× SSC, 0.1% Triton ×-100. 2.3 Immunofluorescence

1. IF fixation buffer: 4% PFA, 1× PBS. 2. 50 mg/mL RNase-free BSA. 3. 1× PBS. 4. Blocking solution: 1× PBS, 0.1% BSA. 5. Primary antibody in blocking solution. Here we use an antialpha-tubulin antibody (1:1000) resuspended in 2 mM Na azide, 1% BSA, and 1×PBS. 6. Secondary antibody in blocking solution. Here we use a goat anti-mouse secondary antibody (1:1500) conjugated to Alexa Fluor 647. 7. 100% EtOH. 8. Mounting solution with DAPI. 9. Nail polish, microscopy grade.

2.4

Equipment

1. 26 °C shaking incubator. 2. 125 mL Erlenmeyer flask. 3. Spectrophotometer at 600 nm. 4. Whatman filter paper. 5. 50 mL falcon tubes and 1.5 mL microcentrifuge tubes. 6. Tabletop shaker at room temperature. 7. Refrigerated centrifuges for 50 mL and 1.5 mL tubes. 8. 30 °C water bath. 9. Phase contrast microscope with a 20× objective. 10. Microscope glass slides. 11. Vacuum aspirator. 12. Vacuum centrifuge with heating (e.g., Vacufuge). 13. Parafilm. 14. Forceps. 15. Hybridization chamber (Fig. 2b) (see Note 4). 16. Three Coplin jars with 1× PBS. 17. IF chamber (Fig. 3b) (see Note 5).

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2.5 Imaging and Analysis

3

1. Wide-field epifluorescence microscope (see Note 6). 2. smFISH analysis software, e.g., FISH quant [18].

Methods

3.1 Preparation of Coverslips

Coverslips are washed with acid to ensure that they are clean and to facilitate cell adherence. 1. Boil 200 coverslips in 500 mL 0.1 N HCl in a 1 L beaker for 30 min. 2. Extensively rinse with water to ensure all acid is rinsed out. 3. Autoclave rinsed coverslips. 4. Store the clean coverslips in 70% EtOH at 4 °C for up to a year.

3.2 Cell Culture Setup, Cell Fixation, and Permeabilization

1. Inoculate cells in 25 mL SC medium and grow overnight in a 26 °C shaking incubator (see Note 7). 2. Dilute cells to an OD600 of 0.1 and grow at 26 °C under shaking until they have reached an OD600 of 0.3–0.4. For strain background BY4741, the growth time corresponds to approximately 3 h (see Note 8). 3. While cells are growing, coat five coverslips per 25 mL cell culture with poly-L-lysine by placing acid-washed coverslips on a clean surface, e.g., a sheet of Whatman filter paper. Aspirate any remaining EtOH and rinse twice with DDW. Aspirate excess water and let the coverslips air-dry. Add ~250 μL 0.1% poly-L-lysine in DDW to each coverslip to cover the whole surface, and incubate at RT for 30–60 min. Aspirate excess poly-L-lysine, and let the remainder air-dry on the coverslip. Wash twice with DDW. Aspirate excess water and let the coverslips air-dry completely. Using forceps, place the coated coverslips in a clean 12-well culture plate, coated side up. Prepare coverslips fresh for each experiment. 4. Prepare 50 mL tubes with 3.15 mL 32% paraformaldehyde (see Note 9). 5. Transfer 21.85 mL cell culture to the PFA-containing tubes. Immediately mix to evenly distribute the PFA. Incubate at RT for 45 min on a tabletop shaker to fix the cells (see Note 10). 6. Collect cells at 2400 g for 3 min at 4 °C. Wash three times with 10 mL cold buffer B. 7. Resuspend cells in 1 mL cold buffer B, and transfer to a 1.5 mL tube. Collect cells at 2400 g and aspirate all buffer B. Prepare the spheroplast buffer during the washes. 8. Resuspend cells in 480 μL spheroplast buffer.

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9. Add lyticase enzyme at 25,000 units per OD of cells (see Note 11). 10. Incubate cells in a 30 °C water bath for 7–10 min to allow for digestion of the cell wall. After 6 min, add 3 μL of digested cells onto a microscope slide, and cover with a coverslip. Check the progress of digestion using a phase contrast microscope with a 20× objective. Digested cells will appear darker than non-digested cells (Fig. 1d) (see Note 12). 11. Once approximately 50% of the cells are digested, collect cells at 2400 g at 4 °C for 3 min (see Note 13). 12. Stop the digestion reaction by washing cells with 1 mL cold buffer B. Collect cells at 2400 g at 4 °C for 3 min. Keep the cells on ice. 13. Resuspend the cells in 1 mL buffer B. Add ~150 μL cells to each poly-L-lysine coated coverslip placed in a 12-well culture plate, and distribute throughout the whole surface to ensure an even cell density on the coverslip. Let cells settle at 4 °C for 1 h. 14. Slowly add 2 mL of buffer B to each well. Aspirate the liquid to remove excess cells. This will leave a monolayer on the coverslips. 15. Slowly add 2 mL ice-cold 70% EtOH to each well. Close the 12-well plate, and wrap with parafilm to avoid evaporation. Incubate at -20 °C overnight. Cells can be stored in 70% EtOH at -20 °C for up to 6 months. 3.3 Probe Hybridization

1. Rehydrate cells by placing coverslips in a new 12-well culture plate. Incubate for 5 min at room temperature in 2 mL 2× SSC. Wash once more with 2× SSC for 5 min. This should remove any detached cells, and wash off the EtOH (see Note 14). 2. Pre-hybridize the cells by adding 2 mL FISH pre-hybridization buffer, and incubate at room temperature for 30 min. During the incubation, prepare the hybridization solution consisting of smFISH probes, E. coli tRNA, ssDNA, solution F, and solution H with a final volume of 25 μL per coverslip (see Note 15). 3. Mix 5 μl of tRNA/ssDNA mix per coverslip with smFISH probe at a final concentration of 125 nM in the final hybridization mix with a volume of 25 μL per coverslip. For example, for five coverslips to hybridize, and a smFISH probe stock solution of 25 μM, add 0.625 μL probe to 25 μL tRNA/ssDNA mix (see Note 15, 16). 4. Dehydrate the probe, tRNA, and ssDNA solution in a vacuum centrifuge with heating, preheated to 45 °C, until all liquid has evaporated, approximately 1 minute per μL probe mix.

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5. Prepare solution F. Boil for 2 min at 95 °C. Let cool in room temperature. 6. Prepare solution H. Make sure the VRC is homogenized by vortexing before use. 7. Add 12.5 μL of solution F and 12.5 μL of solution H per coverslips to the dried probe tRNA/ssDNA mix. Mix by pipetting up and down until the pellet is fully reconstituted. 8. Place a sheet of parafilm in the FISH chamber with the clean side up. Add drops of 20 μL hybridization solution for each coverslip. Make sure to maintain distance between the drops to keep the coverslips separated. Five coverslips can easily fit in one hybridization chamber (see Note 4) (Fig. 2b). 9. Using forceps, place coverslips, cells facing down, onto the hybridization mix. Be careful not to generate any air bubbles in this process. 10. Fill a cap from a 15 mL tube with DDW, and place inside the chamber. Close the chamber, and seal using parafilm. Incubate at 37 °C for 3 h. From this point on, the samples need to be protected from light to avoid bleaching of the fluorophore. 11. Preheat FISH pre-hybridization buffer to 37 °C for washes following hybridization. 12. Using forceps, place the coverslips back into the 12-well culture plate containing the pre-hybridization solution. 13. Aspirate the buffer and wash with 2 mL pre-hybridization solution. Wrap the 12-well culture plate in aluminum foil to protect the samples from light. Incubate at 37 °C for 15 min. 14. Repeat the wash step. 15. Wash once with 2 mL permeabilization buffer for 5 min at room temperature. Be careful not to extend this wash step, to maintain the integrity of the cells. 16. Wash once with 2 mL 1× SSC for 10 min at room temperature. 3.4 Immunofluorescence

1. Fix the coverslips with 2 mL IF fixation buffer for 10 min at room temperature. 2. Wash with 2 mL 1× PBS for 5 min at room temperature. 3. Using forceps, move the coverslips to the IF chamber. Place coverslips with cells facing up on top of the tube caps (Fig. 3b) (see Note 5). 4. Block by carefully adding 190 uL blocking solution to each coverslip. Incubate in closed IF chamber for 30 min at room temperature.

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5. Remove IF buffer and incubate cells at RT with 190 μL primary antibody at recommended dilution in blocking buffer for 45 min. Here, we used a mouse anti-alpha-tubulin antibody at 1:1000 dilution. 6. Fill the Coplin jars with 1× PBS, and wash the coverslips by dipping them into the Joplin jars sequentially. Put coverslips back in the IF chamber, add 190 μL 1× PBS to each coverslip, and incubate in closed chamber at room temperature for 5 min. Repeat this step three to five times. 7. Remove PBS by inverting the coverslips. Incubate cells at RT with 190 μL secondary antibody at recommended dilution in blocking buffer for 45 min. Here, we used a goat anti-mouse antibody conjugated to an Alexa 647 fluorophore at 1:1500 dilution. 8. Repeat the washes in step 6 (see Note 17). 9. Dehydrate the coverslips by dipping each coverslip into 100% EtOH. Let the coverslips dry completely protected from light. 10. Add a drop of mounting solution to a microscope slide and place the coverslip, cell side down, on top of the mounting solution with DAPI (Fig. 3c). Make sure no bubbles form. Allow the mounting solution to polymerize overnight at room temperature, protected from light. 11. Seal the coverslips with nail polish. Ensure the nail polish has completely dried before imaging to avoid damage to the objective. 12. Slides can be stored at 4 °C for several days. For long term storage for multiple months, -20 °C is recommended. Always protect your slides from the light to avoid bleaching of the fluorophores. 3.5 Image Acquisition and Analysis

1. Collect smFISH-IF data using a wide-field fluorescence microscope with a high numerical aperture (see Note 6). 2. Acquire 41 Z-stacks every 200 nm for each stage position to cover the full volume of the cells. Repeat over 10–15 stage position per sample (see Note 18). Use an exposure of 750–1000 ms at 100% light power when imaging the smFISH Q570, the IF is imaged at 300–500 ms with 100% light power, and DAPI is imaged at 50 ms exposure at 12.5% light power. Example images are provided in Fig. 4 where the tubulin IF, cell morphology, and DAPI staining are used to determine the cell cycle phase of individual cells which can be correlated to the expression of the CLB2 mRNA. 3. For single-molecule detection and quantification, we use FISH-quant [13, 18] (see Note 19).

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Fig. 4 Example smFISH-IF images. Top. Schematic of budding yeast cells at different stages of the cell cycle as indicated below schematic. Morphological markers used to identify the cell cycle phase are shown. The nucleus stained with DAPI is shown in blue, microtubules visualized by anti-alpha-tubulin IF is shown in purple/magenta, and approximate size of the bud relative to the mother is depicted. Green dots represent the CLB2 mRNA. Bottom. Example images of budding yeast cells at different cell cycle phases as indicated above each image. Colorway the same as the top schematic. Orange arrow indicates transcription site. Scale bar is 3 μm

4 Notes 1. Activate VRC at 65 °C for 10 min before use. Store in -20 °C. 2. β-Mercaptoethanol is toxic. Work in chemical fume hood while handling this substance. 3. We ordered CLB2 smFISH probe sequences [14] from LGC Biosearch Technologies. The probe mix is reconstituted in TE buffer to a final concentration of 25 μM. Aliquot and store protected from light long term in -80 °C and short term in 20 °C. Probe design against RNA of interest can be done using the LGC Biosearch Technologies Stellaris Probe Designer. gcttcttggaaggaaatagc

tctattgggttggacatcta

taaggccaacctttgtacat

agagagagagggggtaatca

attctgtgagttttctgtgt

gctggattattcgcattact

ccttctttactgagtttagt

gtttcactttcggtatttct

cggattgggataatttaggt

aattgaggaatccgacttcc

gggcagttcttgttcaacaa

tatcgtagtcatctgctttc

gatcttgatacgctttctct

ccattttcatcgcacacttt

atgagggtagtatcccaata

cattttctttcagggttctt

ttcccattcatattcttctt

cataaacatcgcagctgctg

ttgtttttggggagttggat

ggatcatttacatcttctgc

cctttacctaacatttttct

ggtcttggttttctttatca

atcattgacgtactcgctga

tattagatttccatcccatt

gcaccaaattgctgactact

taatgacctctagttggtgg

agttcttctttagtataccc (continued)

RNA and Protein Detection by smFISH-IF in S. cerevisiae

55

cccctcctttatagaagtta

ctccggtaataagccgaatt

aagcccattggacggaaatt

tatatttgcaggcagctcag

cctgtccattatgttaatgg

catagccgttttttctaacc

gctgaggaggattcttgtaa

atgatgtgccaaccaattgt

ttctcattcatgcaaggtca

gttatggactacctcagtgt

acctcttcatatttagaggc

gcagccatacattttatatg

gaagtttttcgttttctacc

gtctgtctctgatgcgaaat

aggcgctaaaattgggtgtg

tgaagttgacttctagctgg

aaatgaatttctccccttct

tggtgtggtgttattagtga

4. The hybridization chamber is made from a 15 cm petri dish wrapped in aluminum foil to protect the samples from light. A cap from a 15 mL tube filled with DDW is placed inside the dish to maintain a humid environment. 5. The IF chamber is made from a 15 cm petri dish wrapped in aluminum foil. Caps from 1.5 mL tubes are glued upside down on the bottom of the petri dish (for illustration, see Fig. 3b). DDW is added to the bottom of the petri dish to maintain a humid environment. 6. The microscope used to acquire images in this experiment was an Olympus BX-63 wide-field fluorescence microscope equipped with a CCD camera (Hamamatsu, pixel size 6.45 μm) and a 100× objective with 1.35 numerical aperture. An X-Cite 120 PC EXFO lamp was used as light source. Shift filters used were DAPI-5060C-Zero, Cy3-4040C-Zero, and Cy5–4040-Zero from Semrock. The microscope was controlled using the Metamorph software from Molecular Devices. 7. Do not forget to include controls. Control can include but are not limited to (a) a positive control using a previously successful smFISH probe set against a gene other than your gene of interest labeled with a different dye as an internal control for the smFISH assay and (b) a negative control strain that is not expected to express the gene of interest, e.g., a gene knockout strain. This will allow you to identify potential unspecific binding and background signal from smFISH probes and antibodies. 8. Do not grow the cultures denser. Harvesting the cells at log phase is important to minimize autofluorescence. Additionally, optimal cell density facilitates cell segmentation during image analysis, which can be labor-intensive. 9. Beware of toxic fumes when working with PFA. Work in a chemical fume hood. 10. Over-fixation of the cells might increase autofluorescence and reduce the efficiency of cell wall digestion.

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11. When working with multiple samples, ensure equal digestion across samples by pipetting the lyticase enzyme onto the side of each tube (not into the liquid). Close all tubes, and invert to mix the enzyme into the cell solution. 12. Digestion times may vary depending on strain and how freshly prepared the lyticase is. The efficiency between different batches of lyticase might also vary. It is therefore important to carefully check the progression of digestion frequently. Over-digested cells stand the risk of digesting the cell membrane in addition to the cell wall, compromising the integrity and morphology of the cells and thereby also accurate analysis of mRNA and protein distribution. The digestion step is crucial for the cells to adhere to the poly-L-lysin coated coverslips and for the probe and antibodies to permeabilize. 13. After lyticase digestion, the cells will be fragile. Make sure not to centrifuge the cells at a higher g and never vortex, as this will increase the risk of cell lysis. 14. This is a good point to check if the cells are attached to the coverslip using a phase contrast microscope. If the cells have detached and you have no cells/very few cells on your coverslips, the acid washing might not have been done properly to allow the poly-L-lysine to adhere to the glass or the poly-Llysine might not have dried completely onto the coverslip. Another explanation could be that the cells are under- or over-digested. 15. If you are doing smFISH against multiple genes, simply add all probe sets at equimolar concentration (125 mM) to the hybridization buffer. 16. Example of a hybridization mix done for five coverslips using one probe set can be found in Table 1. 17. If you are using antibodies against multiple proteins, repeat steps 5–8. 18. The number of stage positions will depend on how densely the cells are seeded on the coverslip. We use 500–1000 cells for smFISH quantification. Adjust the number of stage positions accordingly. 19. FISH-quant [18] has a detailed manual, tutorials, and accompanying YouTube video tutorials (https://www.youtube.com/ @muellerfresearch/videos) to guide the user through the analysis process step by step. We recommend using the CellProfiler software [19] for cell segmentation which can be imported for analysis in FISH-quant.

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Table 1 Example hybridization mix for five coverslips Solution

Stock

Volume 1× (μL) Volume 5× (μL)

E. coli tRNA/ssDNA

10 mg/mL

5

25

smFISH probe

25 μM

0.125

0.625

Formamide

100%

2.5

12.5

NaHPO4, pH 7.5

200 mM

0.625

3.125

9.375

46.875

tRNA/ssDNA/probe

Solution F

DDW Solution H SSC

20×

2.5

12.5

BSA

20 mg/mL

1.25

6.25

VRC

100 mM

1.25

6.25

7.5

37.5

DDW

References 1. Bruggeman FJ, Teusink B (2018) Living with noise: on the propagation of noise from molecules to phenotype and fitness. Curr Opin Syst Biol 8:144–150 2. Raj A, Peskin CS, Tranchina D, Vargas DY, Tyagi S (2006) Stochastic mRNA synthesis in mammalian cells. PLoS Biol 4:e309 3. Femino AM, Fay FS, Fogarty K, Singer RH (1998) Visualization of single RNA transcripts in situ. Science 280:585–590 4. Raj A, van den Bogaard P, Rifkin SA, van Oudenaarden A, Tyagi S (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5:877–879 5. Zenklusen D, Larson DR, Singer RH (2008) Single-RNA counting reveals alternative modes of gene expression in yeast. Nat Struct Mol Biol 15:1263–1271 6. Tutucci E, Livingston NM, Singer RH, Wu B (2018) Imaging mRNA in vivo, from birth to death. Annu Rev Biophys 47:85–106 7. Das S, Vera M, Gandin V, Singer RH, Tutucci E (2021) Intracellular mRNA transport and localized translation. Nat Rev Mol Cell Biol 22:483–504

8. Pichon X, Lagha M, Mueller F, Bertrand E (2018) A growing toolbox to image gene expression in single cells: sensitive approaches for demanding challenges. Mol Cell 71:468– 480 9. Duncan S, Olsson TSG, Hartley M, Dean C, Rosa S (2016) A method for detecting single mRNA molecules in Arabidopsis thaliana. Plant Methods 12:13 10. Long X, Colonell J, Wong AM, Singer RH, Lionnet T (2017) Quantitative mRNA imaging throughout the entire drosophila brain. Nat Methods 14:703–706 11. Levsky JM, Shenoy SM, Pezo RC, Singer RH (2002) Single-cell gene expression profiling. Science 297:836–840 12. Eliscovich C, Shenoy SM, Singer RH (2017) Imaging mRNA and protein interactions within neurons. Proc Natl Acad Sci 114: E1875–E1884 13. Maekiniemi A, Singer R, Tutucci E (2020) Single molecule mRNA fluorescent in situ hybridization combined with immunofluorescence in S. Cerevisiae: dataset and quantification. Data Brief 30:105511

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14. Tutucci E, Maekiniemi A, Snoep JL, Seiler M, Rossum K, van Niekerk DD, Savakis P, Zarnack K, Singer RH (2022) Cyclin CLB2 mRNA localization determines efficient protein synthesis to orchestrate bud growth and cell cycle progression. bioRxiv 2022.03.01.481833. https://doi.org/10. 1101/2022.03.01.481833 15. Trcek T, Larson DR, Moldo´n A, Query CC, Singer RH (2011) Single-molecule mRNA decay measurements reveal promoter- regulated mRNA stability in yeast. Cell 147:1484– 1497 16. Pereira G, Schiebel E (2002) The role of the yeast spindle pole body and the mammalian centrosome in regulating late mitotic event. Curr Opin Cell Biol 13:762–769

17. Tutucci E, Singer RH (2020) Simultaneous detection of mRNA and protein in S. cerevisiae by single-molecule FISH and immunofluorescence. In: Heinlein M (ed) RNA tagging: methods and protocols. Springer, New York, pp 51–69 18. Mueller F, Senecal A, Tantale K, Marie-NellyH, Ly N, Collin O, Basyuk E, Bertrand E, Darzacq X, Zimmer C (2013) FISH-quant: automatic counting of transcripts in 3D FISH images. Nat Methods 10:277–278 19. Kamentsky L, Jones TR, Fraser A, Bray M-A, Logan DJ, Madden KL, Ljosa V, Rueden C, Eliceiri KW, Carpenter AE (2011) Improved structure, function and compatibility for CellProfiler: modular high-throughput image analysis software. Bioinforma Oxf Engl 27:1179– 1180

Chapter 4 Fluorescence In Situ Hybridization as a Tool for Studying the Specification and Differentiation of Cell Types in Nematostella vectensis Oce´ane Tournie`re, Henriette Busengdal, James M. Gahan, and Fabian Rentzsch Abstract The sea anemone Nematostella vectensis is a genetically tractable cnidarian species that has become a model organism for studying the evolution of developmental processes and genome regulation, resilience to fluctuations in environmental conditions, and the response to pollutants. Gene expression analyses are central to many of these studies, and in situ hybridization has been an important method for obtaining spatial information, in particular during embryonic development. Like other cnidarians, Nematostella embryos are of comparably low morphological complexity, but they possess many cell types that are dispersed throughout the tissue and originate from broad and overlapping areas. These features have made two-color fluorescence in situ hybridization an important method to determine potential co-expression of genes and to generate hypotheses for their functions in cell fate specification. We here share protocols for single and double fluorescence in situ hybridization in Nematostella and for the combination of fluorescence in situ hybridization and immunofluorescence. Key words Cnidaria, Gene expression, Neurogenesis, Evolution of development, Cell type specification

1

Introduction Animals and other multicellular organisms are composed of morphologically, physiologically, and molecularly diverse cell types. In bilaterians, specific functions of the organism are often executed by organs, in which specific cell types are arranged in a spatially defined manner to facilitate their interaction. During embryonic development, the cell types constituting specific organs often arise from territories that have been delimited by combinatorial activities of signaling molecules and transcription factors. Migration of cells from other regions of the embryo into the forming organs can further contribute to their full cell type complement, for example,

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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when neural crest-derived progenitors of neurons colonize the intestine in mammals [1, 2]. The presence of functionally specialized organs is common in bilaterians, but rare in non-bilaterian animals, which often show a broad distribution of specific cell types that potentially reflects an ancestral state for extant animals. An informative group of animals for studying cell type development in the absence of organ-level tissue organization are the cnidarians. They are the sister taxon to bilaterians, with an estimated time of divergence of >550 million years ago [3, 4]. Comparing cell type development and embryonic patterning in cnidarians and bilaterians might provide insight into the evolution of organ development and into the mechanisms that regulate proportionality of cell types in different organismal contexts [5, 6]. Cnidarians include jellyfish, corals and sea anemones, and both solitary and colonial species. They can be divided into three major clades, anthozoans, medusozoans, and the parasitic endocnidozoans [7, 8]. A sessile polyp is found in the life cycle of both anthozoans and medusozoans, whereas a free-swimming medusae stage is only found in medusozoans. The body of polyps resembles a hollow cylinder with only one opening that serves for the intake of food and the release of feces. This opening is called the mouth, and the site at which it is located is called the head. The mouth is surrounded by tentacles that are used for catching prey, and the gastric cavity extends to the aboral end of the polyp (or foot). The tissue of cnidarians derives from only two germ layers, which are traditionally called ectoderm and endoderm. In at least some cases, they possess multi- or even pluripotent adult stem cells that maintain tissue homeostasis and contribute to the remarkable regenerative capacity of both polyps and medusae [9, 10]. In the last two decades, several cnidarian species have been developed into model organisms amenable to genetic manipulation like transgene insertion, genome editing, and gene knockdown [11–14]. Among anthozoan cnidarians, the currently most prominent model species for studying embryonic development is the sea anemone Nematostella vectensis. Nematostella can easily be maintained in laboratory culture, large numbers of gametes can be obtained year-round, and the generation time of 3–4 months is compatible with genetic approaches [11, 15, 16]. Embryonic development includes a hollow blastula stage, gastrulation by invagination from the oral pole, and a free-swimming planula larva with a prominent apical tuft of long cilia. Apart from the oral opening and the apical tuft, there are no readily visible morphological landmarks at these stages (Fig. 1a, b). The planula larva gradually develops into a primary polyp that settles and starts feeding with initially four tentacles [17–19]. Despite their relatively simple body plan, Nematostella (as other cnidarians) has a rich repertoire of cell types. Single cell sequencing data suggest over 30 neural cell types and over 20 different types of gland/secretory cells in adult polyps [20, 21]. Many (potentially

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Fig. 1 Limited morphological complexity and variation in gene expression in Nematostella embryos. (a–d) Colorimetric in situ hybridizations of lateral views oriented with the oral pole to the right. Developmental stages are indicated on the top, gene IDs for the probes on the left. (a, b) illustrate the rather few features visible after colorimetric in situ hybridization. No signal is visible at gastrula stage, scattered cells in the aboral ectoderm at mid-planula stage. Note that the apical tuft at the aboral pole is not visible in (b), though it often is still present after in situ hybridization. (c, d) show embryos at the same developmental stage. Despite similar overall distribution and number of labeled cells, their positions vary between specimens. Ecto – ectoderm; endo – endoderm; gc – gastric cavity; pha – pharynx. Scale bars: 100 μm

all) of the neural and gland/secretory cell types can be traced back to a population of embryonic progenitor cells characterized by the expression of NvSoxC and NvSoxB2a [21–23], but the genetic control of the development of individual cell types remains largely unexplored. Cells expressing these genes are broadly distributed across the ectoderm and endoderm at gastrula and planula stages when the embryos already consist of >5000 cells [21, 24, 25]. Genes that identify subpopulations of cells derived from these progenitors can either be distributed in a similarly broad manner [26, 27] or be spatially more restricted, for example, being more abundant in some area along the oral-aboral axis [28, 29]. The number and distribution of cells expressing a particular gene can vary substantially at any given developmental stage

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(Fig. 1c, d). Cell types expressing specific genes thus do not have defined positions, and the embryos have only few morphological landmarks for orientation. Together, this prevents inferring co-expression of genes by comparing colorimetric in situ hybridizations or single-color fluorescence in situ hybridizations (FISH) in different specimens, which often reveal superficially similar expression patterns of scattered cells (Fig. 2). Double fluorescence in situ hybridization (DFISH) and FISH in the background of transgenic reporter lines have therefore become important methods for

Fig. 2 Cells of the neural/secretory lineage are distributed broadly in Nematostella embryos. (a, b) Double fluorescence in situ hybridizations at blastula stage with the probes indicated on top. Though NvSoxB(2) and NvRFamide expression are similar (A″, B″), only NvRFamide shows co-expression with NvPOU4 (A, B). Note that the lack of co-expression of NvPOU4 and NvSoxB(2) is due to sequential expression in cells that first express NvSoxB(2) [27]. (c) Fluorescence in situ hybridization for NvPOU4 combined with IF for the cnidocyst protein NvNCol3. The oral pole is oriented to the top. Scale bars: 50 μm

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studying gene expression in Nematostella with the aim to generate hypotheses about gene-regulatory interactions. While DFISH provides direct information about co-expression at the transcript level, it cannot distinguish between scenarios in which two genes are either expressed in different populations of cells or are expressed sequentially in the same cells. If a transgenic reporter line is available for one of the genes, FISH can reveal whether a second gene is expressed in cells that have previously expressed the reporter transgene. The growing number of transgenic reporter lines will likely increase the applicability of this approach in the future. We here describe protocols for FISH, DFISH, and FISH in combination with immunofluorescence (IF) in Nematostella as tools for studying gene expression during embryonic development. Together with increasingly sophisticated functional approaches, single-cell RNA sequencing and mapping of chromatin features, these methods can be expected to provide an improved understanding of the gene-regulatory networks that govern the specification and differentiation of cell types in Nematostella and how these regulatory programs may have changed in the evolutionary transition to organ-level tissue organization. Despite the additional information that can be obtained by FISH, we note that in our hands colorimetric in situ hybridizations are more sensitive and that we therefore typically first characterize the expression pattern of a given gene by colorimetric in situ hybridization. For the generation of transgenic reporter lines, we refer the reader to previously published protocols [30, 31].

2

Materials Prepare all buffers and reagents using autoclaved Milli-Q (MQ) water unless otherwise noted. Detergents cannot be autoclaved and therefore should be added to autoclaved buffers just prior to use. To reduce the amount of plastic waste, we use the same tube for preparing solutions from the fixation step to the hybridization step. For all other steps, it is important to use clean and RNasefree tubes. Unless otherwise noted, the buffers used are prepared and kept at room temperature (20–25 °C).

2.1 Embryo Preparation

• Nematostella medium: filtered artificial or natural sea water (diluted to 14–16 ppt). • De-jellifying solution: 3% cysteine (l-cysteine) in Nematostella medium, pH adjusted to 7.4–7.6 with 1 M sodium hydroxide (NaOH). Dissolve the l-cysteine crystals in Nematostella medium in a tube (e.g., 50 mL) by inverting or shaking, prior to pH adjustment.

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• Plastic transfer pipettes or Pasteur pipettes with smooth opening (see Note 1). • Petri dishes (see Note 1). 2.2

Fixation

• Phosphate buffered saline (PBS 10×): 18.2 mM NaH2PO4 × H2O, 84.1 mM Na2HPO4 × 2H2O, 1.75 M NaCl. • PTW: 1 × PBS (autoclaved), 0.1% Tween20 from a 25% Tween20 stock. • Magnesium chloride (immobilization agent): 1 M MgCl2 (dissolved in autoclaved MQ water). • Fixative 1: 3.7% formaldehyde and 0.25% glutaraldehyde in Nematostella medium, cooled to 4 °C. Prepared on the day shortly before use. • Fixative 2: 3.7% formaldehyde in PTW, cooled to 4 °C. Prepared on the day, can be stored in a fridge, or kept on ice up until use. • Eppendorf tubes (RNase-/DNase-free 1.5 ml and 2 ml) (see Note 1). • Tube rotator.

2.3

Probe Synthesis

Using Ambion RNAqueous kit, extract the total RNA of embryos (the developmental stages should include the stage when your gene of interest is expressed), and then synthesize the cDNA using Super Script III Reverse Transcriptase. If the timing of expression of your gene of interest is unknown, we recommend to extract the total RNA of embryos at various developmental stages separately, synthesize cDNAs for each stage, and then mix the cDNAs for PCR amplification. Design primers for your gene of interest. In our hands, probes of approximately 1 kb work best. Since the prediction for UTRs are often unreliable for genes annotated in the Nematostella genome, we typically amplified fragments of the coding sequence. Using the cDNA previously synthesized as well as your designed primers, amplify your fragment of interest by PCR. The PCR product can be cloned into a vector (we use pGEM-T Easy vector that contains promoters for SP6 and T7 RNA polymerases) for amplification and sequencing. Using the M13 primers present on the vector, it is then possible to amplify and generate the probe template by PCR. Labeled antisense riboprobes are finally synthesized from the probe templates using T7 (or SP6) RNA polymerase, digoxigenin (DIG), fluorescein (FITC), or dinitrophenol (DNP) modified nucleotide mix. We use Invitrogen MegaScript kits according to the manufacturer’s instructions. The remaining DNA is then removed using TURBO DNase for 30 min at 37 °C. Using a 50% LiCl wash overnight, the synthesized RNA is precipitated and can be spun down into a pellet at the bottom of the tube. The pellet of RNA is washed in 75% ice-cold

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ethanol, before being air-dried and dissolved in RNase-free water. = Stocks of the probes are kept at -20 °C in 50% deionized formamide in RNase-free water. 2.4 FISH Buffers and Reagents

• PTW: 1× PBS (autoclaved), 0.1% Tween20. • PBTx: 1× PBS (autoclaved), 0.1% Triton ×-100. • Endogenous peroxidase inactivation reagent: 3% hydrogen peroxide (H2O2) in methanol. Prepared fresh on the day of use. • Proteinase K solution: 10 μg/ml proteinase K in PTW. Prepare fresh. • Proteinase K stop solution: 4 mg/ml glycine in PTW. Prepare fresh. • Basket rack and basket wash solution: 1 M NaOH. • TEA buffer: 1% triethanolamine (TEA) (≥99.0% (GC)) in PTW. Prepare fresh on the day of use. • 0.25% acetic anhydride solution: 0.25% acetic anhydride in TEA buffer. See Note 2. • 0.5% acetic anhydride solution: 0.5% acetic anhydride in TEA buffer. See Note 2. • 20× saline-sodium citrate buffer (20× SSC): 3.0 M sodium chloride, 0.3 M sodium citrate, pH 7.0. • Hybridization buffer (HB): 50% formamide, 5× SSC, 1% SDS, 50 μg/ml heparin, 100 μg/ml salmon sperm DNA, 9.25 mM citric acid, 0.1% Tween20. • Tris-NaCl-Tween buffer (TNTw): 0.1 M Tris-HCl pH 7.5, 0.15 M NaCl, 0.1% Tween20. • Tris-NaCl-Triton ×-100 buffer (TNT×): 0.1 M Tris-HCl pH 7.5, 0.15 M NaCl, 0.2% Triton ×-100. • TNBlock: 0.1 M Tris-HCl pH 7.5, 0.15 M NaCl, 0.5% blocking reagent (Akoya Biosciences). In an autoclaved 200 ml bottle, mix 0.5 g blocking reagent and 100 ml TN buffer. Heat up gradually to 55 °C in a water bath; swirl the bottle gently until the blocking reagent is completely dissolved. Aliquot and store at -20 °C. • Anti-digoxigenin antibody conjugated with horseradish peroxidase (POD) (Roche), anti-fluorescein-POD (Roche), or antiDNP HRP (Perkin Elmer). • Fluorescence signal detection kits: TSA Plus Fluorescein/TSA Plus Cy3 (Akoya Biosciences) (see Note 3). • StopPOD1: 0.1 M glycine, 0.1% Tween-20, pH adjusted to 2.0 with drops of fuming HCl. • StopPOD2: 3% hydrogen peroxide in PTW. • ImmunoBlock: 5% normal goat serum in PTW.

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• Hoechst (1:100) or DAPI (1:100) nuclear stain in PBTx. • ProLong™ Gold Antifade Mountant with or without DNA Stain DAPI. • Primary antibody and secondary antibody (see Note 15). 2.5

FISH Equipment

• Fume hood: all the steps with formaldehyde, methanol, or formamide should be performed in a fume hood. • FISH baskets (Fig. 3a): the baskets used in this protocol are homemade and were first generated by the Technau Lab. Using a warmed scalpel (warmed up by the flame of a Bunsen burner), cut the tip off of a 1 ml pipette tip. Press the cut tip-end against a piece of mesh (70 μm pore size) lying on a hotplate covered with aluminum foil. When the mesh is attached by the melted plastic, remove the basket from the plate and trim the edges of the mesh with scissors or a scalpel. Ensure the mesh is fully attached before use. It is important to ensure the cut of the basket is straight to avoid embryos sliding and accumulating on one side of the basket during the protocol. The exact size of the baskets depends on the pipette tips used for making them. In our case, the diameter at the bottom is approx. 7 mm and the height ca 20 mm. At this size, each basket can be used for about 100 specimens (blastula to planula stage). • Basket rack box (Fig. 3b): an empty 1 ml pipette tip rack, typically the rack which the pipette tips came in and a box to place the tip rack in (at least two boxes will be needed for

Fig. 3 Baskets, racks, and mounting procedure for in situ hybridizations. (a) Baskets are generated by cutting 1 ml pipette tips and gluing them to a mesh. (b) Insets and lids of tip racks can be used for processing up to 96 samples. (c) For the hybridization step, the baskets are transferred into a standard tube rack. (d) Scotch tape is used as spacer, (e) samples in mounting medium are placed on the slide, (f) mounting medium is applied to the corners of the coverslip, and (g) the coverslip is placed on the sample

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alternating washes) (see Note 4). Up to 96 samples can be processed in a rack; the volume of solutions used in these steps is 25–30 ml. • Eppendorf tube rack. • Eppendorf tubes (1.5 and 2 ml). • Water bath in a fume hood for the hybridization and posthybridization washes. • Tweezers with pointed tips for transfer of baskets. • Plastic and aluminum foil to cover the Eppendorf tube rack.

3

Methods Subheading 3.1, steps 1–5 are derived from protocols originally used for colorimetric in situ hybridizations in Nematostella [32, 33].

3.1 Embryo Preparation

1. Induce spawning by exposing male and female polyps’ boxes to bright light at 24 °C for 12 h [34]. 2. The following day, discard egg packages that were laid overnight, and collect freshly laid egg packages after 2 h. To fertilize, incubate these for 20 minutes in medium collected from the box containing male polyps. 3. Collect fertilized egg packages in a petri dish, and proceed to de-jellify them. Transfer the egg packages to a petri dish containing 30 mL de-jellifying solution. Incubate the egg packages in this solution for 15–20 min on a horizontal shaker with intermittent mixing. 4. Collect the fertilized eggs in the center of the dish by swirling the dish, and transfer the eggs into a new petri dish with minimum de-jellifying solution carryover. Wash them by transfer into a new petri dish containing fresh Nematostella medium at least six times. 5. Put the de-jellified, fertilized eggs into a 21 °C incubator to grow until the desired developmental stage.

3.2

Fixation

1. Transfer embryos to a 1.5 ml Eppendorf tube (see Note 5), wait for them to collect at the bottom, remove the Nematostella medium, and add Fixative 1. Incubate tubes on ice for 90 s, while the embryos sink back to the bottom. 2. Replace the solution with Fixative 2, and incubate for 1 h at 4 °C. Use a tube rotator to rock the samples during this step. 3. After 1 h, replace the fixation solution with PTW. Wait for embryos to settle to the bottom, and repeat three more times

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in PTW, then one time in 1× PBS, and one time in water before incubating them for 5 min in 50% methanol and then 100% methanol. 4. Fixed embryos can be stored in 100% methanol (MeOH) at -20 °C until the start of the protocol. 3.3 Rehydration, Proteinase K Treatment, and Refixation

All washes using the basket rack are performed on a horizontal shaker at low speed unless otherwise stated. Before starting this protocol, wash all the equipment (baskets, tube/basket racks) with 1 M sodium hydroxide for at least 20 min (to degrade any remaining RNAs from previous experiments and prevent RNase contamination), and then wash them thoroughly with autoclaved MQ H2O. Between washes when using the basket rack, the easiest workflow is to use two boxes for alternating transfer of the rack/ baskets between the wash steps. 1. Allow embryos in methanol to reach room temperature, remove methanol, and add 3% hydrogen peroxide in methanol. Incubate for 20 min to quench endogenous hydrogen peroxidase activity. 2. Rehydrate the samples via a series of three washes of 5 min each in MeOH/PTW (diluted to 75/25, 50/50, 25/75). Wash three times for 5 min each with PTW before transferring the samples into the FISH baskets (see Note 6). 3. Incubate the rack with baskets in proteinase K solution for 5 min at room temperature without shaking. Samples should be incubated without shaking until the refixation step (Step 6) (see Note 7). 4. Proceed directly to a wash in proteinase K stop solution for 5 min. Repeat this step one more time. 5. Transfer the basket rack into TEA buffer for 5 min, followed by incubation with 0.25% acetic anhydride solution for 5 min and then another 5 min of incubation with 0.5% acetic anhydride solution. 6. Next wash the samples three times for 5 min in PTW followed by a 30-min incubation in Fixative 2. 7. Wash the samples three times in PTW to remove all the fixative.

3.4 Pre-hybridization and Hybridization

1. Transfer the basket rack into warm hybridization buffer solution (preheated to 60 °C). Put the lid on and place the tray in the water bath for at least 2 h at 60 °C for pre-hybridization. Pre-hybridization can be prolonged to overnight to make the protocol more convenient. To ensure stability in the water bath, put a small weight on top of the basket rack. 2. During this time, add the probes to the hybridization buffer in a 1.5 mL tube (digoxigenin, fluorescein, or DNP-labeled

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riboprobes) at a final concentration of 0.1–1 ng/μl. Preheat the probe/hybridization mix at 85–95 °C for 5 min. 3. Hybridization is performed in an Eppendorf tube rack (Fig. 3c). First, add the probes to the wells (300 μl per well), and then transfer the basket into it using tweezers (see Note 8). Wrap the tube rack first in plastic foil (to avoid probe evaporation), then in aluminum foil (to maintain a homogeneous temperature), and finally in a plastic bag (to avoid water drops entering the rack), and put it back into the water bath with a weight onto it for at least 60 h at 60 °C (see Note 9). 4. Remove the rack from the water bath. Using tweezers, transfer each basket from the tube rack back to the basket rack (see Note 10). 3.5 Posthybridization Washes

1. Excess and unbound probe is removed via a series of four washes of HB/2x SSC solutions (diluted to 75/25, 50/50, 25/75, 0/100 (v/v)) of 30 min each at 60 °C. This is followed by 20 min in 0.2× SSC and finally two washes of 20 min each in 0.1× SSC. 2. This is followed by a series of 10-min washes at room temperature in 0.1× SSC/PTW solutions [diluted to 75/25, 50/50, 25/75, 0/100 (v/v)] on a horizontal shaker at low speed. 3. Samples are then washed four times for 5 min in TNTw on a horizontal shaker at low speed.

3.6 First Probe Labeling and Washes (All Steps Are Performed on a Horizontal Shaker at Low Speed Unless Otherwise Stated)

Depending on the number of baskets to process, either pipette the embryos out of the basket into Eppendorf tubes using a 1 ml pipette (few baskets), or transfer the baskets to a 48-well dish (many samples) for the steps of blocking and antibody incubation. For the TSA reaction, we perform the reaction in tubes to limit use of reagents. 1. Samples are incubated in 500 μl TNblock for 1 h at room temperature. 2. They are then incubated overnight at 4 °C with the antibody (anti-digoxigenin-POD (1:100) or anti-fluorescein-POD (1: 250) or anti-DNP-POD (1:250) (see Note 11) in blocking solution without shaking. 3. On the next day, unbound antibody is removed with a series of ten washes of 15 min each in TNTx. At this step, transfer the samples back to the baskets for more convenient handling of multiple samples. Or carry on in tubes/alternating transfers in 48-well dishes. 4. Samples are then transferred into 2 ml tubes (to limit the quantity of kit reagents used) and incubated in 100–150 μl of fluorophore tyramide amplification reagent for 30–40 min

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(fluorophore diluted 1:50 in 1× working solution) (see Note 12). 5. After the first TSA reaction, it is important to keep samples protected from light exposure until the end of the protocol. 6. Samples are washed two times for 5 minutes in TNT× (for double fluorescence in situ hybridization, proceed directly to Step 3.7/for immunostaining, proceed directly to Step 3.8). If only one probe was used, proceed to nuclei staining by incubating your samples in Hoechst (1:100) in PBT× for 30 min, followed by two washes of 10 min in PBT× and one wash in PBS. 7. Remove the PBS and add 50–100 μl of ProLong™ Gold Antifade Mountant with or without DNA Stain DAPI (this increases the DNA signal without any additional steps in the protocol). Place the tubes with samples in dark at 4 °C until they settle at the bottom (1 day usually but can be kept longer). Once settled, it is possible to pipette them out and mount them on a slide. 8. To mount them on a slide, it is recommended to use two layers of tape on each side of the slide (Fig. 3d). Cut off the tip of a 200 μl tip, pipette the embryos out of the tube, and drop them between the two layers of tapes (Fig. 3e). Using the same tip, add four drops of mounting reagent at each corner of a coverslip (18 × 18mm; Fig. 3f). Drop the cover slip onto the tape (Fig. 3g). This mounting strategy prevents squashing the embryos and helps having an even thickness of the mounting medium. Leave the mounted slide at room temperature at dark overnight to dry. Then store the slides at 4 °C. 9. Samples can now be imaged on either a compound microscope with a camera or on a confocal microscope. 3.7 Double Fluorescence in Situ Hybridization (All Steps Are Performed on a Horizontal Shaker Unless Otherwise Stated)

1. For double labeling, following the TNT× washes, samples are transferred back into baskets (see Note 13) and further washed in StopPOD1 for 10 min to block the peroxidase activity (see Note 14). 2. Rinse the samples in three successive washes of 5 minutes in TNT×. 3. Transfer the basket rack into TNblock for 1 h before overnight incubation with anti-digoxygenin (or anti-fluorescein or antiDNP horseradish peroxidase) in blocking solution at 4 °C without shaking. 4. On the next day, proceed to ten post-antibody washes for 15 min each in TNT×. 5. Samples are then transferred into 2 ml tubes and incubated in 100–150 μl of fluorophore tyramide amplification reagent for

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40 min (fluorophore diluted 1:50 in 1× working solution) (see Note 12). 6. Samples are washed twice in TNT× and in PTW prior to incubation in Hoechst (1:100) in PBT× for 30 min, followed by two washes of 10 min in PBT× and one wash in PBS. 7. Samples can finally be mounted in ProLong™ Gold Antifade Mountant with DNA Stain DAPI as described above. 3.8 Immunofluorescence

If using transgenic animals or a primary antibody against an endogenous protein, an alternative protocol can be to use one probe followed by an immunostaining. In this case, fix the embryos and proceed to the fluorescence in situ protocol as described above. 1. Samples are then incubated in ImmunoBlock for 1 h at room temperature before overnight incubation with primary antibodies (diluted in ImmunoBlock) at 4 °C (see Note 15). 2. On the next day, they are washed four times for 30 min with PTW, before being incubated for 1 h at room temperature in ImmunoBlock and then overnight at 4 °C in Alexa Fluor conjugated secondary antibodies (1:200). 3. On the final day, samples are washed four times for 30 min with PTW to remove excess antibody and then incubated for 30 min in Hoechst (1:100) and mounted in ProLong™ Gold Antifade Mountant with DNA Stain DAPI as previously described.

4

Notes General comments: rinse the basket racks with autoclaved MQ H2O between each wash. At the end of the protocol, rinse all the equipment (baskets, tube/basket racks) with 1 M sodium hydroxide and then with autoclaved MQ H2O. 1. Nematostella eggs and embryos stick to fresh plasticware and glass pipettes. Prerinse the pipettes in Nematostella medium, and gently scrub and wash petri dishes in water several times before use to avoid them being sticky. Reuse dishes and pipettes. For fixation, fill the tubes with Nematostella medium or with Fixative 2 for >1 h prior to use. To avoid damage to embryos and to plastic dishes, use blunted Pasteur pipettes or plastic transfer pipettes. The blunted glass Pasteur pipettes are made by briefly placing the Pasteur pipette tip in a Bunsen burner flame to smoothen the edges of the opening without reducing the diameter. 2. Acetic anhydride in solution has a short half-life and should be prepared right before use. Typically, acetic anhydride is added

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to the TEA buffer 30 seconds before adding the solution in the basket rack. 3. TSA Plus Fluorescein and TSA Plus Cy3 (Akoya Biosciences) detection kits use horseradish peroxidase (HRP, coupled to the antibody detecting the labeled probe) to catalyze the formation of radicals of TSA-fluorescein (or Cy3) conjugates. These radicals are then deposited close to the site where the probe localizes. In our hands, these kits improve the signal and the signal to noise ratio considerably. 4. The tip rack (the piece of plastic that the 1 ml tips rest on in a pipette tip box) can be cut to desired size by using a warm scalpel blade (use a scalpel with metal shaft). Feet for the rack can be made of cut 1 ml pipette tips (the upper part of the tip) melted and glued to the bottom of the tip rack. The height of the feet should be so that the baskets are not resting at the “bottom” of the rack but will have some distance to the bottom of the box allowing buffer to flow freely. Ideally use a rack that fits perfectly in the pipette tip box lid. Then two lids can serve as top and bottom of the basket rack box. But any plastic box that is chemical and heat resistant and has a lid can be used to hold the tip rack, as long as the bottom part of the baskets will be submerged in the solutions. 5. From planula stage on, embryos are able to contract. When working with swimming planulae, relax them by adding a large drop (0.5–1 ml) of magnesium chloride to the petri dish prior to fixation. Same, when working with primary polyps, it is essential to relax them properly with magnesium chloride prior to fixation. It might be that the amount of magnesium chloride added to the dish needs to be adjusted. Once fully relaxed (tentacles out with no movements upon touching them with a pipette), proceed to fixation. For primary polyps, it is easiest to add a few drops of Fixative 2 to the dish in a fume hood. After 2 min, it will be possible to collect them easily at the center of the petri dish, transfer them into a 2 ml tube, and proceed to Fixative 2. Before pipetting them, prerinse the pipette with PTW, to ensure polyps do not stick to the pipette tip walls. 6. Do not add more than approximately 200 embryos per baskets; otherwise, they will accumulate on top of each other, limiting exposure to the buffers and reagents along the protocol. The individual baskets are difficult to label, and make sure that you can identify the baskets/samples by their position in the rack, for example, by leaving one of the corner slots empty. 7. After the proteinase K treatment, it is imperative to not shake samples before re-fixation step.

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8. We have no indication that cooling of the samples during the step of adding probes affects the level of background. However, we do not recommend to place probes on ice after denaturation at 85 °C, as the hybridization buffer contains SDS that can precipitate from the solution at low temperatures. Rather keep the denatured probes at room temperature, and work safe and quick to add them to the Eppendorf rack. 9. It is recommended to do this step over the weekend. 10. Used probe in HB can be reused if kept at -20 °C. 11. Whether you use anti-digoxigenin, anti-fluorescein, or antiDNP antibody first does not matter. However, it is recommended to start with the strongest probe and finish with the weakest one. As the last probe will endure fewer washes, the staining might be better. If in doubt, use the anti-digoxigenin antibody first; in our experience, digoxigenin-labeled probes tend to give stronger signal, but also more background; many washes often improve its signal. Do not use anti-fluorescein antibody after using the TSA-fluorescein signal detection kit; the antibody would bind to the fluorescein signal from the first probe. 12. After a couple of washes, following the incubation in fluorophore tyramide amplification reagent, it is recommended to pipette few samples out of the tube and check their fluorescence under a fluorescence stereomicroscope. If the signal is not strong enough, it is possible at this step to repeat the labeling one more time for 30 min. This can be helpful and improve the signal for very weak probes. 13. If you have many samples, transfer the samples back into baskets in order to quicken the following washing steps. It is, however, possible to keep them in tubes. 14. This step is crucial to ensure that only the second probe is detected by the second TSA reaction. It is recommended to include a control sample in the experiment. This control will be incubated only with the first probe but will be stained with both TSA kits. If by the end of the protocol, both staining perfectly co-localize, it is clear that the inactivation of the peroxidase did not work properly. In this case, add the following extra washing step to your protocol to ensure the inactivation of the peroxidase. Incubate samples in StopPOD2 in complete darkness for 30–40 min, and then proceed to the TNT× washes. 15. The following primary antibodies were used after a FISH protocol: to detect NvInsm1::memGFP, anti-GFP (mouse, abcam1218, 1:200) [22]; to detect mOrange, anti-dsRed (rabbit, Clontech 632,496, 1:100) [35]; anti-NCol3 [36]. We noticed that some lot numbers of anti-GFP (and anti-dsRed)

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worked better than others. It is therefore recommended to always test them first in a regular immunostaining before using them after FISH. For combining FISH and IHC, we also observed a significant level of variation between batches of embryos, with some batches not giving any staining. In our hands, optimization of antibodies and the label for the probe has to be repeated for different antigen-target gene combinations. References 1. Kang YN, Fung C, Vanden Berghe P (2021) Gut innervation and enteric nervous system development: a spatial, temporal and molecular tour de force. Development 148:dev182543 2. Nagy N, Goldstein AM (2017) Enteric nervous system development: a crest cell’s journey from neural tube to colon. Semin Cell Dev Biol 66:94–106 3. Park E, Hwang DS, Lee JS, Song JI, Seo TK, Won YJ (2012) Estimation of divergence times in cnidarian evolution based on mitochondrial protein-coding genes and the fossil record. Mol Phylogenet Evol 62:329–345 4. dos Reis M, Thawornwattana Y, Angelis K, Telford MJ, Donoghue PC, Yang Z (2015) Uncertainty in the timing of origin of animals and the limits of precision in molecular timescales. Curr Biol 25:2939–2950 5. Technau U, Steele RE (2011) Evolutionary crossroads in developmental biology: Cnidaria. Development 138:1447–1458 6. Technau U, Genikhovich G, Kraus JE (2015) Cnidaria. In: Wanninger A (ed) Evolutionary developmental biology of invertebrates, vol 1. Springer, Vienna, pp 115–163 7. Chang ES, Neuhof M, Rubinstein ND, Diamant A, Philippe H, Huchon D, Cartwright P (2015) Genomic insights into the evolutionary origin of Myxozoa within Cnidaria. Proc Natl Acad Sci U S A 112:14912– 14917 8. Kayal E, Bentlage B, Pankey MS, Ohdera AH, Medina M, Plachetzki DC, Collins AG, Ryan JF (2018) Phylogenomics provides a robust topology of the major cnidarian lineages and insights on the origins of key organismal traits. BMC Evol Biol 18:1–18 9. Gold DA, Jacobs DK (2013) Stem cell dynamics in Cnidaria: are there unifying principles? Dev Genes Evol 223:53–66 10. Watanabe H, Hoang VT, Mattner R, Holstein TW (2009) Immortality and the base of multicellular life: lessons from cnidarian stem cells. Semin Cell Dev Biol 20:1114–1125

11. Layden MJ, Rentzsch F, Rottinger E (2016) The rise of the starlet sea anemone Nematostella vectensis as a model system to investigate development and regeneration. WIREs Dev Biol. https://doi.org/10.1002/wdev.1222 12. Frank U, Nicotra ML, Schnitzler CE (2020) The colonial cnidarian Hydractinia. EvoDevo 11:7 13. Vogg MC, Galliot B, Tsiairis CD (2019) Model systems for regeneration: Hydra. Development 146:dev177212 14. Weissbourd B, Momose T, Nair A, Kennedy A, Hunt B, Anderson DJ (2021) A genetically tractable jellyfish model for systems and evolutionary neuroscience. Cell 184(5854–5868): e5820 15. Ikmi A, McKinney SA, Delventhal KM, Gibson MC (2014) TALEN and CRISPR/Cas9mediated genome editing in the earlybranching metazoan Nematostella vectensis. Nat Commun 5:5486 16. Renfer E, Amon-Hassenzahl A, Steinmetz PR, Technau U (2009) A muscle-specific transgenic reporter line of the sea anemone, Nematostella vectensis. Proc Natl Acad Sci U S A 107:104–108 17. Hand C, Uhlinger K (1992) The culture, sexual and asexual reproduction, and growth of the sea anemone Nematostella vectensis. Biol Bull 182:169–176 18. Magie CR, Daly M, Martindale MQ (2007) Gastrulation in the cnidarian Nematostella vectensis occurs via invagination not ingression. Dev Biol 305:483–497 19. Kraus Y, Technau U (2006) Gastrulation in the sea anemone Nematostella vectensis occurs by invagination and immigration: an ultrastructural study. Dev Genes Evol 216:119–132 20. Sebe-Pedros A, Saudemont B, Chomsky E, Plessier F, Mailhe MP, Renno J, Loe-Mie Y, Lifshitz A, Mukamel Z, Schmutz S et al (2018) Cnidarian cell type diversity and regulation revealed by whole-organism single-cell RNA-Seq. Cell 173(1520–1534):e1520

FISH and Cell Type Characterization in Nematostella 21. Steger J, Cole AG, Denner A, Lebedeva T, Genikhovich G, Ries A, Reischl R, Taudes E, Lassnig M, Technau U (2022) Single-cell transcriptomics identifies conserved regulators of neuroglandular lineages. Cell Rep 40:111370 22. Tourniere O, Gahan JM, Busengdal H, Bartsch N, Rentzsch F (2022) Insm1expressing neurons and secretory cells develop from a common pool of progenitors in the sea anemone Nematostella vectensis. Sci Adv 8: eabi7109 23. Richards GS, Rentzsch F (2014) Transgenic analysis of a SoxB gene reveals neural progenitor cells in the cnidarian Nematostella vectensis. Development 141:4681–4689 24. Sinigaglia C, Busengdal H, Lecle´re L, Technau U, Rentzsch F (2013) The bilaterian head patterning gene six3/6 controls aboral domain development in a cnidarian. PLoS Biol 11:e1001488 25. Magie CR, Pang K, Martindale MQ (2005) Genomic inventory and expression of sox and fox genes in the cnidarian Nematostella vectensis. Dev Genes Evol 215:618–630 26. Layden MJ, Boekhout M, Martindale MQ (2012) Nematostella vectensis achaete-scute homolog NvashA regulates embryonic ectodermal neurogenesis and represents an ancient component of the metazoan neural specification pathway. Development 139:1013–1022 27. Tourniere O, Dolan D, Richards GS, Sunagar K, Columbus-Shenkar YY, Moran Y, Rentzsch F (2020) NvPOU4/Brain3 functions as a terminal selector gene in the nervous system of the cnidarian Nematostella vectensis. Cell Rep 30(4473–4489):e4475 28. Busengdal H, Rentzsch F (2017) Unipotent progenitors contribute to the generation of sensory cell types in the nervous system of the

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cnidarian Nematostella vectensis. Dev Biol 431:59–68 29. Faltine-Gonzalez D, Havrilak J, Layden MJ (2023) The brain regulatory program predates central nervous system evolution. Sci Rep 13: 8626 30. Renfer E, Technau U (2017) Meganucleaseassisted generation of stable transgenics in the sea anemone Nematostella vectensis. Nat Protoc 12:1844–1854 31. Rentzsch F, Renfer E, Technau U (2020) Generating transgenic reporter lines for studying nervous system development in the cnidarian Nematostella vectensis. Methods Mol Biol 2047:45–57 32. Scholz CB, Technau U (2003) The ancestral role of Brachyury: expression of NemBra1 in the basal cnidarian Nematostella vectensis (Anthozoa). Dev Genes Evol 212:563–570 33. Wolenski FS, Layden MJ, Martindale MQ, Gilmore TD, Finnerty JR (2013) Characterizing the spatiotemporal expression of RNAs and proteins in the starlet sea anemone, Nematostella vectensis. Nat Protoc 8:900–915 34. Fritzenwanker JH, Technau U (2002) Induction of gametogenesis in the basal cnidarian Nematostella vectensis(Anthozoa). Dev Genes Evol 212:99–103 35. Nakanishi N, Renfer E, Technau U, Rentzsch F (2012) Nervous systems of the sea anemone Nematostella vectensis are generated by ectoderm and endoderm and shaped by distinct mechanisms. Development 139:347–357 36. Zenkert C, Takahashi T, Diesner MO, Ozbek S (2011) Morphological and molecular analysis of the Nematostella vectensis cnidom. PLoS One 6:e22725

Chapter 5 SABER-FISH in Hydractinia Miguel Salinas-Saavedra Abstract In situ hybridization allows the detection of nucleic acid sequences in fixed cells and tissues. The gelatinous nature of cnidarians and Hydractinia demands extensive and exhausting protocols to detect RNA transcripts with traditional methods (e.g., colorimetric in situ hybridization). Signal amplification by exchange reaction (SABER) fluorescence in situ hybridization (FISH) enables simplifying and multiplex imaging of RNA targets in a rapid and cost-effective manner. In one enzymatic reaction, SABER-FISH uses a stranddisplacing polymerase and catalytic DNA hairpin to generate FISH probes with adjustable signal amplification, allowing highly sensitive detection of nucleic acids and reducing the number of required probes. Here I describe the methodology to detect transcripts within the cells of Hydractinia by SABER-FISH in wholemount samples. Key words SABER-FISH, Cnidaria, Hydractinia, RNA, In situ hybridization

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Introduction Cnidarians are of great relevance to understanding cellular and molecular mechanisms in an evolutionary context since they are the sister taxa of all bilaterian animals. Cnidarians are classical model organisms to study development and regeneration due to their remarkable regenerative ability, plasticity, and rapid embryogenesis [1–3]. However, within the Cnidaria, only a few species count with established molecular biology protocols essential to functionally investigate biological processes, including visualization of RNA molecules to determine their spatiotemporal expression. The structural organization of cnidarian tissues and their high mucous levels demand extensive washes and probe development periods with classical in situ hybridization (ISH) methods [4–7]. Recent resource development of ISH protocols in diverse organisms has enabled researchers to optimize the visualization of RNAs by increasing signal/noise ratio and by multiplexing [8, 9]. SABER-FISH [10] uses the simplicity of the primer-exchange reaction (PER) to synthesize single-stranded (ssDNA) concatemers

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Diagrammatic illustration of the PER reaction for probe synthesis and hybridization. (a) The PER reaction combines a strand-displacing polymerase, a catalytic DNA hairpin, and ssDNA primers in a single tube to create FISH probes with adjustable signal amplification (extended concatemer). (b) In SABER-FISH, there are two hybridization steps. First, the probe sets (with extended concatemer) are incorporated and incubated with the polyps at 43
C, hybridizing with the target mRNAs. Then, short fluorescent oligonucleotides (fluor oligos) are incubated with the samples at 37
C, hybridizing with the extended concatemers

of a desired length (programmable) in vitro. The concatemers attached to oligonucleotide-based probes give a rapid signal amplification detected by secondary hybridization with fluorescent oligonucleotides (fluorescent imagers), enhancing the functionality of single-molecule RNA FISH probe pools. Probe synthesis by PER reaction (Fig. 1) is similar to PCR, takes 1–3 h, and requires widely available and inexpensive reagents [11]. Fixation and pre-hybridization treatments are practically identical to classical FISH protocols. However, post-hybridization washes and signal development only take a few hours, including the secondary hybridization of fluorescent imagers. Hence, SABERFISH is a simple, rapid, and accessible alternative to more expensive and restricted modern FISH methods. These characteristics are essential for researching emerging model organisms, allowing a cost-effective assessment of unknown and low-expressed transcripts. Here I describe a protocol for detecting the expression of individual RNAs by SABER-FISH in the colonial hydrozoan Hydractinia symbiolongicarpus.

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Materials 1. Probes: Use Oligominer software (https://github.com/ beliveau-lab/OligoMiner) to design oligonucleotide-based RNA FISH probes according to the protocol described in Kishi et al. [10], integrating the Hydractinia genome. The Oligominer software will generate a list of unique oligo sequences per each target gene. This protocol works using 1 and up to 15 oligonucleotide-based probes. Choose at least five oligos when possible. RNA FISH probes are in vitro synthesized with primer sequences on their 30 ends [10], extended into concatemers using a catalytic hairpin. Ideally, select nonadjacent oligo sequences that do not end in a poly (A) or poly(T) since they may interfere with the hairpin hybridization reaction. Aim to amplify concatemers between 400 and 700 base pairs for better penetrance. This protocol is optimized to work with Piwi1 and Vasa transcripts (Fig. 2) as well as other genes in previously published works [12, 13], using primer sequences number 30, 27, and 25 (sequences number from Kishi et al. [10]) with their respective fluor oligos [10] (see Note 1). 2. 4% MgCl2: prepare 4% MgCl2 W/Vol in 1:1 solution of fresh seawater (FSW) and distilled water. 3. PBSTw: 1 phosphate buffered saline (PBS), 0.1% Tween-20 in nuclease-free water. 4. Fixation glass dishes: 40 mm soda lime glass petri dishes. 5. Fixative I: 0.4% glyoxal, 4% formaldehyde in FSW at room temperature (RT) (see Note 2). 6. Fixative II: 0.1% glyoxal, 4% formaldehyde in ice-cold PBSTw.

Fig. 2 Double mRNA SABER-FISH of a regenerating Hydractinia polyp. (a) Piwi1 labeling using hairpin 30 and Atto633 fluor oligo probes. (b) Vasa Piwi1 labeling using hairpin 27 and Atto565 fluor oligo probes. (c) Merge composite including nuclear marker (Hoechst; blue). Blastema is at the top. Scale bars: 20 μm

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7. N-acetyl cysteine: 7.5% stock in pure water. 8. 0.2 M HCl in DEPC water. 9. Methanol. 10. Acetone. 11. Hydrogen peroxide (H2O2). 12. Glycine: prepare stock at 2 M concentration, and filter it once in solution with PBTw. Make 0.05 M dilution fresh every time. 13. Triethanolamine. 14. Acetic anhydride. 15. 24-well tissue culture plate. 16. Platform rocker. 17. Sealed humid chamber: plastic container with airtight lid (e.g., lunchbox). 18. Hybridization oven or incubator. 19. Wash hybridization (Whyb): 2 SSC (from 20 stock, pH 7), 1% Tween-20, 40% deionized formamide. 20. Hyb 1 (for pre-hybridization): 2 SSC (from 20 stock, pH 7), 1% Tween-20, 40% formamide, 10% dextran sulfate. 21. Hybe buffer (for primary probe hybridization): 40% formamide, 5 SSC, 0.05 mg/mL heparin, 0.25% Tween-20, 1% SDS, 1 mg/mL salmon sperm DNA, 1 mg/mL Roche blocking buffer powder. Dissolve all the solutions in DEPC water. 22. Hyb 2 (for fluorescent detection): 1 PBS, 0.2% Tween-20, 10% dextran sulfate. 23. WHyb 2: 1 PBS, 0.1% Tween-20, 30% formamide. 24. 2 SSCTw: 2 SSC pH 7.0, 0.1% Tween-20. 25. Hybe/Probe mix: 80% Hybe buffer, probes, bring to volume with ddH2O. Try using each probe for the first time at a concentration of 1 μg/120 μL volume. For example, 96 μL Hybe buffer, 5 μL 200 ng/μL probe 1, 5 μL 200 ng/μL probe 2, 14 μL ddH2O. 26. Fluor/Hyb2 mix: 80% Hyb2 buffer, Fluor Oligos (0.2–1 μM each), bring to volume with ddH2O. Try using each Fluor Oligo for the first time at a concentration of 0.2 μM. For example, 96 μL Hyb2, 2.4 μL Fluor Oligo 1 (10 μM), 2.4 μL Fluor Oligo 2 (10 μM), 19.2 μL ddH2O. 27. Hoechst (20 mg/mL). 28. 80% glycerol. 29. 97% 2,20 -thiodiethanol (TDE) in PBS. 30. Microscope slides and coverslips. 31. Access to a confocal microscope

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Methods Fixation

1. Relax animals in 2 mL 4% MgCl2 at RT for 10–30 min. 2. Replace the solution with 2 mL Fixative 1 to the samples. Leave for 90 s at RT (see Note 3). 3. Remove Fixative 1. Add 2 mL Fixative 2; place animals on rocker at 4
C for 1 h. 4. Remove Fixative 2. Wash samples 3 with PBSTw, 5 min each time (see Note 4).

3.2 Postfixation Wash and Dehydration for Storage

Carry out all procedures with ice-cold reagents unless otherwise specified. 1. Wash 2 with 25% methanol in 75% PBSTw, 5 min each time. 2. Wash 2 with 50% methanol in 50% PBSTw, 5 min each time. 3. Wash 1 with 75% methanol in 25% PBSTw, 1 min. 4. Wash 3 with 100% methanol, 1 min each time. 5. Place samples at 220
C for storage (see Note 5).

3.3

Bleaching

1. 1% H2O2 in 100% methanol for 45-min wash (in a rocker at 4
C).

3.4

Permeabilization

1. Wash 1 with 75% methanol in 25% acetone, 1 min. 2. Wash 1 with 50% methanol in 50% acetone, 1 min. 3. Wash 2 with 25% methanol in 75% acetone, 1 min each time. Leave for 20 min on ice after second wash. 4. Wash 1 with 50% methanol in 50% acetone, 1 min. 5. Wash 1 with 75% methanol in 25% acetone, 1 min. 6. Wash 1 with 100% methanol, 1 min. Continue to rehydration or store samples (see Note 5).

3.5

Rehydration

1. Wash 1 with 75% methanol in 25% PBSTw, 5 min. 2. Wash 1 with 50% methanol in 50% PBSTw, 5 min. 3. Wash 1 with 25% methanol in 75% PBSTw, 5 min. Carry out all the following procedures at room temperature reagents unless otherwise specified. 4. Wash 2 with 100% PBSTw, 1 min each time. Additional optional washes to improve signal/noise ratio 5. Wash 1 with 0.2 M HCl (in DEPC water) for 20 min in a rocker (see Note 6). 6. Wash 2 with DEPC water, 1 min each time. 7. Wash 3 with 100% PBSTw, 1 min each time.

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Additional optional washes to quench formaldehyde activity 8. Wash 2 with 0.05 M glycine in PBSTw, 5 min each time. 9. Wash 3 with PBSTw, 5 min each time. 10. Move the samples into a 24-well tissue culture plate, and distribute according to how many samples/controls one will have (recommendation: ten polyps per well). 3.6 Charge Removal for Nonspecific Binding in Tissues

1. Wash 1 with 1% triethanolamine (pH 8 – titrate pH with HCl) in PBSTw, 5 min. 2. Wash 1 with 1% triethanolamine (pH 8) in PBSTw – premixed with 6 μL of acetic anhydride (3 μL for embryos), 5 min (see Note 7). 3. Wash 1 5 min 1% triethanolamine (pH 8) in PBSTw – premixed with 12 μL of acetic anhydride (6 μL for embryos), 5 min (see Note 7). 4. Wash 3 with 100% PBSTw, 5 min each time.

3.7

Pre-hybridization

1. Remove most of the PBSTw without drying the samples. 2. Replace PBSTw with Whyb buffer (prewarmed to 43
C). 3. Place in hybridization oven set to 43
C, 10 min minimum. 4. Replace Whyb buffer with Hyb1 buffer (prewarmed to 43
C), and place in a sealed humid chamber overnight (or 4–6 h) in oven set to 43
C (see Note 8). Pause point: samples can be stored in Hyb1 after pre-hybridization at 20
C for several weeks.

3.8

Hybridization

1. Preheat Hybe/Probe mix at 60
C, 3 min. 2. Remove old Hyb1 buffer and add the prewarmed Hybe/ Probe mix (see Note 9). 3. Incubate at hybridization temperature in oven at 43
C from 16 h (minimum) to 2 days (or during the weekend).

3.9 Posthybridization

Carry out all procedures at hybridization temperature and prewarmed reagents unless otherwise specified. 1. Replace Hybe/Probe mix with Whyb, 10-min wash. 2. Wash 2 with Whyb, 30 min each time. 3. Wash 1 with 50% Whyb in 50% 2X SSCTw, 10 min. 4. Wash 2 with 100% 2X SSCTw, 10 min. 5. Return to room temperature. Pause point: sample can be stored in 2X SSCT or PBSTw at 4
C for several weeks.

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1. Replace 2X SSCTw with PBSTw, two washes, 1 min each time at room temperature. 2. Set oven to 37
C. 3. Transfer chamber to 37
C for hybridization and subsequent wash steps. 4. Once samples are warm, remove PBSTw and add Hyb2/fluor solution (prewarmed at 37
C). 5. Incubate for at least 10 min to 1 h at 37
C (see Note 10). 6. Replace Hyb2 with prewarmed Whyb2, 10 min wash maximum (see Note 11). 7. Replace Hyb2 with prewarmed PBSTw, 1 min wash. 8. Wash 2 with prewarmed PBSTw, 5 min each time. 9. Return to room temperature. Pause point: samples can be stored at 4
C without obvious signal loss for at least 1 week. 10. (Optional) Add preferred nuclear marker (e.g., Hoechst (20 mg/mL) 1:2000 in PBSTw), and incubate for 30 min at RT. 11. Wash 2 with PBSTw, 5 min each time.

3.11 Sample Mounting

1. Mount the samples for imaging. Transfer the samples onto microscope slides, and cover them with 30 μL of 80% glycerol or 97% TDE mounting media for clearing. 2. Carefully place a coverslip on top, and seal its edges with nail polish. Allow it to harden for 30 min. 3. Store the samples for future imaging (see Note 12).

3.12 Imaging and Image Analysis

1. To view the mounted samples, use a confocal microscope, and select the appropriate laser and filter combination for the chosen fluorophores. 2. Locate the sample using a 20 objective, and adjust the magnification with a 40 or 60 objective to detect and capture the fluorescent signal. 3. Calibrate the laser power and acquisition parameters for minimal background and puncta detection of the target mRNA in the cell type and tissue of interest. 4. Raw images can be visualized and analyzed using ImageJ/Fiji software. 5. Images can be imported into Adobe Photoshop to further brightness and contrast adjustment.

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Notes 1. When choosing the right fluorophore combination, it is relevant to note that Hydractinia tissues may emit autofluorescence in the green channel. Adding glyoxal to the fixative can significantly decrease this technical artefact [14]. 2. Optional additive, 0.375% N-acetyl cysteine (from 7.5% stock). This fixative additive may help to dissolve mucus and preserve mRNA. 3. Consider using dedicated fixation glass dishes when fixing animals. Clean these dishes with water and 70% ethanol after each use. 4. Alternative fixative solutions – Fixative 1: 0.2% glutaraldehyde, 4% formaldehyde in FSW at RT; Fixative 2: 4% formaldehyde in ice-cold PBSTw. Although these fixative solutions are highly effective, they can cause an increase in autofluorescence in tissues. It is recommended to use them for troubleshooting fixative-related issues. 5. If the results when using this stop point are unsatisfactory, consider not storing the samples, and continue with the protocol. 6. These washes are optional, may increase tissue permeability, and reduce noise signals of the tentacles and connective tissues. However, it may be detrimental to some genes and embryonic tissue. Therefore, it is advisable to perform when necessary. 7. Critical step: check and adjust the solution mix pH ¼ 8.0 before adding acetic anhydride. Failure to do so may result in less efficient charge removal and potentially increase nonspecific binding. Acetic anhydride drop takes 5–10 min to dissolve. To ensure that the acetic anhydride dissolves properly, premix the solution in a rocker at room temperature for at least 10 min before using it for washing purposes. 8. While it is possible to complete all the steps in 1 day, it is advisable to pre-hybridize overnight to achieve optimal results. 9. Hyb1 is very viscous; be careful when removing it. Sometimes it is better to dilute it by adding some Whyb to the solution. 10. The incubation time will depend on your target genes. However, 2 h of incubation may generate an unspecific signal. 11. This step is critical if you have an unspecific signal from your imager. It will depend on the sample and experiment. 12. For best results, it is advised to image the samples within 4 days of the second hybridization with the fluorescent oligonucleotides. If the samples are mounted for more than a week, the fluorescence may spread to other tissues, causing nonspecific background and signal loss.

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Acknowledgments I thank members of the Frank lab for discussions and advice. Confocal images were taken at the Centre for Microscopy and Imaging Core Facility at University of Galway. MSS is a Human Frontier Science Program Long-Term Postdoctoral Fellow (grant no. LT000756/2020-L). References 1. Chrysostomou E, Febrimarsa DBT, Frank U (2022) Gene manipulation in Hydractinia. Methods Mol Biol 2450:419–436 2. Frank U, Nicotra ML, Schnitzler CE (2020) The colonial cnidarian Hydractinia. Evodevo 11:7 3. Technau U, Steele RE (2011) Evolutionary crossroads in developmental biology: Cnidaria. Development 138(8):1447–1458 4. DuBuc TQ, Schnitzler CE, Chrysostomou E, McMahon ET, Febrimarsa GJM, Buggie T, Gornik SG, Hanley S, Barreira SN, Gonzalez P, Baxevanis AD, Frank U (2020) Transcription factor AP2 controls cnidarian germ cell induction. Science 367(6479): 757–762 5. Sanders SM, Shcheglovitova M, Cartwright P (2014) Differential gene expression between functionally specialized polyps of the colonial hydrozoan Hydractinia symbiolongicarpus (phylum Cnidaria). BMC Genomics 15(1):406 6. Sinigaglia C, Thiel D, Hejnol A, Houliston E, Leclere L (2018) A safer, urea-based in situ hybridization method improves detection of gene expression in diverse animal species. Dev Biol 434(1):15–23 7. Wolenski FS, Layden MJ, Martindale MQ, Gilmore TD, Finnerty JR (2013) Characterizing the spatiotemporal expression of RNAs and proteins in the starlet sea anemone,Nematostella vectensis. Nat Protoc 8(5):900–915 8. Young AP, Jackson DJ, Wyeth RC (2020) A technical review and guide to RNA fluorescence in situ hybridization. PeerJ 8:e8806

9. Nielsen BS, JonesIn J (5 ed) (2020) In situ hybridization protocols. Methods in Molecular Biology, vol 2148. Springer US. https://doi. org/10.1007/978-1-0716-0623-0 10. Kishi JY, Lapan SW, Beliveau BJ, West ER, Zhu A, Sasaki HM, Saka SK, Wang Y, Cepko CL, Yin P (2019) SABER amplifies FISH: enhanced multiplexed imaging of RNA and DNA in cells and tissues. Nat Methods 16(6): 533–544 11. Hosoda E, Hiraoka D, Hirohashi N, Omi S, Kishimoto T, Chiba K (2019) SGK regulates pH increase and cyclin B-Cdk1 activation to resume meiosis in starfish ovarian oocytes. J Cell Biol 218(11):3612–3629 12. Chrysostomou E, Flici H, Gornik SG, SalinasSaavedra M, Gahan JM, McMahon ET, Thompson K, Hanley S, Kilcoyne M, Schnitzler CE, Gonzalez P, Baxevanis AD, Frank U (2022) A cellular and molecular analysis of SoxB-driven neurogenesis in a cnidarian. elife 11:e78793 13. Salinas-Saavedra M, Febrimarsa KG, Horkan HR, Baxevanis AD, Frank U (2023) Senescence-induced cellular reprogramming drives cnidarian whole-body regeneration. Cell Rep 42:112687 14. Yao RW, Luan PF, Chen LL (2021) An optimized fixation method containing glyoxal and paraformaldehyde for imaging nuclear bodies. RNA 27(6):725–733

Chapter 6 smFISH for Plants Sahar Hani, Caroline Mercier, Pascale David, Thierry Desnos, Jean-Marc Escudier, Edouard Bertrand, and Laurent Nussaume Abstract Single-molecule fluorescence in situ hybridization (smFISH) is a powerful method for the visualization and quantification of individual RNA molecules within intact cells. With its ability to probe gene expression at the single cell and single-molecule level, the technique offers valuable insights into cellular processes and cell-to-cell heterogeneity. Although widely used in the animal field, its use in plants has been limited. Here, we present an experimental smFISH workflow that allows researchers to overcome hybridization and imaging challenges in plants, including sample preparation, probe hybridization, and signal detection. Overall, this protocol holds great promise for unraveling the intricacies of gene expression regulation and RNA dynamics at the single-molecule level in whole plants. Key words smFISH, Plant, RNA, Transcription, Imaging, Single-molecule

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Introduction During the past few decades, innumerable studies have shed light on the importance of transcriptional regulation, RNA molecules, and their functions. A central regulatory role was thus attributed to RNA. Indeed, apart from its primary function as a messenger for protein synthesis, the roles of RNAs over the years have been found to be more diverse and prevalent than were initially thought. RNA molecules fulfill diverse tasks in the cell, through their secondary structure and interaction with various proteins and nucleic acids and by also being the key constituents of cellular machineries (ribosomes, tRNAs, etc.). Tight regulations at the level of expression, posttranscriptional modifications, and RNA localization have been shown to be compulsory for many biological functions of RNAs [1, 2].

Sahar Hani, Caroline Mercier and Pascale David contributed equally with all other contributors. Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Intracellular transcript localization has gained a lot of interest over the years as it is a ubiquitous mechanism taking place in a wide range of organisms, from bacteria to humans. It offers local protection from degradation and restricts the transcript spatially and temporally to discrete sites within the cell, where it is of use. It is also an efficient means of mass production of localized proteins, which would facilitate protein-protein interactions, folding pathways, and posttranslational modifications [3–5]. RNA localization is a highly regulated process, indispensable to cell development, polarity and differentiation, determination of cell fate and tissue functionality, signaling, and physiology. Nonuniform subcellular mRNA distribution has been described in animal oocytes [6, 7] as well as specialized cells [8–11] and shown to be essential for proper embryonic patterning during development [12, 13]. More recently, these discoveries extended outside of the animal kingdom, whereby specific intracellular mRNA targeting was observed also in fungi [14, 15], bacteria [16], and plants (reviewed in [4, 17]). While intracellular RNA localization is a conserved process across all kingdoms, less effort has been put in its study in higher plants as compared to the animal phyla. One reason may be linked to the pecto-cellulosic cell wall, which acts as a physical barrier often reducing the efficiency of cell biology techniques. Besides, mature plant tissues are composed of differentiated cells containing a large vacuole compartment which pushes the cytoplasm to the periphery of the cell, whereas more cytoplasmically dense cells as meristematic ones are very small in size, making the observation of RNA in confined areas difficult. Despite this, a number of studies focused on the differential localization of transcripts in plants and their fundamental roles in various cellular processes. Evidence for polar localization of mRNA in plant cells include the differential subcellular localization of expansin mRNA in xylem cells of Zinnia elegans, exclusively to their apical or basipetal end depending on the expansin gene and organ. Other examples include the accumulation of profilin mRNA at the tips of emerging root hair in higher plants [18] and the formation of basal/apical gradients of mRNA with development-specific patterns of distribution in the unicellular green alga Acetabularia acetabulum [19]. Additionally, various types of mRNA have been shown to be asymmetrically distributed and concentrated in particular subcellular domains such as in developing rice endosperm [17], in ER compartments, in proximity to the mitochondria [20], and in chloroplast [21]. In addition to intracellular RNA localization, RNA species can move over long distances in plants, between different tissues and organs, to ensure cell-to-cell communication and the coordination of plant growth, development, and adaptation to biotic and abiotic stresses [22–25]. Indeed, plants rely on systemic signals where

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RNAs provide an efficient and specific remote-control system to orchestrate developmental and physiological processes. The systemic migration of such a high number of RNAs between plant cells raises numerous questions regarding the pivotal roles of RNAs in distant organs, the subtle mechanisms that allow such specificity and fine-tuning of directional transport, and how they may function in regulation of vital adaptational processes. Transcript profiling and RNA analysis provide powerful tools to unravel the complexity of molecular events taking place within biological systems. While various methods have been developed and used to reliably detect genome-wide changes in gene expression [26–29], they differ considerably in their abilities to detect distinct steps of gene expression and to identify various RNA species. Additionally, they often provide collective averaged data of many RNA molecules without considering the heterogeneity in the population or the fact that a transcript’s stability varies throughout its life span [30, 31]. More importantly, these techniques lack cellular resolution and fail to measure RNAs at the level of single cells or within cellular compartments [28, 30]. Hence, despite the progress in unraveling RNA functions, understanding the interplay between the different molecules and machineries remains a difficult task without the ability to visualize them in intact cells. Consequently, the implementation of techniques to visualize sequencespecific RNA is indispensable. In situ hybridization (ISH) has evolved significantly over the years, transitioning from a time-consuming monthlong assay to a much faster technique capable of detecting single transcripts in just a couple of days [32, 33]. This transformation was facilitated by the development of various enhancements, such as the use of radioactively labeled probes initially [33], followed by histochemical detection methods [34] and, notably, the introduction of fluorescent oligonucleotides [32]. Single-molecule fluorescent in situ hybridization (smFISH) became a powerful application of ISH that employs multiple fluorescently labeled DNA probes to visualize individual mRNA molecules as diffraction-limited fluorescent spots, enabling the study of gene expression patterns, RNA distribution, and transcription kinetics with cellular resolution. SmFISH has been used in various organisms and different cell types with variable success. As opposed to older methods, it provides cellular resolution and allows the study of gene expression patterns and asymmetric RNA distribution within cells and revealed cell-to-cell variability existing in tissues. Multiple technological advancements made it the method of choice for quantifying low-abundance mRNAs and providing insights on transcription kinetics and the bursty nature of gene expression [35, 36], mRNA export [37], translation, and even decay [38, 39]. Various improvements also enabled high quality three-dimensional imaging, multiplexing different RNA species, and co-visualization of

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RNA with proteins [40, 41] and paved the way for image-based transcriptomics [42, 43]. Although well documented, the smFISH workflow in general holds some drawbacks. A first inconvenience is the cost of synthesis of fluorescent oligonucleotide probes. Another major impediment is that smFISH always suffers from background and nonspecific binding of stray probes which generates false-positive signals and affects the ability to differentiate true targets from background noise. Minimizing these artifacts necessitates the use of a larger number of probes. To achieve this and reduce costs, a single-molecule inexpensive FISH (smiFISH) was developed [44]. The approach is based on the use of an increased number of unlabeled primary probes specific to the targeted RNA, which carry an extra readout sequence that can be detected by fluorescently labeled secondary probes. This in general yields higher signal-to-noise ratio and enhanced signal quality at low costs [43, 44]. In plants, the permeability of cell walls creates difficulties and restricts efficient probe penetration. Hence, hybridization outcomes were limited. Classically, mRNA tissue hybridization used sections of biological samples to allow better probe accessibility to deeply embedded cell types [45]. This provides access to the localization of mRNA in various plant tissues [46–48] and organs using Dig-labeled probes. It was shown to be reliable for the analysis of transcript localization in different developmental processes [49] but could not reach single-molecule resolution. Multicolor whole-mount ISH was also successfully implemented in plants [50]. To overcome the limited sensitivity provided by these techniques, more direct labeling of transcripts with fluorescently labeled probes has been employed. However, the optical properties of plant cells and tissues present considerable challenges for fluorescence microscopy. Many endogenous molecules emit high levels of background, and autofluorescence adversely affects detection efficiency. Nevertheless, recent advances done in that regard allowed to circumvent these problems and use smFISH in Arabidopsis roots [51]. This also enabled the quantification of mRNAs per cell and exploration of cell-to-cell variations to study RNA polymerase II transcription and gene bursting [36] and to explore different steps associated with the RNA regulation (Fig. 1). In the present work, we describe smFISH protocols that are successfully used in plants. We focus on Arabidopsis root since our genes of interest were highly expressed in that organ and rapidly altered due to nutritional stress [36]. However, the protocol also works well for leaves and stems. Overall, the technique provides a resource for plant researchers investigating transcription dynamics, RNA metabolism, and singlemolecule studies. Adequate computational tools and algorithms must be combined for image analysis, molecule counting, and

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Fig. 1 Diagram illustrating the events that can be studied by using FISH technique. Transcription is an event that can occur discontinuously over time, a phenomenon known as bursting. Within the transcription points, the RNA polymerase (brown shape) running through the DNA to produce the associated RNA carries out this task at a certain speed, which can vary from one gene to another. The RNAs are then spliced and exported to the cytoplasm, organelles of the cell where they are produced (e.g., chloroplast, mitochondria, etc.), or to other cells, tissues, or organs of the plant. Some of the RNA produced is also degraded, ensuring its turnover. All these events can be monitored using FISH

spatial organization characterization. Consequently, smFISH holds great promise for visualizing transcripts from birth to death and unraveling the intricacies of gene expression regulation in plants.

2 2.1

Materials Reagents

1. Sterile petri dishes. 2. Autoclaved plant growth medium: Murashige and Skoog medium diluted tenfold (MS/10), 5 g/L sucrose and 8 g/L agar, pH 5.7 (described in [52]), with 0.5 mM KH2PO4 or without phosphate (0.013 mM KH2PO4). 3. Sterilized Arabidopsis seeds. 4. Carbonate buffer: 0.1 M Na HCO3 pH 8.8. 5. Micropore tape. 6. x50, 0.7 M, pH 5.8 (or 7 if we want to observe GFP signal) stock solution of 2-(N-morpholino) ethane sulfonic acid (MES) buffer solution. 7. 4% PFA/MES: 4% paraformaldehyde (PFA) diluted in x1 (or 14 mM) MES buffered at pH 5.8 or 7 if we want to observe GFP. 8. Liquid nitrogen. 9. Forceps. 10. Glass microscopy slides (0.17-mm-thick) and coverslips. 11. Razor blade.

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12. 100%, 80%, 70% ethanol. 13. 20
saline-sodium citrate buffer (SSC). 14. 15% formamide solution: 15% formamide, 1
SSC in DDW. 15. Cy3 or Cy5 labeled oligonucleotide probes (amino-modified – C6 dT and labeled in vitro; see Subheading 3.2). 16. 20 mg/ml E. coli tRNA. 17. Autoclaved water. 18. 20 mg/mL RNAse-free BSA. 19. 200 mM vanadyl-ribonucleoside (nonmandatory).

complex

(VRC)

20. 40% dextran sulfate (100 mg/mL). 21. Anti-fading mounting medium containing DAPI (Prolong Diamond Gold, Invitrogen). 22. 3 M sodium acetate pH 5.2. 23. DMSO. 24. Tris-EDTA buffer. 25. Probe mix 1: 1xSSC, 360 ng/μL E. coli tRNA, 15% formamide, 0.4 ng/μL probe mix (see Notes 1 and 2). 26. Probe mix 2: 200 μg/μL RNase-free BSA, 2 mM VRC (vanadyl-ribonucleoside complex), 10.8% dextran sulfate. 2.2

Equipment

1. Plant growth chamber for Arabidopsis plants under a 16-h light/8-h dark regime with 25  C/22  C, respectively. 2. Heating block. 3. Horizontal shaker. 4. Zeiss Axioimager Z1 wide-field upright microscope equipped with a camera sCMOS ZYLA 4.2 MP (Andor), using a 100
, NA 1.4 Plan Apochromat oil objective. 5. Dragonfly (Oxford Instruments) equipped with four laser lines and an ultrasensitive EMCCD camera (iXon Life 888, Andor) mounted on a Nikon Eclipse Ti2 microscope body, a 40
, NA 1.3 Plan Fluor oil objective or a 60
, NA 1.4 Plan Apochromat oil objective coupled with a supplementary lens of 2
.

2.3

Software

Microscope image acquisition and analysis software (e.g., ImageJ, HotSpot [53], FISH-quant [54]).

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Methods

3.1 Probe Design (Fig. 2)

3.2 Probe Labeling Cy3 or Cy5

For DNA oligonucleotide design, aim for about 20–25 oligonucleotides of ~on average 45 (35–55) bases with 65–70  C Tm and a GC content below 60%. We use http://biotools.nubic.northwest ern.edu/OligoCalc.html, set with 50 mM salt, nearest neighbor, 10 nM primers, ssRNA. Use the reverse complementary strand for the mRNA of interest. Search for an area with a convenient Tm and where you can locate two internal T (on the probe strand) separated from each other and from extremity by at least ten bases. Adjust the length of the oligos to reach a proper Tm. Label the two internal T which will be modified into C6dT. Then add a 50 T X at the beginning of the oligo and X T at the 30 end. Both X will also correspond to C6dT modified base. Order the oligonucleotide with the C6dT modified bases. If gene belongs to multigenic family, choose the probes in location with at least less than 65% of homology. 1. One vial of Cy3 or Cy5 (Cytiva PA23001 or PA25001, Amersham) allows the preparation of 15 μg oligonucleotides. 2. For one vial, add 30 μL DMSO and vortex twice for 30 s. probe

…

5’ A

Y

3’

min 10 nt

min 10 nt

35/55 nt

other important parameters Tm

65 to 70°C

< 60%

For mulgenic family sequence < 65% homology

Fig. 2 Rules for designing the FISH probes. To study the transcription of a gene by FISH, you need to create 20–25 probes. Each probe must be between 35 and 55 nucleotides long. Only the thymidines (T) will be labeled. Thus, it is therefore important that a probe has at least two T, each spaced ten nucleotides apart and located at least ten nucleotides from the first and last positions of the probe. During probe synthesis, the T are replaced by a labeled nucleotide C6dT. The probe must also meet other criteria, such as having a melting temperature (Tm) of between 60  C and 68  C and a percentage of C/G (cytosine/guanine) of between 30% and 50%. To facilitate the application of these parameters, it is recommended to work from the reverse complement of the mRNA to be targeted. Moreover, if the gene belongs to a multigene family, it is important to take this parameter into account and select probes with less than 65% homology with the other members of the family. Once the probe has been created, two nucleotides will be added 50 and 30 , respectively, AX and XA (blue bases), where X will be a labeled base and A an adenosine. Y (yellow base): any nucleotide except a thymine; T thymidine, A adenosine, X labeled nucleotide as C6dT; green: any base; blue: added bases “AX” or “XA”; Tm: melting temperature

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3. To label three probes, take 5 μg of each unprotected oligonucleotide, and add 23 μL of 0.1 M carbonate buffer pH 8.8 and 10 μL of DMSO/Cy3 solution per probe. 4. Vortex twice for 30 s and leave overnight (O/N) at room temperature in the dark. 5. Add 10 μg of E. coli tRNA and precipitate with 3.0 M sodium acetate, pH 5.2 (1/10 volume), and 3 vol 100% ethanol. Leave O/N at 20  C, and wash with 80% ethanol. Supernatant and pellet should be red or green according to the fluorochrome used. 6. Resuspend oligo in 100 μL H20/0.3 M sodium acetate. Precipitate again with 3 vol 100% ethanol, and wash with 300 μL 80% ethanol. Repeat this procedure until the supernatant becomes transparent and only the pellet is colored. 7. Resuspend in 250 μL TE (final concentration 20 ng/μL). Adapt volumes to optimize the number of probes to label (5 maximum), for example, with five probes: 3 μg oligonucleotide, 13.8 μL buffer, and 6 μL DMSO/Cy3 per probe. We do not advise to label reduced amount of oligonucleotides to avoid loss of material during precipitation procedure. 3.3 Sample Preparation and Fixation (Fig. 3)

1. Prepare petri dishes with in vitro plant growth medium, suitable for the experimental requirements. Once set, sow sterilized Arabidopsis seeds. 2. Seal the petri dish with two rounds of micropore tape, and store overnight at 4  C for stratification. 3. Transfer the plate to a growth cabinet with 16-h light/8-h dark regime with 25  C/22  C, respectively. 4. Grow the seedlings for the appropriate duration and conditions in which the gene of interest is induced. In parallel, grow negative control seedlings in which the gene of interest is repressed or deleted. 5. Transfer the plants into a small glass dish containing freshly prepared 4% PFA/MES buffer (pH ~5.8 or 7 depending if GFP signal should be conserved), and incubate for 20 min at room temperature in a fume hood (see Note 3). 6. Isolate material to label (here roots but can be other tissue), and remove from the fixative solution. Wash twice with 1
MES buffer. 7. Arrange three to four roots onto a microscopic slide, and cover with a coverslip. Gently squash each root onto the slide using your thumb, and be careful to avoid breaking the coverslip. Aim to splay the roots sufficiently to produce multiple files of isolated cells in a single cell layer.

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Fig. 3 Diagram of the FISH protocol. Fixation: after incubating the seedling in MES and then fixing it in PFA, the seedling is squashed between slide and coverslip before being immersed in liquid nitrogen for a few seconds by using forceps to hold the slide and the coverslip together. The slide is then removed with a scalpel. The slide is immersed in a Coplin jar containing ethanol. At this stage, the slides can be stored for up to 1 week. Hybridization: the slide is washed in MES and then dipped in formamide. After drying, a mix of probes is placed on the seedling and covered with a coverslip, trying to avoid the formation of bubbles as much as possible. The slide is placed in a petri dish containing a container of water to maintain humidity. The petri dish is covered with an aluminum foil to ensure darkness and then incubated overnight at 37  C. Washing/mounting: after removing the coverslip, followed by several formamide baths at 37  C, the mounting medium is applied to the seedling and then covered with a coverslip, again avoiding the formation of bubbles. The object can be observed immediately but can also be stored at 20  C for several years

8. Use tweezers to hold the squashed roots under the coverslip, and immerse each slide in liquid nitrogen for ~5 s. After removal from the nitrogen, ease a razor blade between the coverslip and the slide, and flick the coverslip off. 9. Leave samples to air-dry at room temperature for a minimum of 30 min. To avoid increased levels of autofluorescence, do not leave to dry for longer than 2 h. 10. Permeabilize the samples by immersing the slides into a Coplin jar containing 70% ethanol overnight at 4  C. Fixed roots can be stored at 2–8  C in 70% ethanol up to a week prior to hybridization. 3.4 In Situ Hybridization (Fig. 3)

1. Rinse plants once with MES, and then aspirate the liquid. 2. Incubate the slides in 15% formamide solution (see Notes 3 and 4) for 15 min at room temperature:

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3. During the incubation time, prepare Mixes 1 and 2 separately on ice according Subheading 2.1. 4. A volume of 100 μL (Mix 1 + Mix 2) is sufficient for one slide (22
50 mm). 5. Vortex thoroughly Mix 2. Heat Mix 1 at 85  C for 3 mins in order to denature secondary structures, and then place on ice (see Note 5). 6. Then add Mix 1 to Mix 2 and vortex again. 7. Add 100 μL of the hybridization mix on top of the fixed plants on the slide, and then lay a coverslip on top to prevent evaporation. Be careful to avoid air bubbles. 8. Arrange the slides in a square petri dish. Put a cap of a falcon tube containing some water inside the petri dish to create humidity, and close the petri dish. 9. Wrap a Parafilm sheet around it, and incubate at 37  C overnight. 3.5 Washing and Mounting (Fig. 3)

1. Remove the coverslip and place the slides in a Coplin jar containing freshly prepared 15% formamide (same as in Subheading 3.4). Put the jars on a horizontal shaker for 45 min at 37  C. Repeat this step twice. 2. Rinse the slides twice in 1xMES buffer before mounting. 3. Drop 20 μL of an anti-fading mounting medium containing DAPI on the slide, and lay the coverslip on top (see Note 6). Observations can be performed immediately.

3.6 Imaging and Image Analysis

1. Observe and image plants on a microscope. We use either a spinning disk confocal or a wide-field microscope. For spinning disk microscopy, we use a Dragonfly (Oxford Instruments) equipped with four laser lines and an ultrasensitive EMCCD camera (iXon Life 888, Andor) mounted on a Nikon Eclipse Ti2 microscope body, using a 40
, NA 1.3 Plan Fluor oil objective or a 60
, NA 1.4 Plan Apochromat oil objective coupled with a supplementary lens of 2
, using z-stacks of about 50–80 slices with a 0.5 μm or 0.4 μm step. For widefield imaging, we use a Zeiss Axioimager Z1 wide-field upright microscope equipped with a camera sCMOS ZYLA 4.2 MP (Andor), using a 100
, NA 1.4 Plan Apochromat oil objective. For these z-stacks, a step of 0.3 or 0.4 μm is used. 2. Images properties (e.g., brightness, contrast, colors) can be adjusted using ImageJ (Fig. 4).

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Fig. 4 Imaging SPX1 transcription in fixed plant mature tissues by using smFISH technique. (a) Maximum projection image of mature Col0 root grown on medium without phosphate, a condition necessary for SPX1 expression. The roots were crushed and treated for smFISH with probes hybridized against endogenous SPX1 mRNA. (b) Cropped image of a nucleus from panel a indicated by a white square. Based on the number of transcription sites (green arrows), this nucleus shows polyploidy; single RNAs are visible outside the nucleus (white arrows). The nuclei are stained with DAPI. (c) Negative control, maximum projection image of mature Col0 root grown on medium with 500 μM phosphate, a condition repressing SPX1 expression. The roots were crushed and treated for smFISH with probes hybridized against endogenous SPX1 mRNA Scale bars: 30 μm (panel a and c) or 10 μm (panel b)

3. Quantification of smFISH spots and transcription sites is performed using FISH-quant. Follow the detailed instructions of the developers. Use the negative control plants as basis to determine the background of FISH spots.

4 Notes 1. We also obtained good results with an alternative hybridization buffer (100 mg/mL dextran sulfate and 10% formamide in 2Xspiepr146 SSC). 2. The amount of probes can be increased if the signal is too low or decreased if the background is too high. 3. If the plants used are transgenic and fluorescent, we recommend fixing in 2% paraformaldehyde for 12 min to maintain the fluorophore or replacing formamide with 2.4 M urea [55].

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4. In case of high background, the formamide concentration can be increased from 15% to 50% for both the washes and hybridization buffer (or use 8 M urea). 5. In our hands, the protocol described here works also for smiFISH in plants (but turns out to be not as good as smFISH). For smiFISH, step 5 in Subheading 3.4 for denaturation of secondary probes must be omitted; otherwise, the secondary probes would detach. 6. For long-term storage, the slides can be kept at 20  C for future imaging.

Acknowledgments CM and TD were financed by ANR-18-CE20-023 (BioPhyt) and LN, SH, and PD by ANR-21-CE12-0016 (ULTIM). References 1. Blein T, Balzergue C, Roule´ T et al (2020) Landscape of the noncoding transcriptome response of two Arabidopsis ecotypes to phosphate starvation. Plant Physiol 183(3): 1058–1072 2. Mangiavacchi A, Morelli G, Orlando V (2023) Behind the scenes: how RNA orchestrates the epigenetic regulation of gene expression. Front Cell Dev Biol 11:1123975 3. Chin A, Le´cuyer E (2017) RNA localization: making its way to the center stage. Biochim Biophys Acta Gen Subj 1861(11):2956–2970 4. Tian L, Chou HL, Fukuda M et al (2020) mRNA localization in plant cells. Plant Physiol 182(1):97–109 5. Medioni C, Mowry K, Besse F (2012) Principles and roles of mRNA localization in animal development. Development 139(18): 3263–3276 6. Berleth T, Burri M, Thoma G et al (1988) The role of localization of bicoid RNA in organizing the anterior pattern of the Drosophila embryo. EMBO J 7(6):1749–1756 7. Frigerio G, Burri M, Bopp D et al (1986) Structure of the segmentation gene paired and the Drosophila PRD gene set as part of a gene network. Cell 47(5):735–746 8. Lawrence JB, Singer RH (1986) Intracellular localization of messenger RNAs for cytoskeletal proteins. Cell 45(3):407–415 9. Garner CC, Tucker RP, Matus A (1988) Selective localization of messenger RNA for

cytoskeletal protein MAP2 in dendrites. Nature 336(6200):674–677 10. Kloc M, Zearfoss NR, Etkin LD (2002) Mechanisms of subcellular mRNA localization. Cell 108(4):533–544 11. Palacios IM, Johnston DS (2001) Getting the message across: the intracellular localization of mRNAs in higher eukaryotes. Annu Rev Cell Dev Biol 17(1):569–614 12. Jeffery WR, Tomlinson CR, Brodeur RD (1983) Localization of actin messenger RNA during early ascidian development. Dev Biol 99(2):408–417 13. Le´cuyer E, Yoshida H, Parthasarathy N et al (2007) Global analysis of mRNA localization reveals a prominent role in organizing cellular architecture and function. Cell 131(1): 174–187 14. Long RM, Singer RH, Meng X et al (1997) Mating type switching in yeast controlled by asymmetric localization of ASH1 mRNA. Science 277(5324):383–387 15. Zarnack K, Feldbru¨gge M (2010) Microtubule-dependent mRNA transport in fungi. Eukaryot Cell 9(7):982–990 16. Keiler KC (2011) RNA localization in bacteria. Curr Opin Microbiol 14(2):155–159 17. Chou HL, Tian L, Washida H et al (2019) The rice storage protein mRNAs as a model system for RNA localization in higher plants. Plant Sci 284:203–211 18. Balusˇka F, Salaj J, Mathur J et al (2000) Root hair formation: F-actin-dependent tip growth

smFISH for Plants is initiated by local assembly of profilinsupported F-actin meshworks accumulated within expansin-enriched bulges. Dev Biol 227(2):618–632 19. Okita TW, Choi SB (2002) mRNA localization in plants: targeting to the cell’s cortical region and beyond. Curr Opin Plant Biol 5(6): 553–559 20. Vincent T, Vingadassalon A, Ubrig E et al (2017) A genome-scale analysis of mRNA s targeting to plant mitochondria: upstream AUG s in 50 untranslated regions reduce mitochondrial association. Plant J 92(6):1132–1142 21. Cheng SF, Huang YP, Chen LH et al (2013) Chloroplast phosphoglycerate kinase is involved in the targeting of bamboo mosaic virus to chloroplasts in Nicotiana benthamiana plants. Plant Physiol 163(4):1598–1608 22. Thieme CJ, Rojas-Triana M, Stecyk E et al (2015) Endogenous Arabidopsis messenger RNAs transported to distant tissues. Nat Plants 1(4):1–9 23. Hannapel DJ, Sharma P, Lin T (2013) Phloemmobile messenger RNAs and root development. Front Plant Sci 4:257 24. Ham BK, Lucas WJ (2017) Phloem-mobile RNAs as systemic signaling agents. Annu Rev Plant Biol 68:173–195 25. Kehr J, Kragler F (2018) Long distance RNA movement. New Phytol 218(1):29–40 26. Schena M, Shalon D, Davis RW et al (1995) Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science 270(5235):467–470 27. Mortazavi A, Williams BA, McCue K et al (2008) Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nat Methods 5(7):621–628 28. Wissink EM, Vihervaara A, Tippens ND et al (2019) Nascent RNA analyses: tracking transcription and its regulation. Nat Rev Genet 20(12):705–723 29. Bustin SA (2000) Absolute quantification of mRNA using real-time reverse transcription polymerase chain reaction assays. J Mol Endocrinol 25(2):169–193 30. Rodriguez J, Ren G, Day CR et al (2019) Intrinsic dynamics of a human gene reveal the basis of expression heterogeneity. Cell 176(1): 213–226 31. Yamada T, Akimitsu N (2019) Contributions of regulated transcription and mRNA decay to the dynamics of gene expression. Wiley Interdiscip Rev RNA 10(1):e1508 32. Femino AM, Fay FS, Fogarty K et al (1998) Visualization of single RNA transcripts in situ. Science 280(5363):585–590

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33. Gall JG, Pardue ML (1969) Formation and detection of RNA-DNA hybrid molecules in cytological preparations. Proc Natl Acad Sci 63(2):378–383 34. Edgar BA, Odell GM, Schubiger G (1987) Cytoarchitecture and the patterning of fushi tarazu expression in the Drosophila blastoderm. Genes Dev 1(10):1226–1237 35. Raj A, Peskin CS, Tranchina D et al (2006) Stochastic mRNA synthesis in mammalian cells. PLoS Biol 4(10):e309 36. Hani S, Cuyas L, David P et al (2021) Live single-cell transcriptional dynamics via RNA labelling during the phosphate response in plants. Nat Plants 7(8):1050–1064 37. Paul B, Montpetit B (2016) Altered RNA processing and export lead to retention of mRNAs near transcription sites and nuclear pore complexes or within the nucleolus. Mol Biol Cell 27(17):2742–2756 38. Trcek T, Larson DR, Moldo´n A et al (2011) Single-molecule mRNA decay measurements reveal promoter-regulated mRNA stability in yeast. Cell 147(7):1484–1497 39. Trcek T, Sato H, Singer RH et al (2013) Temporal and spatial characterization of nonsensemediated mRNA decay. Genes Dev 27(5): 541–551 40. Raj A, Van Oudenaarden A (2008) Nature, nurture, or chance: stochastic gene expression and its consequences. Cell 135(2):216–226 41. Chen KH, Boettiger AN, Moffitt JR et al (2015) Spatially resolved, highly multiplexed RNA profiling in single cells. Science 348(6233):aaa6090 42. Shah S, Takei Y, Zhou W et al (2018) Dynamics and spatial genomics of the nascent transcriptome by intron seqFISH. Cell 174(2):363–376 43. Pichon X, Lagha M, Mueller F et al (2018) A growing toolbox to image gene expression in single cells: sensitive approaches for demanding challenges. Mol Cell 71(3):468–480 44. Tsanov N, Samacoits A, Chouaib R et al (2016) smiFISH and FISH-quant–a flexible single RNA detection approach with super-resolution capability. Nucleic Acids Res 44(22):e165– e165 45. Brewer PB, Heisler MG, Hejátko J et al (2006) In situ hybridization for mRNA detection in Arabidopsis tissue sections. Nat Protoc 1(3): 1462–1467 46. Hejátko J, Blilou I, Brewer PB et al (2006) In situ hybridization technique for mRNA detection in whole mount Arabidopsis samples. Nat Protoc 1(4):1939–1946 47. Javelle M, Vernoud V, Rogowsky PM et al (2011) Epidermis: the formation and functions

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of a fundamental plant tissue. New Phytol 189(1):17–39 48. Hamant O, Heisler MG, Jonsson H et al (2008) Developmental patterning by mechanical signals in Arabidopsis. Science 322(5908): 1650–1655 49. Bleckmann A, Dresselhaus T (2016) Fluorescent whole-mount RNA in situ hybridization (F-WISH) in plant germ cells and the fertilized ovule. Methods 98:66–73 50. Bruno L, Muto A, Spadafora ND et al (2011) Multi-probe in situ hybridization to whole mount Arabidopsis seedlings. Int J Dev Biol 55(2):197–203 51. Duncan S, Olsson TS, Hartley M et al (2016) A method for detecting single mRNA molecules in Arabidopsis thaliana. Plant Methods 12(1): 1–10

52. Hanchi M, Thibaud MC, Le´geret B et al (2018) The phosphate fast-responsive genes PECP1 and PPsPase1 affect phosphocholine and phosphoethanolamine content. Plant Physiol 176(4):2943–2962 53. Pichon X, Bastide A, Safieddine A et al (2016) Visualization of single endogenous polysomes reveals the dynamics of translation in live human cells. J Cell Biol 214(6):769–781 54. Imbert A, Ouyang W, Safieddine A et al (2022) FISH-quant v2: a scalable and modular tool for smFISH image analysis. RNA 28(6):786–795 55. Sinigaglia C (2019) A widely applicable ureabased fluorescent/colorimetric mRNA in situ hybridization protocol. Bio-protocol 9(17): e3360–e3360

Chapter 7 Fluorescent In Situ Detection of Small RNAs in Plants Using sRNA-FISH Kun Huang, Blake C. Meyers, and Jeffrey L. Caplan Abstract Plant small RNAs are 21–24 nucleotide, noncoding RNAs that function as regulators in plant growth and development. Colorimetric detection of plant small RNAs was made possible with the introduction of locked nucleic acid probes. However, fluorescent detection of plant small RNAs has been challenging due to the high autofluorescence from plant tissue. Here we report a fluorescent in situ detection method for plant small RNAs. This method can be applied to most plant samples and tissue types and also can be adapted for single-molecule detection of small RNAs with super-resolution microscopy. Key words Small RNAs, FISH, In situ hybridization, Quantification, RNAs, Plant, Autofluorescence, Confocal, Linear spectral unmixing, Spectra, STORM, SIM

1

Introduction The fluorescent in situ hybridization (FISH) technique was first developed in 1982 for localization of DNA sequences in Drosophila [1]. Later, in 1991, FISH was applied in plant tissues for the mapping of two highly repetitive genomic sequence in rye roots [2]. FISH detection later transitioned from DNA to RNA targets, eventually including small RNAs. Plant small RNAs are 21–24 nucleotide, noncoding RNAs that control many aspects of plant growth and development, including microRNAs (miRNAs) [3] and small interfering RNAs (siRNAs) [4]. In situ detection of plant small RNA was first achieved using a digoxigenin (DIG)labeled concatemeric probe of miR172 in Arabidopsis [5]. Single copy miRNA probes containing miRNA and partial miRNA precursor sequences were later used for detection of miR165 and miR166 [6]. Locked nucleic acid (LNA)-modified oligonucleotides exhibit increased hybridization efficiency and specificity toward complementary DNAs and RNAs [7]. In 2006, LNA-modified oligonucleotide probes were used to detect

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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15 miRNAs in mouse embryos [8]. In 2012, this method was modified to localize small RNAs in plant samples with colorimetric detection [9]. Fluorescent detection of small RNA has been challenging. Autofluorescence in plant tissues spans many regions in the visible light spectrum [10, 11]. This goal has been achieved by sRNA-FISH, using multiphoton and spectra unmixing microscopy in combination with LNA-modified probes [12]. sRNA-FISH can also be adapted to super-resolution imaging that detects and localizes single molecules, bypassing the light-diffraction limit [13]. In this chapter, we described a protocol for sRNA-FISH that can be used for fluorescent detection of small RNAs in plants. This method is compatible with single molecule, localization-based super-resolution using direct stochastic optical reconstruction microscopy (dSTORM) and super-resolution structured illumination microscopy (SR-SIM).

2

Materials All solutions are prepared using autoclaved, ultrapure water under room temperature. Diethyl pyrocarbonate (DEPC)-treated water is not suitable for imaging purposes; hence, it should be avoided for this protocol. All reagents should be stored at room temperature unless specified otherwise.

2.1 Sample Preparation for Embedding

1. 16% paraformaldehyde: store at 4 °C. 2. 20 mL glass scintillation vials. 3. 50 mL tubes. 4. Dissecting scope. 5. Ethanol gradient solutions: 10%, 30%, 50%, 70%, 90%, and 100% ethanol. Mix 50 mL, 150 mL, 250 mL, 350 mL, and 450 mL ethanol, and bring to 500 mL with water. Ethanol gradient solutions should be prepared right before use. 6. Ethanol/histoclear gradient series: 75% ethanol/25% histoclear, 50% ethanol/50% histoclear, 25% ethanol/75% histoclear. Prepare right before use. 7. Fixation buffer: 4% paraformaldehyde in 1× PHEM buffer. To prepare, add 20 mL 2× PHEM buffer to 50 mL tubes. Add 10 mL 16% paraformaldehyde, and bring to 40 mL with water. Paraformaldehyde is hazardous; this step should be carried out in a fume hood. 8. Forceps. 9. PBS buffer (10×): 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4. pH 7.4. Weigh 80 g NaCl, 2 g

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KCl, 14.4 g Na2HPO4, and 2.4 g KH2PO4. Add water to 1 L, and adjust pH to 7.4. Autoclave before use. 10. PHEM buffer (2×): 60 mM PIPES, 5 mM HEPES, 10 mM EGTA, 2 mM MgSO4, pH 8. Add 18.14 g of PIPES, 6.5 g of HEPES, 3.8 g of EGTA, and 0.99 g of MgSO4 to 400 mL of water, and bring to a final volume of 500 mL. Autoclave and store solution at 4 °C. 11. Razor blades. 12. Vacuum pump and bell jar. 2.2 Paraffin Embedding

1. 58 °C oven. 2. Ethanol. 3. Forceps. 4. Histoclear. 5. Hot plate. 6. Tissue embedding and processing cassettes. 7. Paraffin wax pellets.

2.3 Sample Preparation for Hybridization

1. #0 watercolor paint brushes. 2. 37 °C incubator. 3. Enzyme solution: 50 mg/mL protease. Dissolve 0.5 g protease in 10 mL water. Predigest the solution at 37 °C for 4 h. Store aliquots at -20 °C. 4. Ethanol gradient solutions: 100% (two of them), 95%, 80%, 70%, 50%, 30%, 10%, and 0% ethanol. Mix 500 mL, 475 mL, 400 mL, 350 mL, 250 mL, 150 mL, 50 mL, and 0 mL ethanol with 0 mL, 25 mL, 100 mL, 150 mL, 250 mL, 350 mL, and 450 mL water. 5. Glass staining dish. 6. Glycine solution: 0.2% glycine. To prepare, add 5 mL 10% glycine to 245 mL PBS buffer (1×). 7. Histoclear. 8. Paraffin microtome. 9. Slide holder. 10. Slide warmer. 11. Superfrost glass slides. 12. TE buffer: 10 mM Tris-HCl, 1 mM disodium EDTA, pH 8.0. Add 100 mL 100 mM Trish-HCL and 100 mL 10 mM ethylenediaminetetraacetic acid (EDTA). Bring it up to 1 L with water. 13. TEA buffer: 1.3% (v/v) triethanolamine, 0.4% (v/v) HCl, and 0.5% (v/v) acetic anhydride. Add 5.2 mL of triethanolamine

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and 1.6 mL of HCl to 393 mL of water. Add 2 mL of acetic anhydride right before use. TEA buffer should be prepared in the chemical fume hood. 14. TE-protease solution: 130 μg/mL protease in 1xTE buffer. To prepare, pre-warm 250 mL TE buffer to 37 °C. Add 659 μL enzyme solution right before use. 15. Water. 2.4 In Situ Hybridization

1. Denhardt’s solution: 50×. 2. Dextran sulfate solution: 50% dextran sulfate. Add 5 g dextran sulfate to 7 mL water. Heat the solution at 80 °C for until completely dissolve. Bring the volume to 10 mL with water. Mix well and store 2.5 mL aliquots at -20 °C. 3. Digoxigenin NHS-ester (DIG)-labeled probes. 4. Flat-bottom slide container. 5. Formamide: deionized, store at 4 °C. 6. Heating block for 1.5 mL tubes. 7. Hybridization oven. 8. Hybridization salts: 3 M NaCl, 100 mM Tris-HCl (pH 8.0), 100 mM sodium phosphate (pH 6.8) and 50 mM EDTA; store aliquots at -20 °C. 9. Hybridization solution: 12.5% hybridization salt (v/v), 5% Dextran sulfate (w/v), 1.25 ug/uL tRNA, 50% formamide, and 2.5% of 50% Denhardt’s solution (v/v). To prepare, add 1.25 mL of hybridization salts, 5 mL of deionized formamide, 2.5 mL of 50% dextran sulfate, 250 μL of 50× Denhardt’s solution, and 125 μL of tRNA solution to 875 μL water. Filter sterilize and store 1 mL aliquots at -20 °C. 10. Membrane hybridization cover slips. 11. Parafilm. 12. Plastic wrap. 13. SSC buffer (0.2×): Add 10 mL SSC buffer (20×) to 990 mL water. 14. SSC buffer (20×): 3 M NaCl, 0.3 M sodium citrate, pH 7.0. To prepare, add 175.3 g of NaCl and 88.2 g of sodium citrate to 800 mL water. Adjust pH to 7.0 and bring up the volume to 1 L. 15. TBS buffer (1×): 50 mM Tris-Cl, 150 mM NaCl, pH 7.5. Add 100 mL 500 mM Tris-Cl, 30 mL 5 M NaCl to water, and bring it up to 1 L. 16. tRNA solution: 100 mg/mL tRNA. To prepare, add 1 g tRNA to 1 mL water. Mix and dissolve, store at -20 °C.

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1. Blocking buffer: 1% blocking reagent in TBS (1×) buffer. Add 1 g of blocking reagent to 70 °C TBS (1×) buffer, stir till cooled down to room temperature. 2. Blocking reagent. 3. Bovine serum albumin (BSA). 4. Flat-bottom dishes. 5. Primary anti-DIG antibody. 6. Secondary antibody. 7. TBS buffer (1×): 50 mM Tris-Cl, 150 mM NaCl, pH 7.5. Take 100 mL of 500 mM Tris-Cl and 30 mL 5 M NaCl, and dilute it to 1 L. 8. Triton X-100. 9. Washing buffer: 1% (wt/vol) BSA and 0.3% (vol/vol) Triton X-100. Add 8 g BSA to 800 mL TBS buffer (1×). Add 240 μL Triton X-100. Stir till the solution turns clear.

2.6 Sample Wash and Mount

1. Clear nail polish. 2. Cover glasses. 3. Dish shaker. 4. Glass dishes. 5. Mounting medium. 6. Washing buffer: 1% (wt/vol) BSA and 0.3% (vol/vol) Triton X-100. To prepare, add 8 g BSA to 800 mL of TBS buffer (1×). Add 240 μL of Triton X-100. Stir till the solution turns clear.

2.7

Imaging

1. Confocal microscopes. 2. dSTORM imaging buffer contains three buffers and should be mixed freshly before use: Buffer A (30 nM Tris/Cl pH 8.5, 1 mM EDTA, 6.25 uM glucose oxidase, and 2.5 μM catalase), Buffer B (250 mM cysteamine-HCL, pH 3), and Buffer C (250 mM glucose). 3. Software for imaging process. 4. Super-resolution microscopes.

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3.1 Probe Design and Preparation

For each sRNA target, a probe is designed by adding LNA bases to the reverse complementary sequence. Usually 6–9 LNA bases are ideal for a 20–22 nt sRNA target. Designed probes can be ordered through the QIAGEN website: https://www.qiagen.com/us/ products/discovery-and-translational-research/custom-lnaoligonucleotides.

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1. Reverse complement the target sRNA sequence. 2. Calculate the melting temperature of the reverse complemented sequence. The backbone sequence chosen for probe design should have Tm ≤ 60 °C. If the Tm ≥ 60 °C, the backbone length can be reduced to 18–19 nt. 3. Manually introduce LNA bases to the backbone. Avoid stretches of more than four LNA bases. Avoid stretches of more than three Gs or Cs. 4. Analyze the Tm, secondary structure, and dimer tendencies of the oligo, and look for probe with highest Tm and lowest tendencies to form dimers and secondary structures. 5. Probes should be diluted to 100 μM or 250 μM concentration and stored as aliquots at -80 °C. 3.2 Sample Dissection and Fixation

1. Use forceps and razor blade to dissect the sample under a dissecting microscope. Sample size could be 1 mm to 1 cm in diameter. 2. Prepare fixation buffer right before dissecting. Add 10 mL fixation buffer to each 20 mL glass scintillation vial, and place sample in the fixation buffer immediately after dissecting. 3. Apply 0.1 mPa vacuum to the sample in a vacuum bell jar. Vacuum for 15 min and then release the pressure. Gently tap and swirl the sample. Repeat this step three to five times until the sample sinks into the fixation buffer and to the bottom of the vial (see Note 1). 4. Store fixed samples overnight at 4 °C.

3.3 Paraffin Embedding and Slides Preparation

1. Rinse fixed sample three times with PBS (1×) buffer, 30 min each time. 2. Dehydrate the sample by applying ethanol gradient series, 30 min each step. 3. Incubate the sample in 100% ethanol twice, 30 min each time. 4. Treat the sample with ethanol/histoclear gradient series, 1 h each step. 5. Immerse the sample in histoclear three times, 1 h each time. 6. Discard the histoclear. Add 5 mL fresh histoclear to the sample, and add 15 mL of wax pellets to fill the jar. Incubate in a 58 °C oven overnight. 7. Add in fresh wax pellets every hour, till the scintillation vials are completely filled. 8. Replace wax with fresh melted wax every 3 h. 9. Repeat Step 8 four times. 10. Transfer the processed samples into tissue embedding and processing cassettes on a hot plate. Slightly cool at room

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temperature while holding the tip of the sample with forceps in position. 11. Completely cool the sample to room temperature, and store at 4 °C. Embedded samples can be stored at 4 °C up to 6 months. 12. Trim paraffin embedded samples with razor blades. A trapezoid shape is preferred for section. 13. Section the sample to a 6–10 μm thickness. Transfer two to four sections onto a glass slide using paint brushes. 14. Add water beneath the section so that the sections are floating on the water. 15. Completely stretch the section by incubation on a 37 °C slide warmer for 2 h. 16. Discard the water, and dry the slides on the 37 °C slide warmer for 2 days. Dried slides can be stored at 4 °C for up to 1 week. 3.4 In Situ Hybridization

1. Start a 37 °C incubator and a heating block at 90 °C. 2. Deparaffinize by incubating the slides in 100% histoclear solution twice, 10 min each time. 3. Rehydrate the slides by immersing in ethanol gradient solutions, 1 min each step. 4. Rinse the slides in PBS buffer (1×) twice, 2 min each time. 5. Digest the slides in TE-protease solution for 20 min in a 37 °C incubator (see Note 2). 6. Rinse the slides in glycine solution for 10 min at room temperature. 7. Rinse the slides in PBS buffer (1×) twice, 2 min each time. 8. Incubate the slides in TEA buffer for 10 min. This step is optional and can be omitted for LNA probes. 9. Dehydrate the slides in ethanol gradient solution, 1 min each step. 10. Store slides in 100% ethanol at 4 °C for at least 2 h. Slides can be stored in 100% ethanol at 4 °C for up to 1 week. 11. Warm up slides to room temperature. Air-dry the slides for 5 min. 12. Prepare hybridization probe mix by mixing 1 μL probe (see Note 3), 9 μL formamide, and 10 μL water. Denature the probe mix by incubating at 90 °C for 3 min, and chill on ice immediately after incubation. 13. Prepare hybridization mix by adding 80 μL hybridization solution to the probe mix. 14. Apply 100 μL hybridization probe mix to each slide.

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Fig. 1 Assembly for in situ hybridization. A moist chamber is made by sandwiching wet paper between a flat-bottom container and two overlapped pieces of Parafilm with corners cut. Lay the slides flat on top of the Parafilm for overnight hybridization

15. Gently cover the slides with membrane hybridization cover slips. 16. Make a hybridization chamber (Fig. 1) by laying wet paper in a flat-bottom slide container. Cut two corners of paraffin film, and lay on top of the wet paper. 17. Hybridize in a hybridization oven overnight at 53 °C (see Note 4). 18. Wash the slides with SSC buffer (0.2×) twice at 53 °C, 1 h each time. 19. Prepare blocking buffer and washing buffer. 20. Lay the slides flat in a flat-bottom slide container, and apply 100 mL blocking buffer. Gently agitate for 45 min. 21. Wash the slides with washing buffer (1×) for 45 min with gentle agitation. 3.5 Antibody Detection

1. Prepare the antibody solution for the primary antibody by diluting the primary anti-DIG antibody in washing buffer (1×) (see Note 5). 2. Decant the washing buffer, apply 100 μL antibody solution containing primary antibody, and incubate overnight at 4 °C.

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3. The next day, wash the slides with washing buffer (1×) for three times, 20 min each time. 4. Prepare the antibody solution for the secondary antibody by diluting the secondary antibody in washing buffer (1×) (see Note 5). 5. Apply 100 μL secondary antibody to each slide, and incubate overnight at 4 °C. 6. The next day, wash the slides with washing buffer (1×) for three times, 20 min each time. 7. Wash the slides in PBS buffer (1×) for 5 min. 3.6 Slide Mounting, Imaging, and Imaging Processing

1. For confocal microscopy and SR-SIM, slides can be mounted with anti-fade mounting medium and sealed with nail polish. 2. For super-resolution microscopy, slides can be stored in PBS buffer (1×) until imaging and then mounted with imaging solution, such as dSTORM imaging buffer, during image acquisition. 3. Image mounted slides with a laser scanning confocal microscope or super-resolution microscope. 4. If spectra of the samples need to be acquired for distinguishing true signal from autofluorescence background, slides should be imaged with a laser scanning confocal microscope with spectra unmixing capability. 5. Process images with imaging processing software (see Note 6) (Fig. 2).

Fig. 2 Example of sRNA-FISH demonstrating localization of miR2275 (red-to-white dots) in premeiotic maize anthers using dSTORM. (a) miR2275 was detected in the tapetal layer and archesporial cells. TA, tapetal layer; MI, meiocyte. (b) Higher magnification images of boxed area showing localization around the nucleus (Nu). Nuclei were counterstained with DAPI (4′,6-diamidino-2-phenylindole)

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Notes 1. Fixation is critical for successful detection of sRNAs. Good fixation is evident by full and efficient penetration of fixation buffer in the tissue. Some plant samples are extremely difficult to fix. In this case, samples can be stored in a vacuum bell jar overnight at 4 °C. Samples can also be placed on a rotating platform overnight at 4 °C. For large samples, fixative needs to be changed three times for a better fixation result. 2. Protease concentration and digestion time are determined by sample thickness and desired signal intensity. Start with 130 μg/mL protease and a digest time of 20 min. If the signal is too weak, increase the protease concentration, and/or extend the digestion time. If nonspecific signal is observed, decrease the protease concentration and/or shorten the digestion time. 3. Optimal probe concentration is determined based on sample type and background fluorescence. Start with 1 ng/μL probe working concentration. Increase the probe concentration if weak signal is observed. Probe can be used up to 2.5 μM without causing background issue in most plant samples. 4. Ideal hybridization temperature is determined by the melting temperature (Tm) of each LNA probe. Generally, LNA probes with a Tm around 80 °C should use a hybridization temperature between 45 °C and 60 °C. 5. Optimal antibody concentration is determined experimentally. To start, two antibody concentration combinations can be used: 0.2 μg/mL and 1 μg/mL for the primary antibody and 1:100 and 1:1000 dilution for the secondary antibody. The optimal antibody concentration is determined by positive controls that show bright and specific signals. 6. Images acquired with Lambda mode should be spectrally unmixed using Zen (Carl Zeiss) or any other spectra unmixing software, with the pure fluorophore spectrum as a positive control and unstained tissue spectrum as a negative control. Super-resolution images should be processed with Zen PALM processing or any other super-resolution images processing package, such as ThunderSTORM [14] in ImageJ.

Acknowledgments This project was supported by the US NSF Plant Genome Research Program, awards 1649424, 1611853, and 1754097. We would like to thank members of the Meyers and Caplan labs for help and

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support. Microscopy access was supported by grants from the NIH-NIGMS (P20 GM103446, P20 GM139760) and the State of Delaware. References 1. Langersafer PR, Levine M, Ward DC (1982) Immunological method for mapping genes on Drosophila polytene chromosomes. Proc Natl Acad Sci USA 79(14):4381–4385 2. Leitch IJ, Leitch AR, Heslopharrison JS (1991) Physical mapping of plant DNA-sequences by simultaneous in situ hybridization of 2 differently labeled fluorescent-probes. Genome 34(3):329–333 3. Reinhart BJ, Weinstein EG, Rhoades MW, Bartel B, Bartel DP (2002) MicroRNAs in plants. Gene Dev 16(13):1616–1626 4. Hamilton AJ, Baulcombe DC (1999) A species of small antisense RNA in posttranscriptional gene silencing in plants. Science 286(5441): 950–952 5. Chen XM (2004) A microRNA as a translational repressor of APETALA2 in Arabidopsis flower development. Science 303(5666): 2022–2025 6. Kidner C, Timmermans M (2006) In situ hybridization as a tool to study the role of microRNAs in plant development. Methods Mol Biol 342:159–179 7. Kaur H, Arora A, Wengel J, Maiti S (2006) Thermodynamic, counterion, and hydration effects for the incorporation of locked nucleic acid nucleotides into DNA duplexes. Biochemistry 45(23):7347–7355

8. Kloosterman WP, Wienholds E, de Bruijn E, Kauppinen S, Plasterk RHA (2006) In situ detection of miRNAs in animal embryos using LNA-modified oligonucteotide probes. Nat Methods 3(1):27–29 9. Javelle M, Timmermans MCP (2012) In situ localization of small RNAs in plants by using LNA probes. Nat Protoc 7(3):533–541 10. Berg RH (2004) Evaluation of spectral imaging for plant cell analysis. J Microsc 214:174– 181 11. Harter K, Meixner AJ, Schleifenbaum F (2012) Spectro-microscopy of living plant cells. Mol Plant 5(1):14–26 12. Huang K, Baldrich P, Meyers BC, Caplan JL (2019) sRNA-FISH: versatile fluorescent in situ detection of small RNAs in plants. Plant J 98(2):359–369 13. Galbraith CG, Galbraith JA (2011) Superresolution microscopy at a glance. J Cell Sci 124(10):1607–1611 14. Ovesny M, Krizek P, Borkovec J, Svindrych Z, Hagen GM (2014) ThunderSTORM: a comprehensive ImageJ plug-in for PALM and STORM data analysis and super-resolution imaging. Bioinformatics 30(16):2389–2390

Chapter 8 hcHCR: High-Throughput Single-Cell Imaging of RNA in Human Primary Immune Cells Manasi Gadkari, Jing Sun, Adrian Carcamo, Iain Fraser, Luis M. Franco, and Gianluca Pegoraro Abstract Functional genomics and chemical screens can identify and characterize novel cellular factors regulating signaling networks and chemical tools to modulate their function for the treatment of disease. Screening methods have relied primarily on immortalized and/or transformed cancer cell lines, which can limit the generalization of results to more physiologically relevant systems. Most have also relied on immunofluorescence, or on stably expressed recombinant fluorescent proteins, to detect specific protein markers using high-content imaging readouts. In comparison, high-throughput methods to visualize and measure RNA species have been less explored. To address this, we have adapted an isothermal signal amplification chemistry for RNA FISH known as hybridization chain reaction (HCR) to an automated, high-content imaging assay format. We present a detailed protocol for this technique, which we have named high-content HCR (hcHCR). The protocol focuses on the measurement of changes in mRNA abundance at the singlecell level in human primary cells, but it can be applied to a variety of primary cell types and perturbing agents. We anticipate that hcHCR will be most suitable for low- to medium-throughput screening experiments in which changes in transcript abundance are the desired output measure. Key words RNA, Gene expression, Methods, Fluorescent in situ hybridization, Fluorescence microscopy, High-content imaging, Immunology

1 Introduction Chemical or functional genomics screens are important tools for dissecting signaling networks in biological systems [1] and are central to the drug discovery process [2]. In medium- or highthroughput screens, the cellular effects of large collections of perturbing agents (chemical compounds, RNAi, or CRISPR/Cas9) are tested in a single experiment on up to thousands of cells per

Authors Manasi Gadkari and Jing Sun share the first-author position. Authors Luis M. Franco and Gianluca Pegoraro share the senior-author position. Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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condition. High-content imaging (HCI) assays employ automated liquid handling, image acquisition, and image analysis [3, 4] to measure cellular phenotypic changes and are therefore suitable for medium- or high-throughput screens. HCI and other screening assays have relied primarily on immortalized and/or transformed cancer cell lines, which can limit the generalization of results to more physiologically relevant systems [5–8]. In addition, HCI assays generally rely on the detection of signal from fluorescent dyes, fluorescently labeled antibodies (immunofluorescence), or stably expressed fluorescent proteins, to visualize and quantify nucleic acids, cellular membranes, or protein markers. Assays to visualize and measure specific mRNA expression with HCI have been described [9], but methods in this area have been generally lagging behind protein-based fluorescent markers. Advances in single-molecule RNA fluorescence in situ hybridization (smRNA FISH) have greatly increased the sensitivity of detection of individual RNA molecules [10–12]. However, quantitative detection of single RNA molecules depends on high spatial resolution and the use of Nyquist sampling criteria, with high magnification objectives, large z-stacks of images, and single-plane image analysis, which precludes the application of smRNA FISH in the highthroughput format of chemical or functional genomics screens. To overcome these limitations, we have developed hcHCR, a high-throughput imaging-based method for measuring changes in gene expression at the single-cell level in human primary cells [13]. This method combines HCI with an existing isothermal signal amplification chemistry for RNA FISH known as hybridization chain reaction (HCR) (Fig. 1) [14, 15]. HCR amplification permits the use of lower magnification objectives to visualize mRNA transcripts, thus allowing imaging of larger fields of view (FOV) with hundreds of cells per field, making it compatible with automated HCI instruments. This chapter describes the hcHCR in detail. The major steps of an hcHCR experiment (Fig. 2) are as follows: 1. Blood collection, cell purification, and cell plating 2. Treatment of the cells with one or more perturbing agents and controls 3. Cell fixation and permeabilization 4. Primary probe reaction (HCR)

hybridization

and

hybridization

chain

5. High-content imaging (HCI) acquisition and analysis Although we developed and describe the protocol for use with human primary immune cells, the cell purification and plating step (Subheading 3.2) can be optimized for work with other human primary cell types or with cells from other species. Similarly,

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Fig. 1 Overview of the HCR fluorescence amplification reaction (Adapted from [14]). mRNA in fixed and permeabilized cells is hybridized with DNA oligo probe pair sets that carry a split HCR initiator sequence. Each HCR probe set is composed of >10 probe pairs that hybridize to different regions of the target gene (not shown). The probes in each of the pairs bind the mRNA target next to each other, thus bringing the split initiator sequence in close proximity. When in close proximity, the two halves of the split initiator can trigger the unfolding of one of the two fluorescently labeled metastable HCR hairpins (h1), which opens up and exposes an initiator sequence for the other hairpin (h2), which opens up and exposes the initiator sequence for h1. A combination of h1 and h2 hairpins constitutes an HCR amplifier. The unfolding and hybridization process of the HCR amplifiers in the presence of an initiator sequence is isothermal and autocatalytic and results in the local accumulation of a large number of fluorescently labeled molecules at the location of the mRNA target

Fig. 2 hcHCR workflow. The schematic depicts the experimental workflow steps necessary for hcHCR. Peripheral blood mononuclear cells (PBMCs) are obtained from donors’ blood (1), and the desired cell subset is purified from PBMCs by immunomagnetic selection (2). Purified cells are then seeded in 384-well imaging plates (3) and treated with a perturbing agent (4). After treatment, cells are fixed and permeabilized directly on the plate (5) and then stained with RNA HCR (6) so that mRNA transcript expression can be detected and quantified in an automated manner using high-content imaging (HCI) instruments and software

while here we describe one perturbing agent as a specific example in the cell treatment step (Subheading 3.3), hcHCR is compatible with any chemical or functional genomics perturbation appropriate for the cell type selected.

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Materials

2.1 Blood Collection, Monocyte Purification (Adapted from STEMCELL Technologies Protocol), Cell Plating, and LPS Treatment (See Note 1)

1. Peripheral blood collected in Vacutainer EDTA tubes [Becton Dickinson]. 2. SepMate tubes (STEMCELL Technologies). 3. Ficoll-Paque PLUS density gradient medium (GE Healthcare Life Sciences). 4. Cell culture medium: 10% fetal bovine serum (FBS) in RPMI 1640. Sterilize by passing the medium through a 0.22 μm filter. 5. EasySep Human CD14 Positive Selection Kit II (STEMCELL Technologies). Other methods of cell purification can be used. 6. EasySep magnet “The Big Easy” (STEMCELL Technologies). 7. Countess automated cell counter (Invitrogen). Any other automated or manual cell counting method can be used. 8. Resuspension buffer: 2% BSA in 1X phosphate buffered saline (PBS). Filter and sterilize by passing the buffer through a 0.22 μm filter. 9. PhenoPlate 384-well microplates coated with poly-D Lysine (PDL) (Revvity). 10. 1 mg/mL lipopolysaccharide (Enzo, LPS from Salmonella minnesota R595, Re mutant) (see Note 2).

2.2 Cell Fixation and Permeabilization

1. 16% paraformaldehyde (PFA). If using an automated liquid handler, prepare the necessary volume. We use a BlueCatBio BlueWasher automated liquid handler, and prepare 40 mL of 16% PFA for one 384-well microplate (see Note 3). 2. 1X PBS. 3. 70% ethanol. 4. Aluminum adhesive sealing film.

2.3 Primary DNA Oligo Probe Set Hybridization and Hybridization Chain Reaction (Adapted from [14])

1. Saline-sodium citrate Tween-20 (SSC-T) buffer: 5X SSC, 0.1% Tween-20. 2. Probe-hybridization buffer: 30% formamide, 5X SSC, 9 mM citric acid pH 6.0, 0.1% Tween-20, 50 μg/mL heparin, 1X Denhardt’s solution, 10% dextran sulfate. Prepare 10 mL for a full 384-well plate. 3. HCR primary DNA oligo probe sets (see Subheading 3.1). 4. Fluorescently labeled HCR amplifiers that match the initiator sequences present on the primary DNA oligo probe sets (see Subheading 3.1). 5. TE buffer: 10 mM Tris-HCl pH 8.0, 0.1 mM EDTA.

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6. 30% probe wash buffer: 30% formamide, 5X SSC, 9 mM citric acid (pH 6.0), 0.1% Tween-20, 50 μg/mL heparin. Prepare 40 mL per plate. 7. Amplification buffer (Molecular Instruments): 5X SSC, 0.1% Tween-20, 10% dextran sulfate. Prepare 10 mL for a full 384-well plate. 8. 2.5 mg/mL 40 ,6-diamidino-2-phenylindole (DAPI) stock solution. Aliquot into 0.5 mL Eppendorf microcentrifuge tubes, cover the tubes with aluminum foil (DAPI is lightsensitive), and store at 20  C for up to 1 year. The working DAPI solution should be prepared fresh on the day of the experiment by diluting the stock solution 1:1000 in PBS (2.5 μg/mL). 2.4 High-Content Imaging (HCI) Acquisition and Analysis

1. Automated fluorescence microscope (widefield or spinning disk confocal microscope) equipped, at a minimum, with: (a) Controlling software for programmable and automated image acquisition. (b) Motorized x, y, and z stage. (c) Autofocus mechanism (preferably hardware-based, i.e., near-IR laser or LED). (d) Excitation light sources in four fluorescence channels that match the fluorescently labeled amplifiers (e.g., 405 nm, 488 nm, 561 nm, and 640 nm lasers). (e) Matched dichroic and bandpass emission mirrors that match the fluorescently labeled amplifiers. (f) High NA (numerical aperture) 40X or 60X objectives. (g) One or more sCMOS cameras. 2. Commercial or, preferably, open-source software (e.g., Python [16], ImageJ [17], Julia [18], CellProfiler [19, 20]) to perform image analysis on large datasets of images generated by the automated microscope. 3. Open-source scientific computing software for statistical analysis and visualization of single-cell data generated by the image analysis pipelines. R [21], Python, or Julia can be used for this purpose.

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Methods

3.1 HCR Probe Sets and Fluorescent Amplifiers Design and Ordering

We use the Molecular Instruments ordering web page to order both probe sets and fluorescent HCR amplifiers. Probe sets are either custom designed by the vendor, in which case the user needs to provide the organism species and RNA target sequence,

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or picked from a menu of predesigned probe sets for already available genes. A choice of fluorophores is available for the amplifiers (e.g., Alexa546, Alexa488, and Alexa647, among others), based on the availability of light sources, dichroic mirrors, and emission filters on the automated microscope of choice. For multiplexed experiments (more than one RNA molecule measured in each cell), it is important to remember that probe sets targeting each gene, and their corresponding amplifiers, should be selected to have a different combination of split initiators (One out of the ten different initiator sequences: I1–I10) [14], initiator-matched orthogonal amplifiers (one out of the ten different amplifiers: B1– B10) [15], and fluorophores. Below is a specific example of the set of choices for ordering a probe set for the human TNF mRNA: 1. Organism: human (Homo sapiens) 2. Probe set: TNF 3. RefSeq Accession: NM_000594.3 4. Probe set size: 20 probe pairs 5. For use with HCR amplifier: B1 6. Probe set synthesis scale: 200 pmol 7. HCR amplifier: B1 8. Amplifier label: AlexaFluor488 9. Amplifier synthesis scale: 1000 pmol 3.2 Blood Collection, Monocyte Purification, and Cell Plating (See Note 1)

1. Collect peripheral blood from human donors in Vacutainer EDTA tubes (see Note 4). 2. Proceed immediately to isolate peripheral blood mononuclear cells (PBMCs) by gradient centrifugation in SepMate tubes with density gradient medium (Ficoll-Paque PLUS). 3. Add the Ficoll-Paque PLUS density gradient medium to the SepMate tube by carefully pipetting it through the central hole of the SepMate insert. For a 50 mL SepMate tube and an initial blood sample volume of 4–17 mL, add 15 mL of density gradient medium to the SepMate tube. The top of the density gradient medium will be above the insert (see Note 5). 4. Dilute the blood sample with an equal volume of 1X PBS. Mix gently by pipetting up and down. 5. Keeping the SepMate tube vertical, add the diluted sample by pipetting it down the side of the tube. The sample will mix with the density gradient medium above the insert. 6. Centrifuge the tube at 1200  g for 10 min at room temperature (RT), with the brake on (see Note 6). 7. After centrifugation, pour off the top layer, which contains the enriched mononuclear cells (MNCs), into a fresh tube. Do not

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hold the SepMate tube in the inverted position for longer than 2 s (see Notes 7–9). 8. Wash the enriched MNC population with 1X PBS by centrifuging at 300  g for 8 min at RT, with the brake on. 9. Wash once more to remove residual platelets from the enriched MNC population. Perform this wash at 120  g for 10 min at RT with the brake off (see Note 10). 10. Count live cells on the Countess automated cell counter by diluting 10 μL of cells in 10 μL of trypan blue. Other manual or automated cell counting methods and instruments can be used, depending on availability. 11. After PBMC isolation, do one of the following: (a) Proceed immediately to the immunomagnetic enrichment step for the specific cell subset (monocytes, in this case). (b) Alternatively, incubate PBMCs in 10 mL of RPMI 1640 + 10% FBS at 4  C in a 50 mL conical tube, and isolate monocytes the next day. On the day of the isolation, bring up the volume to 50 mL with 1X PBS, and then centrifuge at 500  g for 5 min at RT. Finally, decant the supernatant, and resuspend the PBMC pellet in the desired volume of resuspension buffer, to a final concentration of 1  108 cells/ml for monocyte isolation. 12. Place the PBMCs in a 14 mL polystyrene tube. This procedure is used for processing a sample volume ranging from 250 μL to 8 mL. If starting with smaller sample volumes, bring up the volume to 250 μL, or for larger volumes, prepare multiple sample isolations. 13. Add the EasySep Positive Selection Cocktail at 100 μL/mL of sample. Mix well, and incubate for 10 min at RT. 14. Vortex the RapidSpheres for 30 s to ensure that they are in a uniform suspension. Add the RapidSpheres at 100 μL/mL of sample (e.g., for 1 mL of cell suspension, add 100 μL of RapidSpheres). Mix well and incubate at RT for 3 min. 15. Bring the cell suspension to a total volume of 5 mL (for a sample volume < 2 mL) or 10 mL (for a sample volume > 2 mL) by adding resuspension buffer. Mix the cells in the tube by gently pipetting up and down 2 to 3 times. Place the tube (without cap) into the EasySep magnet, and incubate on a flat surface for 3 min without moving. 16. Gently pick up the EasySep Magnet, and, in one continuous motion, invert the magnet and tube pouring off the supernatant fraction. The magnetically labeled cells will remain inside the tube, held there by the magnetic field of the EasySep

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magnet. Leave the magnet and the tube in inverted position for 2–3 s, and then return to upright position. Do not shake or blot off any drops that may remain hanging from the mouth of the tube. Discard the supernatant. 17. Remove the tube from the magnet, and repeat Steps 15 and 16 two more times, for a total of 3  3-min separations with the magnet. 18. Remove the tube from the magnet, and resuspend cells in culture medium to a concentration of 100,000 cells per 30 μL. This concentration was experimentally optimized to obtain enough cells per field of view (FOV) at the time of imaging. The positively selected cells are now ready for use. 19. Plate the purified cells at 100,000 cells in 30 μL/well of culture medium in a PDL-coated 384-well imaging plate. The 30 μL/ well volume was chosen so that the total volume of cells, plus LPS (30 μL/well), plus PFA 16% fixative (20 μL/well), was below 80 μL/well (see Subheadings 3.2 and 3.3). Seed at least four wells for each experimental condition. In addition, remember to add at least four wells for each of the negative HCR controls (see Note 11). 20. Let the cells rest in the 384-well plate for 1 h at 37  C and 5% CO2 before proceeding with the experimental treatment. This incubation is necessary to reduce the amount of basal cellular stress associated with cell plating. 3.3 Treatment of the Cells with LPS

The following steps describe the activation monocytes purified from healthy donors with LPS. LPS is known to increase mRNA expression of TNF in monocytes [22, 23]. 1. Prepare the 2X LPS working solutions. (a) In a 1.5 mL sterile Eppendorf microcentrifuge tube, add 2 μL of the 1 mg/mL LPS stock to 198 μl of culture medium, and then vortex, to get a 10 μg/mL LPS solution. (b) In a new 1.5 mL sterile Eppendorf microcentrifuge tube, add 20 μL of the 10 μg/mL LPS solution to 980 μL of culture medium, and then vortex, to get the 2X 100 ng/ mL LPS solution (i.e., 200 ng/mL, to be diluted later to a final concentration of 100 ng/mL). (c) Starting from the 2X 100 ng/mL LPS solution, make additional serial 1:10 dilutions (add 100 μL of each 2X solution to 900 μL of culture medium) to get the following working solutions: 2X 10 ng/mL, 2X 1 ng/mL, 2X 0.1 ng/mL, 2X 0.01 ng/mL. 2. Add 30 μL/well of the LPS 2X working solutions to the 30 μL of cells in culture medium, in the corresponding wells of the

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384-well plate. The final concentrations of LPS in the media will be 100, 10, 1, 0.1, and 0.01 ng/mL, respectively. 3. Add 30 μL/well culture medium without LPS to the negative control wells. 4. To study the kinetics of induction of TNF after LPS treatment, the LPS is added at 120 min, 60 min, 30 min, and 15 min before cell fixation (see Note 12). 3.4 Cell Fixation and Permeabilization

1. For use with a BlueWasher plate dispenser/washer, prepare 40 mL/plate of 16% paraformaldehyde (PFA) in PBS. 2. Prime the tubing with 16% PFA. 3. At the end of the LPS treatment timepoints (see Note 12), dispense 20 μL/well of 16% PFA directly to the 60 μL of cells in culture medium, with or without LPS treatment, for a final concentration of 4% PFA (see Note 13). 4. Incubate the plate for 15 min at RT. 5. Use the BlueWasher magbeads wash program plus decant to wash the plate three times with 50 μL/well of 1X PBS. Leave the plate empty after the final wash (see Note 14). 6. To permeabilize the cells, add 40 μL of 70% ethanol, seal the plate with the adhesive aluminum seal, and store it at 20  C overnight (see Note 15).

3.5 Primary DNA Probe Set Hybridization and HCR (Adapted from [14]) 3.5.1 Pre-Hybridization and Primary Probe Hybridization

1. Warm the probe hybridization buffer to 37  C. 2. Aspirate the permeabilization buffer, and air-dry the plate for 10 min at RT to remove the residual ethanol. 3. Rehydrate the plate by adding 80 μL/well of 5X SSC-T buffer using a multichannel pipette, incubating the plate for 5 min at RT, and then by aspirating the buffer. 4. Repeat Step 3 twice to wash the plate. 5. Add 16 μL/well of prewarmed probe-hybridization buffer, and incubate for 10 min at 37  C to equilibrate the plate. During the pre-hybridization incubation, prepare the master mix containing all the primary HCR probe sets for multiplexing in prewarmed hybridization buffer in 1.5 mL Eppendorf microcentrifuge tubes. Prepare a volume of master mix equivalent to at least 120% of the volume to be used in the experiment to account for pipetting errors and losses. The hybridization buffer is viscous, so pipet it slowly to get accurate amounts when preparing the mix. Dilute the probes in TE buffer. The final concentration of the probe set is 2 nM. Mix the master mix well before adding to the wells. The following table shows example volumes needed to prepare a hybridization master mix containing two primary probe sets for 20 wells of a 384-well plate.

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Reagent

1 well

20 wells (including 20% extra)

Hybridization buffer

9.9 μL

237.6 μL

Probe TNF-B1 odd (0.08 μM)

0.275 μL

6.6 μL

Probe TNF-B1 even (0.08 μM)

0.275 μL

6.6 μL

Probe IL1B-B3 odd (0.08 μM)

0.275 μL

6.6 μL

Probe IL1B-B3 even (0.08 μM)

0.275 μL

6.6 μL

Total volume

11 μL

264 μL

6. Gently aspirate the pre-hybridization buffer from each well with a sterile 10 μL tip. The tip should always touch the same corner of the well to avoid cell loss. Do not aspirate the pre-hybridization buffer form the negative control wells (no primary probe set/amplifiers) (see Note 11). 7. Immediately add 11 μL/well of primary probe hybridization master mix to the plate. Do not add primary probe hybridization master mix to control wells. 8. Immediately seal the plate with the adhesive aluminum seal, and incubate the plate in a humidified, lightproof 37  C incubator for 12–18 h. A dedicated incubator should be assigned for this purpose during the hybridization of primary probe sets, to avoid temperature fluctuations caused by frequent opening and closing of the incubator door. 3.5.2 Plate Washes with Prewarmed Solutions to Remove Excess Probes

1. The next day, warm the probe wash buffer to 37  C in a water bath. 2. After the overnight incubation of the plate, aspirate the probe hybridization mix, followed by four 15-min washes in a 37  C water bath with prewarmed solutions to remove the excess probes (see Note 16). (a) Wash 1: 75% of probe wash buffer/25% 5X SSC-T (e.g., for 10 mL, add 7.5 mL probe wash buffer and 2.5 mL of 5X SSC-T), followed by (b) Wash 2: 50% of probe wash buffer/50% 5X SSCT, followed by (c) Wash 3: 25% of probe wash buffer/75% 5X SSC-T, followed by (d) Wash 4: 100% 5X SSC-T 3. After Wash 4, perform one more wash with 5X SSC-T for 5 min at RT.

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1. To equilibrate the plate before HCR amplification, aspirate the wash buffer, and then immediately add 16 μL/well of amplification buffer. 2. Incubate the plate for 30 min at RT. 3. During the plate equilibration (RT incubation), prepare the fluorescently labeled amplifiers: (a) Thaw the amplifiers on ice. (b) Add the necessary volume for each hairpin (See necessary hairpin quantities and volumes below. Add 10% more of the calculated hairpin volume to account for evaporation while snap-heating) to a new 1.5 mL Eppendorf microcentrifuge tube. Keep the amplifiers on ice. (c) Snap-heat the amplifiers at 95  C for 90 s. (d) Let the amplifiers cool down from 95  C to RT. 4. Add the necessary volume of amplifiers to a new 1.5 mL tube containing the calculated amount of amplification buffer. 11 μL/well of amplification master mix is needed, with each of the two hairpins at a final concentration of 60 nM (see Note 17). To prepare the amplification master mix necessary for 20 wells of a 384-well plate, add these volumes (including 20% extra):

Reagent

1 well

20 wells (including 20% extra)

Amplification buffer

10.56 μL

253.44 μL

Amplifiers-B1-Alexa488 (3 μM)

0.22 μL

5.28 μL

Amplifiers-B3-Alexa647 (3 μM)

0.22 μL

5.28 μL

Total volume

11 μL

264 μL

5. Mix the master mix well by flicking the tube, then briefly centrifuge. 6. Gently remove the equilibrating amplification buffer from the plate using vacuum aspiration. 7. Add 11 μL/well of amplification master mix to the plate. The ‘no primary probe set/no amplifiers’ negative controls should receive only amplification buffer. 8. Incubate the plate for 45 min at RT to perform the HCR amplification. Protect the plate from light from this step on. 9. Gently aspirate the amplification master mix from the plate. 10. Wash the plate with 80 μL/well of 5X SSC-T for 30 min at RT. 11. Discard the washing buffer by inverting and pat drying the plate.

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12. Repeat Steps 13–15 twice. 13. Stain cell nuclei by adding 12.5 μL/well of the DAPI working solution. 14. Incubate the plate with DAPI for 20 min at RT. 15. Aspirate the DAPI solution from the plate, and add 50 μL/well of PBS. 16. Seal the plate with the adhesive aluminum seal and store it at 4  C until imaging (up to 2 weeks). 3.6 HCI Acquisition and Analysis

1. If stored at 4  C, remove the HCR-stained imaging plate from the fridge, and let it equilibrate to RT (see Note 18).

3.6.1 HCI Acquisition Setup

2. Use a paper tissue and EtOH 70% to wipe clean the bottom of the plate to remove residual buffer, salts, dust, or condensation. 3. Load the 384-well imaging plate onto the automated microscope stage. 4. Select the appropriate objective. We routinely use a 60X water objective (NA (numerical aperture) 1.2]) with primary human immune cells because they tend to be small. 5. Select camera binning setting 2  2. 6. Program the microscope controlling software to acquire images in the necessary number (up to 4 in our case) of fluorescence channels. As an example, on our instruments, we use these combinations of excitation lasers sources and emission bandpass filters: (a) Ex: 405 nm, Em: 445/45 nm, DAPI (b) Ex: 488 nm, Em: 525/50 nm, Alexa488 (c) Ex: 561 nm, Em: 600/37 nm, Alexa546 or Alexa568 (d) Ex: 640 nm, Em: 676/29 nm, Alexa647 7. If using an automated spinning disk confocal microscope, select 3D z-stack acquisition for all fluorescence channels; otherwise, select to acquire an image in all channels at the focal plane. First use the DAPI channel to find the focal plane taking an interactive snapshot of a random FOV (see Note 19). If acquiring a z-stack, ensure that it contains the whole volume of most nuclei, and as little as possible empty space below and/or above the nuclei of most cells in the FOV. 8. Inspect the foreground (i.e., nuclei) to background ratio in the DAPI channel at the focal plane. Ideally, this ratio should be  5. 9. Check that the grayscale values in the image are in the dynamic range of the camera. Ensure that the fluorescence signal in the image is not saturated (i.e., the FOV does not include pixels

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Fig. 3 hcHCR to visualize TNF and IL1B induction upon LPS treatment. Primary human monocytes were treated either with vehicle or with 1 ng/mL of LPS and fixed at different time points as indicated in the figure. HCR was then performed with TNF-B1 and IL1B-B3 HCR probe sets and B1-Alexa488 and B3-Alexa647 HCR amplifiers. The negative control to measure fluorescence background (B1-Alexa488 and B3-Alexa647 HCR amplifiers only) is shown in the leftmost panel. The images were collected as 3D z-stacks at 60X magnification and maximally projected. Scale bar for full field of view: 20 microns. Scale bar for zoomed inset: 10 microns.

with grayscale values at or close to 4,095 for a 12-bit camera, or 65,535 for a 16-bit camera). 10. If necessary, change the camera exposure time and/or the excitation light power to satisfy the conditions mentioned in steps 8 and 9. 11. Once the z-offset, exposure time, and excitation power settings are set for the DAPI channel, repeat Steps 7–10 for the remaining HCR channels to ensure that these images are in focus and that the HCR fluorescence signal matches the dynamic range of the camera (Fig. 3; also see Notes 11 and 20). 12. Randomly acquire three to four additional snapshot images in a few random wells and random FOVs to make sure that the image acquisition parameters apply across the experiment. 13. At the end of the setup for the image acquisition settings for the 3D-stack in all channels, select to maximally project the 3D-stacks on the fly (see Note 21). 14. Select an appropriate number of randomly positioned FOVs for each well, aiming to image ~1,000 cells per well. 15. Select all the wells on the plate that need to be imaged. 16. Save the image acquisition settings (see Note 22). 17. Launch the batch image acquisition using the image acquisition settings optimized in Steps 4–15 (see Note 23). 3.6.2 HCI Analysis Setup

The dHCR spots need to be counted on a per cell basis in up to hundreds of thousands of images. For this reason, an automated HCI analysis pipeline is needed to analyze the images. These analysis pipelines can be set up using commercial software or using opensource software for image analysis (e.g., CellProfiler, Python, ImageJ, Julia). These are the steps in the HCI analysis pipeline:

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1. Find and segment nuclei in an automated manner in the DAPI channel image. Nuclei segmentation parameters in nuclear segmentation algorithms based on traditional image processing can be interactively optimized using a random FOV. More recent deep learning-based approaches for nuclear segmentation, such as Cellpose [24, 25] and StarDist [26], which do not require manual optimization of their parameters, can also be used. In either case, the user should verify that the nuclear algorithm correctly segments ~90–95% of the nuclei in a random test image. Once this is achieved, the user should test a few additional random FOVs in different wells to make sure that the segmentation algorithm and its parameters’ performance generalize to other images in the experiment. 2. Measure morphological parameters of the nucleus (area, circularity, length-to-width ratio). 3. Generate an approximate cell body region of interest (ROI) by dilating the nuclear region of interest (ROI) by a fixed percentage or number of pixels. The latter parameter will need to be visually optimized to ensure that the cell body contains most of the HCR signals in most cells while at the same time not extending in regions outside of cells. 4. Find HCR signals in the cell ROI in each one of the HCR channels. Like the nuclear segmentation step (Step 1), use a few random test FOVs in different wells to visually optimize the spot detection parameters. Aim to correctly detect ~90–95% of the HCR spots in each channel. 5. Optional: if manual optimization of the spot detection parameters does not lead to successful spot detection, use a supervised classification statistical learning model to classify spots after spot detection. This step requires first setting up permissive spot detection parameters in Step 3. Fluorescence intensity measurements (mean, sum, contrast, standard, deviation, mean intensity of the nucleus ROI containing the spot) should then be calculated for each spot over the spot ROI. This set of measurements can then be used as predictors to train a classifier to predict whether the detected spot is real, or it is a false positive. Use the HCR negative control wells to help with the annotation of false positives (see Notes 11 and 19). The classifier effectively works as a filter to eliminate spot detection artifacts. 6. Calculate the number of HCR spots per cell in each of the HCR channels as a proxy for transcript abundance (dHCR) [14]. Alternatively, if the gene measured by HCR is expressed at high levels, and the signal appears as diffuse due to a high number of HCR spots that cannot be optically resolved,

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measure the mean fluorescence intensity in the HCR channel as a proxy for transcript abundance (qHCR) [14]. 7. Use a combination of the nuclear ROI morphology parameters to filter out nuclear segmentation errors, such as merged or mis-segmented nuclei. 8. Launch the image analysis in batch on all the images in the experiment. 9. Output single-cell results as tabular text files (one file per well). In these tables, each row corresponds to a single-cell object as identified by the image analysis pipeline. For each row, the results tables should contain the following variables: (a) Plate ID (b) Well ID (c) Field of view ID (d) Object ID (i.e., the nucleus ID) (e) Number of HCR spots (one variable for each of the HCR channels) (f) Optional: mean fluorescence HCR intensity over the cell ROI (one variable for each of the HCR channels) (g) Any other relevant high-content feature/variable that is of interest for the downstream analyses.

4

Notes 1. We describe hcHCR in human primary monocytes purified from the peripheral blood as a specific example of a human primary cell. However, the protocol can be optimized for use with any other cell type. 2. We describe hcHCR after LPS stimulation as a specific example of a perturbation agent. However, the protocol can be optimized for use with any set of perturbations that is relevant to a specific experiment and cell type. 3. Dilute in 1X PBS if working with a different stock concentration of PFA. 4. Collection of human biospecimens, including blood, should be performed under a research protocol that conforms to the Declaration of Helsinki [27] and has been approved by an Institutional Review Board or equivalent human research ethics committee. 5. Small bubbles may be present in the density gradient medium after pipetting, but they will not affect performance.

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6. Different makes and models of centrifuges may provide different rates of deceleration when braking. If a layer of MNCs is not visible following centrifugation or the recovery of MNCs is low, reduce the rate of deceleration (i.e., braking) to medium or low. 7. Some red blood cells (RBCs) may be present on the surface of the SepMate insert after centrifugation. These RBCs will not affect performance. 8. To reduce platelet contamination in the enriched MNCs, pipette off some of the supernatant above the MNC layer before pouring. 9. If the density gradient medium above the SepMate insert appears red after centrifugation (i.e., some of the RBCs have not pelleted), the SepMate tube can be centrifuged at 1200  g for another 10 min at room temperature with the brake on. This step may be necessary when processing samples that were collected more than 24 h before processing. 10. If platelet removal is not required, then the repeated wash for the enriched MNC population with 1X PBS can also be performed by centrifuging at 300  g for 8 min at RT with the brake on, instead of 120  g for 10 min at RT with the brake off. 11. For each hcHCR experiment, several HCR negative controls, in addition to the experimental treatment controls, are necessary [14, 15]. For each combination of primary probe set and fluorescent HCR amplifiers, there should be two types of negative controls, with four wells allocated to each control. The first type of negative HCR control is no probe set plus no amplifiers. This negative control is used to estimate the autofluorescence of the cells. The second type of negative HCR control is no probe set plus amplifiers (Fig. 3). This negative control is used to estimate nonspecific binding of the fluorescent amplifiers, which can lead to HCR amplification in proximity to nontarget mRNA, protein, and lipid or at the bottom of the plate. These nonspecific events will result in small spotlike signals in the no probe set plus amplifiers control. To reduce this background, and during the assay development phase, it is useful to titrate the concentration of HCR amplifiers used in the reaction, to vary the amplification reaction time, and to increase the time and/or the number of washes after the amplification reaction. 12. For time-series experiments, it is important to fix with PFA all the LPS-treated wells on the plate at the same time, instead of treating them with LPS at the same time and fixing them at different time points. This reduces the possibility that PFA fumes from a fixed well can affect cell state and viability in

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adjacent, unfixed wells. For example, if the desired time points are 1, 2, and 3 h after treatment, add LPS to the corresponding wells 3, 2, and then 1 h before fixation, and fix all wells at the same time. 13. It is essential that cells are fixed in medium, without removing the medium or washing with PBS first, to avoid detachment of cells from the bottom of the plate. Most cell types, and in particular primary immune cells that grow in suspension, attach poorly to the plate and are easily dislodged by automated washes. 14. Alternatively, addition of PFA, PBS washes, and aspiration of buffers from the 384-well plate can be performed manually. Be extremely careful when pipetting liquids in wells or when aspirating them. Gently pipette to one corner of the well, and aspirate using vacuum from the same corner of the well to avoid dislodging fixed cells. 15. While proceeding with the HCR protocol immediately after permeabilization is recommended, this is a good stopping point in the protocol. The plate can be stored in 70% ethanol at 20  C for up to 1 month. 16. It is important to seal the plate for the four 15-min washes in a 37  C water bath and for the plate to sit on a raised platform in the water bath with just enough water to cover the sides of the plate without submerging it. 17. We have sometimes observed speckled fluorescence background signal in the “No primary probe set” negative control. The working concentration of fluorescent amplifiers used in the amplification reaction and the amplification reaction times should be optimized empirically by each user before proceeding with the actual experiment. 18. Letting the plate reach RT before starting the image acquisition setup is essential for two reasons. First, the functionality of the hardware autofocus systems is impaired by condensation that can form at the bottom of the plate when the plate is colder than the air surrounding it. This leads to autofocus failures, and it is an issue with air objectives. Second, the position of the focal plane during acquisition tends to be temperature-sensitive, and it can shift up or down by a few microns between lower temperatures and RT. If a cold plate is used to interactively set up the imaging parameters on the microscope, the z-offset defined in the software during the imaging setup is going to be mismatched with respect to the actual position of the focal plane when the plate reaches room temperature during image acquisition. As a result, the images during the automated acquisition will progressively be out of focus, blurry, and not usable for image analysis.

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19. This interactive mode is used to optimize the image acquisition settings. A visual image readout is used to optimize the z-offset, laser power, and camera exposure. As a result, the snapshot images acquired in interactive mode are not saved. 20. Digital HCR [14] fluorescence signals appear as spotlike features in the images close to or sometimes inside nuclei (Fig. 3). With the imaging settings described in this protocol, and with dHCR probe sets designed to target exons, these spots correspond to one or more mature mRNA molecules. The number of dHCR signals per cell varies substantially depending on the expression level of the gene probed by HCR and on the HCR probe set used for the experiment. The concentration of the primary probe set and of the fluorescently labeled HCR amplifiers should be optimized independently on a per cell type and per gene target basis to reduce nonspecific HCR background signal (i.e., the signal in images that are stained with the fluorescent HCR amplifiers but with no primary probe sets) (see Note 12) and to increase the number and foreground to background fluorescence ratio of the spotlike dHCR signals. Ideally, dHCR spots should have a foreground to background fluorescence ratio  3 to allow for spot detection in the downstream image analysis steps. 21. We maximally project the 3D z-stacks for all corrected images on the fly and only save the maximally projected image. We do this to increase the throughput of our measurements and to reduce the amount of data (up to tenfold) stored on disk. This becomes particularly useful for large datasets (up to 500 GB), leading to reduced data transfer time to network storage, and during image analysis. Of course, maximal projection leads to the loss of 3D information and to potential overlap of dHCR signals on different planes, thus leading to undercounting of dHCR signals during the image analysis. Since we consider this assay a high-content assay and we are only interested in changes in the number of dHCR signals per cell (as opposed to absolute values) between experimental conditions, since we acquire ~1000 cells, and since this is a systematic artifact in all different experimental conditions, we consider this a reasonable tradeoff for HCR of genes that are not highly expressed. Of course, researchers with different needs and/or requirements for their applications can decide to acquire just the focal plane or acquire, save, and analyze the full z-stack. 22. Image acquisition settings from previous similar experiments can be reutilized. Before launching a new batch image acquisition routing with old image acquisition settings, verify that they still apply to the current plate (Steps 4–15), and modify them if needed.

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23. Some microscope controlling software allows users to apply a “flat field” or “shading” illumination and background subtraction corrections to the images on the fly (i.e., while the instrument is running) using previously acquired reference images. This step is almost always necessary on spinning disk confocal microscopes, where illumination is uneven, being higher at the center of the FOV, and lower at the edges and corners of the image. The use of an illumination correction step has also been recommended as good practice for widefield fluorescence microscopes [28]. Similarly, on multi-camera instruments, vendors provide an option to run a geometric correction to compensate for translational and rotational differences between images acquired on different cameras. The geometric correction also compensates chromatic aberrations between different channels. We routinely use both sets of corrections on the fly for our measurements.

Acknowledgments GP and AC were supported by the Intramural Research Program (IRP) of the National Cancer Institute at the National Institutes of Health, 1-ZIC-BC011567-09. LMF and MG were supported by the IRP of the National Institute of Arthritis and Musculoskeletal and Skin Diseases at the National Institutes of Health. LMF, IF, and JS were supported by the Division of Intramural Research of the National Institute of Allergy and Infectious Diseases at the National Institutes of Health. The authors declare no conflict of interest. Figures 1 and 2 were made with BioRender and Adobe Illustrator. References 1. Sun J, Li N, Oh K-S et al (2016) Comprehensive RNAi-based screening of human and mouse TLR pathways identifies species-specific preferences in signaling protein use. Sci Signal 9:ra3 2. Hughes JP, Rees S, Kalindjian SB et al (2011) Principles of early drug discovery. Br J Pharmacol 162:1239–1249 3. Pegoraro G, Misteli T (2017) Highthroughput imaging for the discovery of cellular mechanisms of disease 33:604–615 4. Esner M, Meyenhofer F, Bickle M (2018) Livecell high content screening in drug development. Methods Mol Biol 1683:149–164 5. Mittelman D, Wilson JH (2013) The fractured genome of HeLa cells. Genome Biol 14:111

6. Gioia L, Siddique A, Head SR et al (2018) A genome-wide survey of mutations in the Jurkat cell line. BMC Genomics 19:334 7. Zhou B, Ho SS, Greer SU et al (2019) Haplotype-resolved and integrated genome analysis of the cancer cell line HepG2. Nucleic Acids Res 47:3846–3861 8. Lavrentieva A (2018) Essentials in Cell Culture. In: Kasper C, Charwat V, Lavrentieva A (eds) Cell culture technology. Springer, Cham, pp 23–48 9. Querido E, Dekakra-Bellili L, Chartrand P (2017) RNA fluorescence in situ hybridization for high-content screening. Methods 126: 149–155

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10. Femino AM, Fay FS, Fogarty K et al (1998) Visualization of single RNA transcripts in situ. 280:585–590 11. Pichon X, Lagha M, Mueller F et al (2018) A growing toolbox to image gene expression in single cells: sensitive approaches for demanding challenges. Mol Cell 71:468–480 12. Raj A, van den Bogaard P, Rifkin SA et al (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5:877–879 13. Gadkari M, Sun J, Carcamo A et al (2022) High-throughput imaging of mRNA at the single-cell level in human primary immune cells. RNA 28:1263–1278 14. Choi HMT, Schwarzkopf M, Fornace ME et al (2018) Third-generation in situ hybridization chain reaction: multiplexed, quantitative, sensitive, versatile, robust. Development 145: dev165753 15. Choi HMT, Beck VA, Pierce NA (2014) Nextgeneration in situ hybridization chain reaction: higher gain, lower cost, greater durability. ACS Nano 8:4284–4294 16. Python Software Foundation, https://www. python.org/ 17. Schneider CA, Rasband WS, Eliceiri KW (2012) NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671–675 18. Bezanson J, Edelman A, Karpinski S et al (2017) Julia: A fresh approach to numerical computing. SIAM Rev 59:65–98 19. Carpenter AE, Jones TR, Lamprecht MR et al (2006) CellProfiler: image analysis software for identifying and quantifying cell phenotypes. Genome Biol 7:R100 20. Stirling DR, Swain-Bowden MJ, Lucas AM et al (2021) CellProfiler 4: improvements in

speed, utility and usability. BMC Bioinform 22:433 21. R Core Team (2023) R: a language and environment for statistical computing, R Foundation for Statistical Computing, Vienna, Austria 22. Chen AR, McKinnon KP, Koren HS (1985) Lipopolysaccharide (LPS) stimulates fresh human monocytes to lyse actinomycin D-treated WEHI-164 target cells via increased secretion of a monokine similar to tumor necrosis factor. J Immunol 135:3978–3987 23. Kornbluth RS, Edgington TS (1986) Tumor necrosis factor production by human monocytes is a regulated event: induction of TNFalpha-mediated cellular cytotoxicity by endotoxin. J Immunol 137:2585–2591 24. Stringer C, Wang T, Michaelos M et al (2021) Cellpose: a generalist algorithm for cellular segmentation. 18:100–106 25. Pachitariu M, Stringer C (2022) Cellpose 2.0: how to train your own model. Nat Methods 19:1634–1641 26. Schmidt U, Weigert M, Broaddus C, et al (2018) Cell detection with star-convex polygons 27. World Medical Association (2013) World medical association declaration of Helsinki: ethical principles for medical research involving human subjects. JAMA 310:2191–2194 28. Bray M-A, Carpenter A, Imaging Platform, Broad Institute of MIT and Harvard (2012) Advanced assay development guidelines for image-based high content screening and analysis. In: Sittampalam GS, Coussens NP, Brimacombe K et al (eds) Assay Guidance Manual. Eli Lilly & Company and the National Center for Advancing Translational Sciences, Bethesda (MD)

Chapter 9 High-Throughput RNA-HCR-FISH Detection of Endogenous Pre-mRNA Splice Variants Asaf Shilo, Gianluca Pegoraro, and Tom Misteli Abstract RNA-fluorescence in situ hybridization (RNA-FISH) is an essential and widely used tool for visualizing RNA molecules in intact cells. Recent advances have increased RNA-FISH sensitivity, signal detection efficiency, and throughput. However, detection of endogenous mRNA splice variants has been challenging due to the limits of visualization of RNA-FISH fluorescence signals and due to the limited number of RNA-FISH probes per target. HiFENS (high-throughput FISH detection of endogenous pre-mRNA splicing isoforms) is a method that enables visualization and relative quantification of mRNA splice variants at single-cell resolution in an automated high-throughput manner. HiFENS incorporates HCR (hybridization chain reaction) signal amplification strategies to enhance the fluorescence signal generated by low abundance transcripts or a small number of FISH probes targeting short stretches of RNA, such as single exons. The technique offers a significant advance in high-throughput FISH-based RNA detection and provides a powerful tool that can be used as a readout in functional genomics screens to discover and dissect cellular pathways regulating gene expression and alternative pre-mRNA splicing events. Key words RNA-HCR-FISH, Alternative pre-mRNA splicing, High-throughput imaging, RNA imaging, FGFR2 splicing

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Introduction Single molecule RNA-FISH (smFISH) is considered the gold standard method for visualizing and quantifying mRNA molecules in their native cellular context [1]. smFISH techniques enable the examination of abundance and spatial distribution of RNA transcripts at single-cell and single-molecule resolution [2], providing information on gene expression patterns [3], RNA localization [4, 5], and cell-to-cell heterogeneity [6]. Nevertheless, traditional smFISH approaches still require tens of directly labeled DNA probes to generate a sufficiently strong fluorescent signal for detection. These approaches also generally necessitate high magnification and high numerical aperture (NA) objectives for detection, which limits the size of the field of view (FOV) and thus the

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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throughput of the method. For these reasons, there is a need for FISH-based techniques to increase the imaging throughput of RNA-FISH and to make the technique capable of working with short RNA targets. Such approaches are particularly useful for the detection and study of alternatively spliced exons. RNA splicing is the process in which noncoding intron sequences are removed from nascent RNA, and the coding exons are rejoined together to form the mature mRNA [7]. It is estimated that more than 90% of human genes undergo such alternative splicing (AS) [8]. During the process of AS, multiple or single exons, or parts of an exon, can be included or excluded from the final transcript, often in a combinatorial fashion in individual transcripts, thus producing a vast array of splice variants and, after mRNA translation, a variety of different protein isoforms from a relatively small number of genes. AS plays a crucial role in most biological processes, including development, tissue differentiation, response to environmental signals, and disease [8, 9]. Understanding how AS is regulated is important for our understanding of physiological cellular processes and for the development of molecular tools to modulate its activity for therapeutic applications [10, 11]. Given that differences between RNA splice variants are generally limited to short RNA sequences, typically on the order of 30–150 nt, it has been challenging to adapt traditional smFISH methods, especially in a high-throughput fashion, to detect RNA splice variants. Furthermore, most splicing assays have relied on use of minigene reporters, and analysis of splicing behavior of endogenous genes has been difficult. To overcome these challenges, and to adapt smFISH to study AS of endogenous genes at high throughput, we have developed HiFENS (high-throughput FISH detection of endogenous pre-mRNA splicing isoforms) [12]. HiFENS takes advantage of the previously developed hybridization chain reaction (HCR) signal amplification method [13]. In HCR, a DNA probe containing an “initiator” sequence is hybridized to a target RNA molecule, and subsequent incubation with labeled metastable DNA hairpin amplifiers, which only polymerize in the presence of the initiator, leads to polymerization and the accumulation of fluorescent label (Fig. 1). For high-sensitivity, high-throughput detection of alternatively spliced RNA isoforms by HiFENS, HCR-mediated FISH was adapted to a 384-well format and to automated liquid handling, thus enabling functional genomics screens that utilize endogenous RNA splice variant as a visual readout. As proof-of-principle, we have carried out a screen of the human kinome to identify modulators of alternative splicing of the endogenous FGFR2 genes (Fig. 2) [12]. Altogether, HiFENS is a powerful tool for both basic research and translational studies of AS.

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Fig. 1 Schematic representation of RNA-HCR. (a) Hybridization step. Black line represents target RNA molecule. Colored curved lines represent DNA probes. Each probe has three components: black and blue regions represent the specific recognition sequence which is complementary to the RNA target; gray region is a linker sequence; orange region is the initiator sequence. The full initiator is formed only when both split probes are bound in proximity to each other. (b) Amplification step. Blue and purple shapes represent the metastable hairpin amplifiers. The amplifiers are in a closed configuration in the absence of an initiator sequence. However, in the presence of initiator, the amplifiers are in an open configuration and can trigger the amplification chain reaction. Glowing spheres represent the fluorophore attached to the amplifiers

2

Materials

2.1 siRNA Plate Preparation

1. siRNA or sgRNA oligos library. The type of library is dependent on the desired assay. We used the Silencer™ Select Human Kinase siRNA Library (Ambion). 2. Allstars Hs Cell Death Control siRNA (QIAGEN), or appropriate sgRNA positive control for transfection. 3. Silencer Select Negative Control No. 2 siRNA (Ambion), or appropriate sgRNA negative control. 4. RNase-free water. 5. 384-well imaging plates (see Note 1).

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Fig. 2 Outline of the HiFENS pipeline. (a) Reverse siRNA transfection. Different colored lines represent siRNA oligonucleotides. Each well is spotted with a unique siRNA sequence (see Subheading 3.1 for more details). (b) RNA-in situ hybridization performed in a 384-well format in a fully automated manner. Black line represents target RNA molecule, and colored curved lines represent DNA probes (see Subheading 3.3.2 for more details). (c) RNA-HCR in a fully automated manner. Glowing spheres represent the full probe-amplifier complex. Red glowing spheres represent housekeeping gene detection used in our assay (TATA-Binding Protein, TBP). Green glowing spheres represent specific FGFR2 splice variant detection, and blue glowing spheres represent detection of the total FGFR2 transcript (see Subheading 3.3.3 for more details). (d) Highthroughput imaging. From each well, several fields of view were imaged in four channels to detect: DAPI, Alexa-Fluor-488, Alexa-Fluor-546, and Alexa-Fluor-647 (see Subheading 3.4 for more details). (e) Automated image analysis and spot detection. Image analysis is performed using Columbus (PerkinElmer) to identify the nuclei, cytoplasm (white circles), and spots (colored circles around a spot) in different channels. (f) Data analysis. A representative graph ranking all screen targets based on median Z-scores for FGFR2 splice variant spot count detection. (g) One field of view of MCF-7 cells. Purple outlined image was taken using the DAPI channel; red outlined image was taken using the far-red channel to detect total FGFR2 transcript; orange outlined image was taken using the red channel to detect TBP transcript; green outlined image was taken using the green channel to detect FGFR2 splice variant transcript. Lower panel shows magnified images of dotted boxes from the upper panel

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6. Automated liquid washer/dispenser compatible with 384-well plates. We used an Echo525 (Beckman-Coulter) acoustic liquid handler to spot the siRNA oligos in imaging plates. 7. Echo525-compatible source plates. 8. Aluminum adhesive seal. 2.2 Reverse Transfection in 384Well Imaging Plate Format

1. Adherent mammalian cell line of user’s choice (e.g., MCF7). 2. Transfection reagent. For MCF7 cells, DharmaFECT1 (Horizon Discovery) works best in terms of efficiency and lack of toxicity (see Note 2). 3. 20% serum Opti-MEM: 20% FBS serum in Opti-MEM reduced serum medium. 4. 1× PBS (phosphate-buffered saline). 5. Appropriate cell dissociation reagent. For MCF7 cells, we use trypsin/EDTA 0.25%. 6. Cell counter. 7. Automated cell/liquid dispenser compatible with 384-well plates. We used a Multidrop Combi Reagent Dispenser (Thermo-Fisher Scientific) (see Note 3).

2.3 RNA HCR FISH in 384-Well Imaging Plate Format

1. 1× PBS. 2. 8% paraformaldehyde (PFA): dilute 16% or 32% PFA stock solution in 1×PBS. 3. ddH2O (see Note 4). 4. 70% ethanol in water. 5. SSCT (saline sodium citrate – Tween-20): 5× SSC (in water), 0.1% Tween-20. 6. Probes (Molecular Instruments). Stock solution is 1 μM. For working solution concentration, see Subheading 3.3.2 (see Note 5). 7. Amplifiers (Molecular Instruments). Stock solution is 3 μM. For working solution concentration, see Subheading 3.3.3 (see Note 5). 8. Probe-hybridization buffer, probe wash buffer, and amplification buffer (Molecular Instruments) (see Note 5). 9. Humidified chamber at 37 °C for probe hybridization. 10. Automated liquid dispenser compatible with 384-well plates. We used a BlueWasher (BlueCatBio) for washing steps and a Mosquito (SPT Labtech) to add probes and amplifiers (see Note 3). 11. Heat block at 95 °C.

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12. DAPI (4′,6-diamidino-2-phenylindole) 10 mg/mL in PBS.

stock

solution:

13. DAPI staining solution: 2.5 μg/mL DAPI in PBS. 2.4 Automated Image Acquisition and Analysis

1. High-throughput spinning disk confocal microscope. (We used a Yokogawa CV7000.) 2. Image analysis software. We used PerkinElmer Columbus. However, image analysis software can be either commercial or open-source, such as Python, Julia, ImageJ, or CellProfiler. 3. Statistical analysis software. We use R (https://cran.r-project. org/) and RStudio Desktop (https://rstudio.com) for statistical analyses. However, statistical analysis can be performed with any other scientific programming language or platform, such as Python or Julia.

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Methods Before you start, here are some important guidelines for HiFENS: 1. Aliquot buffers (15–50 mL), probes (10–20 μL), and amplifiers (25–50 μL). Avoid repeat cycles of freeze-thaw. 2. Make fresh 8% PFA, 70% ethanol, 5× SSCT, and DAPI working solution on the day of use. 3. Some steps involve incubation at specific temperature. Adjust water bath/thermocycler accordingly before starting the protocol. 4. This protocol is based on the Molecular Instruments protocol for HCR, https://www.molecularinstruments.com/hcrrnafish-protocols. However, it is modified to make it compatible with automated liquid handling and imaging in a 384-well format for a maximum of five to ten plates per batch.

3.1 siRNA Plate Preparation

The siRNA library used in our screen is the Silencer Select Human Kinase library. The library contains 2127 unique siRNAs targeting 709 kinases in the human genome (3 siRNAs per target). The library (at 0.25 nmol synthesis scale) is received from the vendor in dehydrated form in 27 96-well plates. 1. Resuspend the oligo siRNA in 50 μL/well of ddH20 (5 μM final) in the plates using an automated liquid handler. Leave column 12 of the 96-well plate empty. The empty wells will be used for controls in the 384-well plate format used for imaging. See next steps for details.

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2. Freeze the plates o/n, and then thaw them to increase oligo siRNA solubility. 3. Compress the resuspended library from 96-well into 384-well format by transferring the entire volume of each well in the 96-well plate to a well in a 384-well plate using an automated liquid handler (every four 96-well plates yield a single 384-well plate). These 384-well plates are considered the “mother” plates. Columns 23 and 24 in the 384-well mother plate are left empty on purpose, so that siRNA controls can be added at a later stage. 4. For every mother plate, generate one “daughter” plate by transferring 10 μL of siRNA to an ECHO-compatible 384-well low dead volume (LDV) plate using an automated liquid handler. 5. Transfer control siRNAs at 5 μM concentration into a separate 384-well source plate compatible with the liquid handler to be used. 6. Centrifuge the source plates for 2 min at 250 g. 7. Spot 150 nL of control siRNAs at the bottom of each well of an empty imaging plate. For our screens, we used four controls: a negative scrambled control (Silencer Select Negative Control No.2 siRNA), a transfection control (Allstars Hs Cell Death Control siRNA), and two positive biological siRNAs. In our case, we detected FGFR2 pre-mRNA splicing and used siRNAs against its known regulators ESRP1 and ESRP2 as positive controls [14]. The biological positive controls should be determined and tested prior to performing the assay. We spotted eight wells for each control siRNA in every imaging plate (columns 23 and 24) for the screen. siRNA spotting was done using the Echo525, but this step can be done with any liquid handler capable of transferring small volumes (1–2 μL) (see Note 6). 8. Spot 150 nL of library siRNA per well of imaging plate (see Notes 6 and 7). 9. Air-dry the siRNA oligo in the laminar air flow hood (usually takes 45–90 min). 10. Seal the plate with aluminum adhesive seal, and store at -20 °C for up to 2 months. 3.2 Reverse Transfection in 384Well Imaging Plate Format

1. On the day of the transfection, allow the imaging plates containing the siRNA oligos to equilibrate at RT (~ 45 min). 2. Spin the imaging plates at 250 × g for 2 min. 3. Calculate the total volume of Opti-MEM needed. For example, for 6 imaging plates (384 wells per plate, 20 μL per well), the calculation for total Opti-MEM volume is as follows:

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6 plates × 384 wells per plate × 20 μL per well = 46.08 mL. Add extra volume, and account for the dead volume of the device of use. For example, add 1.25× extra volume and 20 mL dead volume. So, for six plates, use 80 mL (see Note 8). 4. Add the necessary volume of transfection reagent to the Opti-MEM. Use 37.5 nL DharmaFECT1 per well (150 μL of DhramaFECT1 into 80 mL of Opti-MEM) (see Note 2). 5. Add 20 μL of transfection reagent-Opti-MEM mix to each well of the imaging plates using an automated liquid dispenser (e.g., Thermo-Fisher Multidrop Combi). Incubate for 20–30 min at RT. Procced immediately to the next step. 6. While the transfection reagent and the Opti-Mem mix are incubating at RT with the siRNA in the plates, harvest cells from cell-culture flasks by trypsinization. 7. Count the cells, and calculate cell concentration. 8. Transfer the appropriate cell number into a new tube. For MCF7 cells, use 1,800 cells, seeded in 20 μL per well (90,000 cells/mL; for our screen, we need to have 7.2 × 10^6 cells in a total volume of 80 mL Opti-MEM) (see Note 9). 9. Centrifuge at room temperature to pellet cells at 250 × g for 5 min. 10. Resuspend cells in 10 mL of PBS. 11. Centrifuge to pellet cells at 250 × g for 5 min. 12. Resuspend cells to 90,000 cells/mL in 80 mL of 20% serum Opti-MEM. 13. Seed cells in the imaging plate by adding 20 μL of cell-OptiMEM-FBS mix per well (containing transfection reagent-OptiMEM mix from Step 5) using an automated liquid dispenser (e.g., Thermo-Fisher Multidrop Combi) (see Note 3). 14. Leave the imaging plate containing cells at room temperature for 30 min in the hood. This step has been shown to help solve issues with uneven cell growth patterns in the wells [15]. 15. Move the imaging plates to the incubator, and incubate the plate at 37 °C, 5% CO2, 80% humidity for 72 h (see Note 10). 3.3 RNA HCR FISH in 384-Well Imaging Plates 3.3.1 Fixation and Ethanol 70% Permeabilization

1. Add 40 μL per well of 8% PFA in PBS directly to the imaging plates containing cells using an automated liquid dispenser (e.g., Blue Washer). (See Notes 3 and 11.) 2. Incubate for 10 min at RT. 3. Wash the imaging plates 3× with 40–50 μL/well of PBS (see Note 12).

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4. Remove the PBS, and add 50 μL per well of 70% ethanol. 5. Seal the plates with aluminum sealer/parafilm, and store at 20 °C for 12–18 h (see Note 13). 3.3.2 Primary HCR Probe Set Hybridization

1. Warm up the hybridization buffer to 37 °C. 2. Carefully aspirate the 70% ethanol solution from the wells of the imaging plates by suction, or place the plate face down on absorbing paper. 3. Air-dry the plates for 10 min at RT to let ethanol residues evaporate. 4. Rehydrate cells in 40–50 μL per well of 5× SSCT for 5 min at RT. 5. Wash twice with 40–50 μL per well with 5× SSCT for 5 min. 6. Add 20 μL per well of 30% probe-hybridization buffer. This is a blocking step (see Note 14). 7. Incubate for 30 min in a 37 °C incubator. 6. During the blocking step, prepare the primary HCR probe sets in 30% probe-hybridization buffer. We find that for most targets, 2 nM final concentration of probes works well. For FGFR2 splice variant detection (four probes), we use 10 nM final concentration probes. However, the exact concentration must be optimized experimentally for each target, cell line, and assay format. 8. Aspirate blocking buffer. 9. Add 10 μL per well of probes in hybridization buffer mix (see Note 15). 10. Seal the plates with aluminum sealer. 11. Incubate the imaging plates containing the probehybridization buffer mix at 37 °C in an incubator for 12–18 h. Make sure a humid environment is maintained in the incubator to avoid drying of the wells.

3.3.3

HCR Reaction

Keep in mind the following before you start this section: • Pre-warm the solutions to be used. • Adjust the temperature of a heating block to 95 °C prior to the start of the experiment. • Thaw fluorescent HCR hairpin probes (see Note 16). • Allow the amplification buffer to warm to room temperature. 1. Wash imaging plates with 40–50 μL per well of a solution composed of ¾ vol/vol of probe wash buffer and with ¼ vol/vol of 5× SSCT for 15 min at 37 °C in incubator (Note:

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do not use water bath. Avoid water drops from entering the plates!). 2. Wash imaging plates with 40–50 μL per well of a solution composed of 1/2 vol/vol of probe wash buffer and with 1/2 vol/vol of 5× SSCT for 15 min at 37 °C in incubator. 3. Wash imaging plates with 40–50 μL per well of a solution composed of ¼ vol/vol of probe wash buffer and with ¾ vol/vol of 5× SSCT for 15 min at 37 °C in incubator. 4. Wash with 40–50 μL per well of 100% 5× SSCT for 15 min at 37 °C in incubator. 5. Wash with 40–50 μL per well 5× SSCT for 5 min at RT. 6. Aspirate the 5× SSCT from the plates, and add 20 μL/well of amplification buffer. 7. Incubate the plates at RT for 45 min. This is a blocking step. Then proceed immediately to next step. 8. During the blocking step, prepare primary HCR probe sets in probe-hybridization buffer. Transfer the needed volume of fluorescent HCR hairpin probes into new 1.5 mL Eppendorf tubes. Each amplifier should be in a separate tube. We find that for most targets, 30 nM final concentration amplifiers work well. For isoform detection, we used amplifiers at a final concentration of 60 nM. However, the exact amplifier concentration must be optimized experimentally for each target, cell line, and assay format. 9. Denature the fluorescent HCR amplifiers by placing the tubes in a hot block at 95 °C for 120 s (see Note 17). 10. Allow the fluorescent HCR hairpin probes to cool down for 30 min at RT protected from light. 11. Add the fluorescent HCR hairpin probes to the amplification buffer. At this point, all the HCR hairpin probes should be in the same tube. 12. Aspirate the blocking buffer from the plates. 13. Dispense 10 μL/well of fluorescent HCR hairpin in amplification buffer mix into the imaging plates (see Note 15). 14. Incubate plates for 45 min at RT protected from light (see Note 18). 15. Wash 5× with 40–50 μL per well of 5× SSCT for 5 min at RT. 3.3.4

DAPI Staining

1. Aspirate the 5× SSCT from the imaging plates, and add 40 μL per well of DAPI staining solution. 2. Incubate plates for 30–60 min at RT. 3. Aspirate the DAPI solution, and add 55 μL per well of PBS.

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4. Continue directly to imaging of the plates, or store at 4 °C for up to 1 week protected from light. 3.4 Automated Image Acquisition

1. If previously stored at 4 °C, allow imaging plates to warm to room temperature for at least 45 min. 2. Clean the bottom of the imaging plates with 70% ethanol. Make sure that dust, condensation, or buffer salt residues are wiped off. 3. Load imaging plate onto the microscope stage. 4. The parameters for high-throughput image acquisition are assay-dependent and need to be optimized. For our assays, we programmed the Yokogawa CV7000 high-throughput microscope to these parameters: (a) Objective: 40× air objective (NA 0.9). Typically, one field of view (FOV) captures about 100–200 MCF7 cells (Fig. 2g). (b) sCMOS camera (2560 × 2160 pixels) binning: 2 × 2. pixel size: 0.325 microns. (c) Excitation lasers and bandpass emission filters. All channels used a fixed 405/488/561/640 nm dichroic excitation mirror and a fixed 568 nm dichroic emission mirror: (i) Ex:405 nm, Em:445/45, for DAPI. (ii) Ex:488 nm, Em:525/50, for Alexa-Fluor-488. (iii) Ex:561 nm, Em:600/37, for Alexa-Fluor-546. (iv) Ex:640 nm, Em:676/29, for Alexa-Fluor-647. (d) Select 3D z-stack acquisition. (e) Select on the fly maximum z-stack projection. 5. To find the optimal setting for z-stacks and exposure time, select a random well and field of view, and test different conditions for the DAPI channel. The z-stack should include in-focus nuclei. Once parameters are set, run an acquisition test by changing projection to maximum projection. If the test image is blurry, out of focus, or overexposed, adjust parameters, and repeat steps until the image appears focused and not overexposed. 6. Repeat this process for all channels. The z-stack should include focused, not overexposed, spots. If the test image is blurry, out of focus, or overexposed, adjust parameters, and repeat the process described above (Step 5), until the image appears focused and not overexposed (Fig. 2g). 7. Test a few wells using the imaging parameters from pervious steps to verify that the image acquisition parameters apply to the rest of the plate.

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8. Select several fields of view per well. We usually aim to image around 1000–1500 cells per well. For MCF7, we usually image eight to nine fields of view per well. 9. Launch the image acquisition program for all plates. 3.5 Image Analysis and Statistical Analysis

4

Analysis of images and downstream statistical analysis are highly dependent on user needs and assay design and are beyond the scope of this protocol. Briefly, our screen image analysis was done using Columbus 2.8.1 (PerkinElmer). First, automated nucleus segmentation and cytoplasm detection were based on the maximal projection of the DAPI signal (e.g., 405 nm). For detection of HCRRNA-FISH spots within these cells, maximal projections of the green (e.g., 488 nm), red (e.g., 561 nm), or far-red (e.g., 640 nm) channels were used. Next, we applied a linear classifier algorithm to separate between positive and negative (background) spots for each channel. Single-cell results generated in Columbus were exported into R software (4.1, R Core Team) and RStudio Desktop (RStudio) and analyzed using custom made scripts. For the full scripts, see GitHub - CBIIT/mistelilab-hifens [12]. We used a median per gene threshold z-score of 1.5 (i.e., at least two out of the three siRNA oligos with a z-score > 1.5 or z-score < 1.5) to identify hits.

Notes 1. The HiFENS protocol was designed and optimized for highthroughput assays in 384-well imaging plates. If using different plates, all volumes need to be adjusted and optimized accordingly. 2. The transfection reagent and its concentration should be optimized beforehand as follows: (a) Spot imaging plate with negative and positive transfection controls (see Subheading 3.1, Step 7). (b) Allow siRNA imaging plate to air-dry. Usually, it takes about 45–90 min. It can be used immediately once dry or frozen at -20 °C until use. (c) Use Multidrop dispenser or another liquid handler to add 20 μL Opti-MEM per well. (d) Use the ECHO525 to dispense different concentrations of transfection reagent into wells already containing 20 μL Opti-MEM (from Step c). For example, add 0, 25, 50, 75, 100,125, and 150 nL of transfection reagent per well. (e) Cover plate with plastic lid, and leave at RT for 30 min in the laminar flow hood.

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(f) During this incubation step, prepare and count your cells. (g) Use Multidrop dispenser to add 20 μL of cells resuspended in 20 μL of Opti-MEM supplemented with 20% FBS. (h) Cover plate with plastic lid, and leave at RT for 30 min in the laminar flow hood. (i) Incubate the plate at 37 °C, 5% CO2, 80% humidity for 72 h. (j) Condition with high efficiency (based on the positive control, meaning high cell death) and minimal toxicity (low cell death in the negative control wells) should be chosen for future screens. 3. Multichannel pipettes (10–300 μL) can be used instead. However, it increases the likelihood for errors and is very timeconsuming. 4. Any procedure that involves water must use high grade water such as UltraPure, DEPC, etc. 5. We purchased probes, amplifiers, and all buffers from Molecular Instruments (https://www.molecularinstruments.com/). 6. In our application, siRNA final concentration was 18.75 nM. However, the final concentration can vary based on need. 7. In our library, we have three different siRNAs per target. We spotted one siRNA per well. The number of replicates should be considered before starting the protocol. 8. Always include extra volume. Consider if the liquid handler has dead volume, and add this to the volume calculations, and then add about 12.5–25% extra volume to ensure sufficient volume. 9. The number of cells per well needs to be calibrated beforehand, as follows: (a) Seed different cell numbers per well in replicates. For example, seed 1000, 2000, 3000, 4000, 5000, and 6000 cells per well. (b) Incubate the plate at 37 °C, 5% CO2, 80% humidity for 72 h. The duration of treatment is based on need; we used 72 h for siRNA treatment. (c) Wells with 70–80% confluency after 72 h should be selected for use. 10. The duration of treatment is assay-dependent. We typically treat cells for 72 h with siRNA. 11. Make fresh 8% PFA each time. 12. Cells can be washed with 2× SSC. We did not observe any difference when washing with PBS or 2× SSC.

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13. Plates can be stored at -20 °C. We have not determined maximum storage time. 14. Make sure the buffer is clear. Aggregate formation in the buffer may cause unsuccessful hybridization. To avoid aggregations, incubate at 37–40 °C, and vortex vigorously. 15. We used the Mosquito dispenser (SPT Labtech) to add probes and amplifiers. Hybridization buffer is very viscous, and some liquid handlers are unable to transfer it. It is advised to check if the liquid handler can transfer hybridization buffer before starting the protocol. Amplification buffer is not viscous and easier to transfer. If using electric multichannel pipettes, consider transferring more than 10 μL per well (12–13 μL per well). 16. Hairpins/amplifiers are light sensitive. All steps involving amplifiers should be done protected from light. 17. For large volume of amplifiers (>200 μL), amplifiers can be split into different tubes to ensure complete denaturation. 18. Incubation time can be longer, based on assay need. References 1. Le P, Ahmed N, Yeo GW (2022) Illuminating RNA biology through imaging. Nat Cell Biol 24(6):815–824 2. Tingey M et al (2022) Technologies enabling single-molecule super-resolution imaging of mRNA. Cells-Basel 11(19) 3. Xia C et al (2019) Spatial transcriptome profiling by MERFISH reveals subcellular RNA compartmentalization and cell cycledependent gene expression. Proc Natl Acad Sci U S A 116(39):19490–19499 4. Weil TT, Parton RM, Davis I (2010) Making the message clear: visualizing mRNA localization. Trends Cell Biol 20(7):380–390 5. Engel KL et al (2020) Mechanisms and consequences of subcellular RNA localization across diverse cell types. Traffic 21(6):404–418 6. Levitin HM, Yuan J, Sims PA (2018) Singlecell transcriptomic analysis of tumor heterogeneity. Trends Cancer 4(4):264–268 7. Wilkinson ME, Charenton C, Nagai K (2020) RNA splicing by the spliceosome. Annu Rev Biochem 89:359–388 8. Pan Q et al (2008) Deep surveying of alternative splicing complexity in the human

transcriptome by high-throughput sequencing. Nat Genet 40(12):1413–1415 9. Scotti MM, Swanson MS (2016) RNA mis-splicing in disease. Nat Rev Genet 17(1): 19–32 10. Stanley RF, Abdel-Wahab O (2022) Dysregulation and therapeutic targeting of RNA splicing in cancer. Nat Cancer 3(5):536–546 11. Ren P et al (2021) Alternative splicing: a new cause and potential therapeutic target in autoimmune disease. Front Immunol 12:713540 12. Shilo A, Pegoraro G, Misteli T (2022) HiFENS: high-throughput FISH detection of endogenous pre-mRNA splicing isoforms. Nucleic Acids Res 50(22):e130 13. Choi HMT et al (2018) Third-generation in situ hybridization chain reaction: multiplexed, quantitative, sensitive, versatile, robust. Development 145(12) 14. Warzecha CC et al (2009) ESRP1 and ESRP2 are epithelial cell-type-specific regulators of FGFR2 splicing. Mol Cell 33(5):591–601 15. Lundholt BK, Scudder KM, Pagliaro L (2003) A simple technique for reducing edge effect in cell-based assays. J Biomol Screen 8(5): 566–570

Chapter 10 Simultaneous In Situ Detection of m6A-Modified and Unmodified RNAs Using DART-FISH Charles J. Sheehan and Kate D. Meyer Abstract N6-methyladenosine (m6A) is an abundant mRNA modification which plays important roles in regulating RNA function and gene expression. Traditional methods for visualizing mRNAs within cells cannot distinguish m6A-modified and unmodified versions of the target transcript, thus limiting our understanding of how and where methylated transcripts are localized within cells. Here, we describe DART-FISH, a visualization technique which enables simultaneous detection of both m6A-modified and unmodified target transcripts. DART-FISH combines m6A-dependent C-to-U editing with mutation-selective fluorescence in situ hybridization to specifically detect methylated and unmethylated transcript copies, enabling the investigation of m6A stoichiometry and methylated mRNA localization in single cells. Key words N6-methyladenosine (m6A), Fluorescent in situ hybridization (FISH), Padlock probe, Epitranscriptomics, RNA modifications, Deamination adjacent to RNA modification targets (DART)

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Introduction N6-methyladenosine (m6A) is an abundant internal mRNA modification which influences the processing and expression of modified transcripts [1–3]. Over the last decade, sequencing-based approaches to map m6A transcriptome-wide have provided invaluable insights into the regulation of modified mRNAs. However, due to the requirement of these methods to purify mRNA, our understanding of how m6A-modified mRNAs are spatially regulated within cells remains poorly understood. Traditional methods for visualizing RNAs within cells rely on the hybridization of complementary oligonucleotide probes which are either directly labeled or detected by secondary methods [4]. However, these strategies are insensitive to RNA modifications such as m6A which do not alter Watson-Crick base-pairing [5]. To overcome these limitations, we developed DART-FISH, a technique to simultaneously visualize m6A-modified and unmodified

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Schematic representation of the DART-FISH method. (a) Expression of APO1-YTH in cells directs C-to-U deamination adjacent to m6A sites transcriptome-wide. (b) Cells are then fixed and subjected to reverse transcription using a targeted locked nucleic acid (LNA) primer. (c) The target RNA molecule is degraded by RNAse H, and padlock probes complementary to the cDNA sequence with either a C or U adjacent to the m6A site are hybridized. The 50 and 30 ends of the padlock probes are ligated together only if the probe and cDNA are perfect complements. (d) Ligated padlock probes are then subjected to rolling-circle amplification, generating a rolling-circle product (RCP) containing iterative copies of the unique detection sequence from the padlock probe. Individual padlock probe hybridization and ligation events are visualized via hybridization of fluorescent oligonucleotides complementary to the detection sequence

target mRNAs of interest in cells [6]. To achieve this, DART-FISH couples m6A-adjacent RNA editing [7] with single-nucleotide variant (SNV)-sensitive fluorescence in situ hybridization [8, 9] to visualize m6A-modfied and unmodified versions of transcripts of interest (Fig. 1). To detect m6A-modified mRNAs, DART-FISH utilizes the expression of a fusion protein consisting of the C-to-U deaminase, APOBEC1, tethered to the m6A-binding YTH domain [7]. Expression of APOBEC1-YTH (hereafter APO1-YTH) in cells directs Cto-U deamination of cytidine residues which immediately follow nearly all m6A-modified adenosines [7, 10]. Delivery of the APO1YTH transgene can be accomplished using a variety of methodologies, including transient transfection, stable cell generation, or viral transduction. Below, we outline the DART-FISH protocol as applied to stable doxycycline-inducible immortalized cell lines. Visualization of APO1-YTH-induced C-to-U mutations adjacent to m6A sites is achieved using padlock probe (PLP) hybridization and rolling-circle amplification (RCA) [8, 9]. First, cells are fixed, and reverse transcription (RT) is performed using a custom RT primer for an RNA of interest. Following RNase H treatment to remove the bound RNA, PLPs are then hybridized to the cDNA. PLPs consist of DNA oligonucleotides which circularize upon hybridization to the target molecule. Each PLP contains homology arms at the 50 and 30 ends of the oligonucleotide which recognize the target RNA and a unique detection sequence within the linker region between the homology arms (Fig. 2). To enable SNV discrimination, the 30 homology arm is designed such that its 30 -most residue hybridizes to a nucleotide variant of interest in the target cDNA. Phosphorylation of the 50 end of the PLP enables ligation

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Fig. 2 Schematic representation of padlock probes targeting the ACTB A1222 m6A site. Two padlock probes are designed for an individual m6A site, one probing for C (a) and the other for U (b) adjacent to the site of interest. The locked nucleic acid (LNA) reverse transcription (RT) primer (orange arrow) is bound to the original mRNA molecule, tethering the cDNA molecule in place. Each padlock probe is designed complementary to the cDNA molecule with homology arms at the 50 and 30 ends of the padlock probe (red nucleotides) and the single nucleotide of interest, either C or U, at the 30 end of the probe. Each padlock probe contains a unique detection sequence (represented by cyan or gold portion of the padlock probe) between two poly(A) linker regions joining the homology arms and detection sequence. Ligation of the 50 and 30 ends of the padlock probe closes the circularized padlock probe allowing for subsequent rolling-circle amplification

and circularization only if the homology arms perfectly pair with the target sequence. Successfully ligated and circularized PLPs then serve as templates for RCA, which generates rolling-circle products (RCPs) that contain repetitive iterations of the unique detection sequence in the PLP (Fig. 2). Visualization of successful PLP ligation events is then achieved using fluorescent oligonucleotides complementary to the unique detection sequence. Thus, this approach translates distinct SNVs into unique fluorescent signals. By coupling this with APO1-YTH expression, the unmodified and m6A-modified variants of an RNA of interest can be simultaneously visualized using distinct PLPs targeting the C and U variants, respectively, of the RNA. Of note, due to the series of enzymatic steps, PLP-RCA on average detects less than 30% of target transcript copies compared to alternative FISH methods. This includes sites of nascent RNA transcription [8, 9].

2 Materials 2.1 Inducible APO1YTH Stable Cell Lines

1. HEK293T cells. 2. pTLCV2 APO1-YTH-T2A-eGFP #178949). 3. pTLCV2 APO1-YTHmut-T2A-eGFP #178950).

plasmid plasmid

(Addgene (Addgene

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4. psPax2 plasmid (Addgene #12260). 5. pMD2.G plasmid (Addgene #12259). 6. 1 mg/mL jet-PEI transfection reagent. 7. HeLa cells. 8. 10 mg/mL puromycin. A limitation of DART-FISH is the requirement to express APO1-YTH in the cells or tissue of interest. However, this can be accomplished using a variety of methods, such as stable cell generation, transient transfection, or viral transduction. Below we describe methodology to generate concentrated APO1YTH lentivirus for the infection and selection of doxycyclineinducible cell lines. This methodology works well in our hands across various mammalian cell lines, including those of both human and mouse origin. Additionally, in parallel to APO1-YTH expression, we also utilize the expression of a control construct with a mutant YTH domain which has reduced affinity for m6A [7]. Although this mutant enzyme still retains some low-level binding to m6A, its expression leads to a substantial reduction in U variant abundance compared to the APO1-YTH protein. 2.2 Reverse Transcription Primer

P-ACTB_ A1222 50 μM 50 G + CC + AT+G C + CA + AT+C T + CA TCT TGT TTT CTG (+ indicates locked nucleic acid). For each target m6A site, there is a single reverse transcription primer used. Reverse transcription primer design is a critical factor of DART-FISH success since the efficiency of this initial step determines the maximum number of transcripts that can ultimately be visualized. To maximize the number of transcripts detected, primers should be placed within 100 nt downstream of the m6A site of interest and contain 7 “locked” nucleic acid bases, which have higher target affinity compared to conventional DNA bases [11] (see Note 1). We recommend designing RT primers using Primer3Plus [12], although other primer design programs should also work. In general, oligonucleotides with longer length (between 25 and 30 nt) and high maximal free energy of binding to the perfect complement (ideally < 45 kcal/moL) provide the best detection. Programs that predict oligonucleotide secondary structure such as IDT OligoAnalyzer [13] or RNAfold (http://rna.tbi.univie.ac.at/cgi-bin/ RNAWebSuite/RNAfold.cgi) can then be used to inspect primers for deleterious structures such as strong hairpins and dimerization with melting temperatures >50  C. Special attention should be taken to ensure LNA bases are not placed within predicted secondary structures. Once a targeted RT primer is designed, it can then be synthesized or ordered from a commercially available source. We recommend purchasing from Integrated DNA Technologies (IDT) as custom oligonucleotides with LNA base substitutions.

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1. PLP-ACTB-C1223 5 μM /5Phos/TTAGTTGCGTTACACCCTTTC AAAAAAAAAAA AAAAAGTAGCCGTGACTATCGACTAAAAAAAAAAAAAA AACTTCTAGGCGGACTATGAC 2. PLP-ACTB-U1223 5 μM /5Phos/TTAGTTGCGTTACACCCTTTC AAAAAAAAAAAAAACCTCAATGCACATGTTTGGCTCC AAAAAAAAAAAAAACTTCTAGGCGGACTATGAT. Underlined text indicates the homology arms. Highlighted text indicates the unique detection sequence. The bold nucleotide indicates the C or U being detected. PLPs can be designed by hand or using software such as ProbeMaker [14]. Each padlock probe consists of two homology arms complementary to the target RNA sequence, a poly(A) linker region (see Note 2), and a unique detection sequence used to visualize the rolling-circle products from specific padlock probes. Complementary PLP arms should generally each be ~20 nucleotides long and designed to hybridize to the cDNA complement of the target mRNA. The single nucleotide of interest, in this case either C or U, should be placed at the 30 end of the PLP, and the 50 -end of the PLP should be phosphorylated. PLPs are ordered as Ultramers from IDT with a 5’phosphate modification (see Note 3).

2.4

Detection Probes

1. DT3-Cy3 5 μM /5Cy3/AGTAGCCGTGACTATCGACT. 2. DT2-Cy5 5 μM /5Cy5/CCTCAATGCACATGTTTGGC TCC. Detection probes should be labeled with two different fluorophores which enable nonoverlapping detection of each respective RCP. We frequently utilize Cy3 and Cy5 since these fluorophores produce bright visible signal and are easily spectrally separated using traditional fluorescence microscopes. The detection sequence should be identical to that of the respective PLP, since RCA generates a DNA cloud containing the complementary sequence. Detector probes can be ordered from IDT as custom oligos with 50 fluorophore modifications and purified by HPLC.

2.5

Reagents

1. Trypsin solution. 2. 0.1 mg/mL poly-D-lysine (PDL). 3. 1 mg/mL doxycycline. 4. Anti-HA antibody (Cell Signaling Technology, 3724) and suitable secondary antibody for Western blot. 5. (Optional) Primary antibody for immunofluorescence. 6. (Optional) Secondary antibody for immunofluorescence (see Note 4).

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7. 4% paraformaldehyde *toxic; handle with care. 8. 0.1 M HCl *highly corrosive; handle with care. 9. 5 M-MuLV reverse transcriptase buffer 10. 200 U/μL RevertAid reverse transcriptase (see Note 5). 11. 25 mM dNTPs. 12. 50 mg/mL BSA. 13. 40 U/μL RiboLock RNAse Inhibitor. 14. 50 μM RT primer. 15. 10 Ampligase Reaction Buffer (BioSearch). 16. Formamide *toxic; handle with care. 17. 2.5 M KCl. 18. 20 U/μL RNAse H. 19. 5 U/μL Ampligase (BioSearch). 20. Glycerol. 21. 10 Phi29 DNA polymerase buffer. 22. 10 U/μL Phi29 DNA polymerase. 23. 1 mg/mL DAPI. 24. Vectashield vibrance or alternative anti-fade mounting media. 2.6 Buffers and Reaction Solutions

1. Cell culture medium: 1 Dulbecco’s Modified Eagle’s Medium, 10% fetal bovine serum, 10 units/mL penicillin, and 10 μg/mL streptomycin. 2. Sterile 20% sucrose in 1 PBS. 3. 2 saline sodium citrate (SSC) buffer: 0.3 M sodium chloride and 0.03 M sodium citrate. 4. 70% ethanol. 5. 1 phosphate-buffered saline (PBS): 137 mM NaCl, 10 mM sodium phosphate, 2.7 mM KCl in Ultrapure water pH 7.4. 6. 0.05% Tween-20 in 1 PBS (v/v) (PBST). 7. RT mix (50 μL per coverslip): 10 μL 5 M-MuLV RT buffer, 1 μL 25 mM dNTPs, 0.2 μL 50 μg/μL BSA, 1.25 μL RiboLock RNAse Inhibitor, 1 μL RT primer, 1.25 μL RevertAid Reverse Transcriptase, 35.3 μL Ultrapure water. 8. Hybridization-ligation mix (50 μL per coverslip): 5 μL 10 Ampligase buffer, 10 μL formamide, 1 μL 2.5 M KCl, 0.33 μL RNAse H, 1.25 μL RiboLock RNAse Inhibitor, 1 μL 5 μM C – PLP, 1 μL 5 μM U – PLP, 5 μL Ampligase, 25.42 μL Ultrapure water.

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9. RCA mix (50 μL per coverslip): 5 μL glycerol, 5 μL 10 Phi29 DNA polymerase buffer, 1 μL 25 mM dNTPs, 0.2 μL BSA, 1.25 μL RiboLock RNAse Inhibitor, 5 μL Phi29 DNA polymerase, 32.55 μL Ultrapure water. 10. Detection probe hybridization mix: 37.95 μL 2 SSC, 1 μL of each 5 μM detection probe, 0.05 1 mg/mL DAPI, 10 μL formamide. 2.7

Equipment

1. 15 cm tissue culture plates. 2. Centrifuge capable of 5000  g. 3. Ultracentrifuge with swinging bucket rotor capable of 32,672  g (Beckman-Coulter, SW28). 4. 0.45 μM filter. 5. Ultracentrifuge tubes. 6. 10 cm tissue culture plates. 7. 12-well tissue culture plates. 8. #1.5 coverslips. 9. Staining container. 10. Whatman paper. 11. Parafilm. 12. 37  C incubator (see Note 6). 13. 45  C incubator. 14. Permanent marker. 15. Microscope (see Note 7). 16. ImageJ/FIJI (https://imagej.net/software/fiji/). 17. CellProfiler (https://cellprofiler.org). 18. FISHQuant (https://fish-quant.github.io).

3

Methods

3.1 Target m6A Site Selection

To identify m6A sites of interest, we recommend performing an initial DART-seq [7] experiment or, for specific RNAs of interest, using targeted RT-PCR and Sanger sequencing to identify sites of C-to-U editing. DART-FISH accurately visualizes C-to-U mutation levels as low as 5%. However, initial optimization of DARTFISH in cell lines of interest using more frequently methylated sites (>10% C-to-U editing) is recommended. Additionally, compared to traditional smFISH methods, PLP-RCA-based detection is much less sensitive due to the series of enzymatic steps required for successful visualization and the use of single probes instead of tiling oligos. Because of this limitation, RNA abundance is an

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important consideration when performing DART-FISH. Individual molecules of low-abundance transcripts may be difficult to detect in sufficient numbers. If RNA-seq datasets from cell lines of interest are available, we recommend looking at the coverage at a given m6A site. This helps ensure sufficient expression of the transcript region containing the target m6A site. In general, we have had good success performing DART-FISH on RNAs with values of >700 normalized transcripts per million (nTPM). 3.2 Generation of Stable Inducible APO1YTH Cell Lines 3.2.1 APO1-YTH Lentivirus Production

1. Culture a 15 cm dish of HEK293T cells in culture medium at 37  C with 5% CO2 until 80% confluent. 2. Transfect cells with 26.75 μg pTLCV2-APO1-YTH-T2AeGFP (see Note 8), 20 μg psPAX2, and 6.25 μg of pMD2.G using 159 μL of jet-PEI according to manufacturer’s instructions. Incubate for 72 h. 3. Collect medium, and centrifuge at 5000  g for 10 min. 4. Collect the supernatant, and filter through 0.45 μm filter into ultracentrifuge tube. 5. Gently underlay 4 mL of sterile 20% sucrose to the supernatant. 6. Centrifuge at 32,672  g for 2 h at 4  C in SW28 swinging bucket rotor. 7. Remove supernatant, and dry tube for 5 min inverted. Remove additional liquid around the rim of the tube. 8. Resuspend pellet in 100 μL of 1 PBS. Rock at 4  C protected from light overnight. 9. Store virus in 20 μL aliquots long term at 80  C.

3.2.2 Infection and Selection of Stable Cell Lines

1. Culture desired cell line (e.g., HeLa) to ~60–80% confluence. 2. Infect with 1:500 APO1-YTH-T2A-eGFP lentivirus (see Note 9). 3. Following 48 h of infection, change culture media to contain 2 μg/mL puromycin. 4. Maintain cells under puromycin selection for at least 72 h or until all cells in a noninfected control plate are dead. 5. Passage cells as necessary into sequentially larger culture dishes up to 10 cm plates. 6. (Optional) For clonal selection, passage and serially dilute cells into a 96-well plate to obtain single-cell clones in a single well (see Note 10). Maintain cells under selection, and allow these clones to grow to fill the culture dish. Passage as necessary back up to a 10 cm plate. 7. To validate doxycycline-inducible APO1-YTH expression, replace media with media containing doxycycline at a final concentration of 1 μg/mL.

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8. Confirm protein expression using immunoblotting targeting the HA tag on the C-terminus of the APO1-YTH protein. Confirm m6A-adjacent C-to-U deamination via RT-PCR and Sanger sequencing. 3.3 Cell Culture and Sample Preparation

1. Wash coverslips three times in 70% ethanol for 15 min each on shaker at room temperature.

3.3.1 Coverslip Preparation

2. Wash cleaned coverslips in PBS three times at room temperature for 3 min each on shaker. 3. Dilute 0.1 mg/mL poly-D-lysine (PDL) to 50 μg/mL in PBS, and incubate coverslips for >30 min in 50 μg/mL PDL at 37  C. 4. Remove PDL, and wash coverslips three times in Ultrapure water for 3 min each on shaker at room temperature. 5. Place each coated coverslip into 12-well culture dish well. 6. Use immediately, or store at 4  C for up to 3 months.

3.3.2

Cell Culture

1. Culture stable cells in a 10 cm plate until 80–90% confluent. 2. Remove media, and wash once with 1 PBS gently. 3. Add 1 mL of trypsin to the plate, gently rock, and return to 37  C incubator for 5 min. 4. During the incubation, prepare a 12-well plate with a PDL-coated coverslip in each well (see Note 11). Add 1 mL of culture media to each well. 5. Remove the cells from the incubator, and resuspend cells by adding 9 mL of culture media to the plate. Triturate to generate a single-cell suspension. 6. Add 25–100 μL of the single-cell suspension to each well depending on the experiment and cell line. 7. Return the cells to the incubator, and incubate under proper conditions for the cells to adhere and grow. 8. Once cells are adherent, 6–24 h after plating, replace media with media containing doxycycline at a final concentration of 1 μg/mL. 9. Visible eGFP expression should be detectable following 18–24 h of doxycycline treatment.

3.3.3 Sample Collection and Storage

1. Following 18–24 h of APO1-YTH expression (see Note 12), remove the media, and wash cells once in 1 PBS. 2. Fix cells with 4% PFA for 10 min at room temperature. 3. Wash cells once with 2 SSC and remove. 4. Add 70% ethanol to each well, and seal the plate with a layer of parafilm. Cells can now be stored at 4  C for up to 2 weeks (see Note 13).

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3.4 In Situ Detection of APO1-YTH-Induced C-to-U Mutations

1. Remove coverslips from 70% ethanol, and wash once in PBST.

3.4.1 Reverse Transcription

4. Preincubate coverslips with 1 RevertAid H-minus reverse transcription buffer at room temperature.

2. Incubate with 0.1 M HCl for 10 min at room temperature. 3. Wash coverslips twice in 1 PBS for 3 min each.

5. Place a layer of Whatman paper wet with 2 SSC into staining container, and cover with a layer of parafilm. Lines for coverslip placement can be drawn on parafilm using permanent marker. 6. Spot 50 μL of the RT mix onto parafilm for each coverslip. Place coverslips cell-side down onto mixture. Seal staining container with parafilm, and incubate at 37  C for 1 h. 3.4.2 RNA Digestion, PLP Hybridization, and Ligation

1. Remove coverslips from staining container, and wash twice in 1 PBS at room temperature. 2. Fix coverslips in 4% PFA for 10 min at room temperature. 3. Wash coverslips twice in PBST for 3 min each. 4. Preincubate coverslips in 1 Ampligase Ligase Reaction Buffer at room temperature. 5. Replace the layer of parafilm on the 2 SSC wet Whatman paper within the staining container. Lines for coverslip placement can be drawn on the parafilm using permanent marker. 6. Spot 50 μL of the hybridization-ligation mix for each coverslip onto the parafilm. Place coverslips cell-side down onto mixture. Seal staining container with parafilm, and incubate at 37  C for 30 min. 7. Incubate at 45  C for 45 min (see Note 6).

3.4.3 Rolling-Circle Amplification

1. Remove coverslips from staining container, and wash twice in PBST for 3 min per wash. 2. Preincubate with 1 Phi29 DNA Polymerase Reaction Buffer at room temperature. 3. Replace the layer of parafilm on 2 SSC wet Whatman paper within the staining container with a fresh piece of parafilm. Lines for coverslip placement can be drawn on the parafilm using permanent marker. 4. Spot 50 μL of the RCA mix for each coverslip onto the parafilm. Place coverslips cell-side down onto mixture. Seal staining container with parafilm, and incubate at 37  C for 60–100 min. RCA duration should be established per probe set to achieve punctate signal with limited RCP shearing. Overamplification and shearing can make it difficult to discriminate individual RNA molecules and lead to challenges during signal quantification (see Note 14).

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To simultaneously detect proteins of interest along with m6A-modified and unmodified transcripts, DART-FISH can be combined with immunofluorescence. We recommend optimizing DART-FISH detection prior to co-staining and including DARTFISH and immunofluorescence alone controls to best optimize this procedure. 1. Remove coverslips from the staining container, and wash twice with 1 PBS for 3 min each at room temperature. 2. Prepare the primary antibody dilution in PBST as recommended for each antibody. Replace the layer of parafilm in the staining container, and draw lines for sample placement with permanent marker. 3. Spot 50 μL of the primary antibody dilution onto the parafilm for each coverslip. Place coverslips cell-side down onto the primary antibody dilution, and incubate coverslips at 4  C overnight or for 1 h at room temperature. 4. Wash coverslips three times in 1 PBS for 3 min each. 5. Replace the layer of parafilm in the staining container, and draw lines for sample placement with permanent marker. 6. From this step onward, samples will be light-sensitive. Take precautions to ensure samples are protected from light during washes and incubations. Prepare the secondary antibody dilution in PBST as recommended for each secondary antibody. Spot 50 μL of the secondary antibody dilution onto the parafilm for each coverslip. Place coverslips cell-side down onto the secondary antibody dilution, and incubate coverslips for 45 min at room temperature. The time and dilution solution may need to be altered based on recommendations for specific secondary antibodies. 7. Remove coverslips, and wash three times in 1 PBS for 3 min each. 8. Fix coverslips in 4% PFA for 10 min at room temperature. 9. Wash coverslips twice in PBST. Proceed to detection probe hybridization.

3.4.5 Detection Probe Hybridization and DAPI Staining

From this step onward, light-sensitive dyes will be used. Take caution to protect detection probes and stained samples from light during handling, incubation, and storage. 1. Remove coverslips from staining container, and wash once with PBST for 3 min at room temperature. 2. Replace the layer of parafilm on 2 SSC wet Whatman paper within the staining container with a fresh piece of parafilm. Lines for coverslip placement can be drawn on the parafilm using permanent marker.

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3. Spot 50 μL of the detection probe hybridization mix for each coverslip onto the parafilm layer. Place coverslips cell-side down onto mixture. Seal the staining container with parafilm, and incubate at 37  C for 30 min. This mixture contains fluorescently labeled oligonucleotides that are light-sensitive; take caution to shield coverslips from light during the incubation. 4. Remove coverslips from the incubator, and wash once in PBST and twice in PBS for 3 min per wash. 5. Mount coverslips onto slides using Vectashield vibrance mounting media. 3.4.6 Image Acquisition and Analysis

1. Slides can be imaged once the mounting media is fully cured, generally after 24 h (Fig. 3). Slides are stored according to the mounting media manufacturer’s recommendations. For Vectashield vibrance, store slides at room temperature. 2. Images can be acquired using a variety of fluorescence microscopes. For initial optimization, we recommend a conventional wide-field epifluorescence microscope to maximize visible signal. 3. It is critical to ensure that proper filter cubes are used to avoid bleed through between individual channels. We utilize Cy3and Cy5-conjugated detection probes to visualize RNAs and Alexafluor488-conjugated secondary antibodies for co-staining using immunofluorescence. All of these dyes can be easily separated with conventional filter cubes (see Note 15). 4. Exposure times should be set to not saturate individual pixels and to allow individual punctate signal from each RCP. Overexposure can lead to challenges during signal quantification due to issues discriminating individual RNA molecules. 5. To capture the maximum number of RNAs, we acquire threedimensional z-stack spanning all focal planes for a given cell. Stacks can be quantified in three dimensions or as a 2D maximal projection. 6. Images can be quantified using a variety of open-source software packages such as ImageJ/FIJI, CellProfiler, and FISHQuant [15–17]. We use FIJI to first transform 3D z-stacks into maximum intensity projections (MIPs). MIPs are then input to CellProfiler to generate masks for individual cells. Individual mRNA molecules are quantified on a per channel basis in three dimensions within each individual cell using FISHQuant.

4

Notes 1. Previous protocols have recommended including between four and LNAs. However, in our experience, four LNAs were not sufficient to provide adequate signal. We have not tested >7 LNAs within a single primer.

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Fig. 3 Representative maximal projection images of DART-FISH targeting the ACTB A1222 m6A site in doxycycline-inducible APO1-YTH HeLa cells. Dotted lines represent the cell outlines. Scale bar ¼ 5 μm. Maximal projections were generated from z-stacks containing 62 (No Dox) or 43 (Dox) focal planes, each of 210 nm thickness. Images were acquired using the following exposure times per z-plane: Cy5/m6A1222, 15 ms (low noise); Cy3/A1222, 10 ms (low noise); DAPI, 50 ms (high well capacity)

2. The linker region within each padlock probe can either be poly (A) or random sequences. Each linker region should be designed to minimize the formation of strong deleterious secondary structures. 3. PLPs can also be ordered as unmodified oligonucleotides and phosphorylated as previously described [8, 9].

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4. For secondary antibodies, we utilize the Alexa-488 dye because following the DART-FISH protocol, the T2A-eGFP signal has been lost due to bleaching. Prior to co-staining for protein detection, ensure that the eGFP signal is no longer visible within your sample. 5. Alternative enzymes, such as alternative reverse transcriptases and ligases, have not been tested for use in DART-FISH; however, other enzymes have been described in other padlock probe rolling-circle amplification protocols [8, 9, 18]. 6. The same incubator can be used for 37  C and 45  C incubations. Allow the incubator to reach 45  C prior to starting the 45-min timer. We observed no difference in the number of detected transcripts between using a single or two different incubators. 7. We utilize an inverted wide-field Leica DMi8 microscope equipped with a 63/1.4 HC PL APO objective, Leica DFC9000 4.2 MP monochrome sCMOS camera, Lumencor SOLA SM light engine, and the following filter cubes: DAPI (EX350/50; DC400; EM460/50), GFP (EX470/40; DC495; EM525/50), RHOD (EX546/10; DC560; EM585/40), and Y5 (EX620/60; DC660; EM700/75). 8. To generate doxycycline-inducible APO1-YTHmut control cell lines, the described protocol can be used with the substitution of the pTLCV2 APO1-YTHmut-T2A-eGFP plasmid (Addgene #178950) during lentivirus production. 9. We recommend using a series of viral dilutions as well as a no infection control. Ideally, viral titer should be limiting and not infecting all cells in the dish to ensure low viral copy-number integration into your cell line of interest. Infection success is cell line-dependent and may require further optimization. 10. For serial dilution of single cells into a 96-well plate, this is a good resource: https://www.corning.com/catalog/cls/ documents/protocols/Single_cell_cloning_protocol.pdf. 11. Coverslips can be coated ahead of time and stored at 4  C for several months. 12. APO1-YTH-induced C-to-U mutations can be detected as early as 2 h after doxycycline treatment. We observe no differences in editing levels between 18 and 24 h of APO1-YTH expression [10]. Prolonged expression of APO1-YTH for longer than 72 h becomes detrimental to cell health. 13. Samples can alternatively be dehydrated with sequential 70%, 85%, and 99.5% ethanol washes for 3 min each. Dehydrated samples can be stored at 20  C for up to 2 weeks or at 80  C for long-term storage.

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14. Rolling-circle product shearing occurs from overamplification which can cause larger non-punctate fluorescent signal and lead to difficulty discriminating individual molecules. Shearing can be diminished by reducing the RCA incubation time. Alternatively, during quantification, single molecules can be discriminated by increasing the minimum distance between individual spots. 15. It is important to experimentally confirm no bleed through signal between individual channels as well as to test whether switching fluorophores changes the proportion of C and U transcripts detected. In our hands, DT3-Cy3 and DT2-Cy5 provide good agreement between DART-FISH %C2U values and those of DART-seq. References 1. Jiang X, Liu B, Nie Z et al (2021) The role of m6A modification in the biological functions and diseases. Signal Transduct Target Ther 6(1):74 2. Zaccara S, Ries RJ, Jaffrey SR (2019) Reading, writing and erasing mRNA methylation. Nat Rev Mol Cell Biol 20(10):608–624 3. Meyer KD, Jaffrey SR (2017) Rethinking m(6) a readers, writers, and erasers. Annu Rev Cell Dev Biol 33:319–342 4. Le P, Ahmed N, Yeo GW (2022) Illuminating RNA biology through imaging. Nat Cell Biol 24(6):815–824 5. Roost C, Lynch SR, Batista PJ et al (2015) and thermodynamics of Structure N6-methyladenosine in RNA: a spring-loaded base modification. J Am Chem Soc 137(5): 2107–2115 6. Sheehan CJ, Marayati BF, Bhatia J et al (2023) In situ visualization of m6A sites in cellular mRNAs. Nucleic Acids Res. https://doi.org/ 10.1093/nar/gkad787 7. Meyer KD (2019) DART-seq: an antibody-free method for global m(6)A detection. Nat Methods 16(12):1275–1280 8. Krzywkowski T, Nilsson M (2018) Padlock probes to detect single nucleotide polymorphisms. Methods Mol Biol 1649:209–229 9. Larsson C, Grundberg I, Soderberg O et al (2010) In situ detection and genotyping of individual mRNA molecules. Nat Methods 7(5):395–397 10. Tegowski M, Flamand MN, Meyer KD (2022) scDART-seq reveals distinct m(6)A signatures

and mRNA methylation heterogeneity in single cells. Mol Cell 82(4):868–878 e810 11. Petersen M, Wengel J (2003) LNA: a versatile tool for therapeutics and genomics. Trends Biotechnol 21(2):74–81 12. Untergasser A, Cutcutache I, Koressaar T et al (2012) Primer3–new capabilities and interfaces. Nucleic Acids Res 40(15):e115 13. Owczarzy R, Tataurov AV, Wu Y et al (2008) IDT SciTools: a suite for analysis and design of nucleic acid oligomers. Nucleic Acids Res 36 (Web Server issue):W163–W169 14. Stenberg J, Nilsson M, Landegren U (2005) ProbeMaker: an extensible framework for design of sets of oligonucleotide probes. BMC Bioinformatics 6:229 15. Schindelin J, Arganda-Carreras I, Frise E et al (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9(7): 676–682 16. Carpenter AE, Jones TR, Lamprecht MR et al (2006) CellProfiler: image analysis software for identifying and quantifying cell phenotypes. Genome Biol 7(10):R100 17. Tsanov N, Samacoits A, Chouaib R et al (2016) smiFISH and FISH-quant – a flexible single RNA detection approach with super-resolution capability. Nucleic Acids Res 44(22):e165 18. Lundin E, Wu C, Widmark A et al (2020) Spatiotemporal mapping of RNA editing in the developing mouse brain using in situ sequencing reveals regional and cell-type-specific regulation. BMC Biol 18(1):6

Chapter 11 Multiplexed Immunofluorescence and Single-Molecule RNA Fluorescence In Situ Hybridization in Mouse Skeletal Myofibers Lance T. Denes, Chase P. Kelley, and Eric T. Wang Abstract RNA fluorescence in situ hybridization (FISH) is a powerful method to determine the abundance and localization of mRNA molecules in cells. While modern RNA FISH techniques allow quantification at single molecule resolution, most methods are optimized for mammalian cell culture and are not easily applied to in vivo tissue settings. Single-molecule RNA detection in skeletal muscle cells has been particularly challenging due to the thickness and high autofluorescence of adult muscle tissue and a lack of in vitro models for mature muscle cells (myofibers). Here, we present a method for isolation of adult myofibers from mouse skeletal muscle and detection of single mRNA molecules and proteins using multiplexed RNA FISH and immunofluorescence. Key words RNA, RNA FISH, RNA localization, Fluorescence imaging, Skeletal muscle, Myofiber, Sarcomere

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Introduction Fluorescence in situ hybridization (FISH) is a widely used technique to visualize the abundance and localization of nucleic acids in cells and tissues [1]. The method relies on the hybridization of labeled oligonucleotide probes to a DNA or RNA sequence of interest. Original in situ hybridization methods used radioactive probes or indirect immunofluorescence as detection modalities, which allowed visualization of tissue-scale expression patterns of genes during development but had a variety of drawbacks and achieved limited spatial resolution [2, 3]. A major breakthrough in RNA imaging came when technical improvements in quantitative fluorescence microscopy were combined with direct fluorescent labeling of oligonucleotide probes. This allowed visualization of mRNAs in single cells with single-molecule resolution for the first time [4]. This technique, referred to as single-molecule RNA FISH

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(smFISH), targets multiple fluorescently labeled probes to an mRNA of interest, concentrating the signal in a diffraction-limited point source at the transcript, which is visualized as a spot. Spots are counted, and the intensity of individual spots is quantified and used to determine the number of molecules present in larger, high copy number locations such as transcription sites and RNA granules [1]. The first smFISH methods used a small number [5–10] of long, multiple internally labeled probes that were expensive to synthesize [4]. Later work optimized the method to work with many (24–48) short, terminally labeled probes, which are more cost-effective and can be obtained from commercial sources [5]. Current methods have improved on the direct probe labeling approach by appending “readout” sequences to unlabeled primary probes and performing a secondary hybridization of labeled readout probes [6]. This further improves cost efficiency and has been combined with clever sequential readout strategies to multiplex smFISH at transcriptome scale [7, 8]. Single-molecule RNA FISH is now widely used and has led to many discoveries involving single-cell gene expression, mechanisms of mRNA processing, and subcellular localization of mRNAs [9–12]. While smFISH has been very successful in cultured cells, it is still a challenge to apply to in vivo tissues. This is largely due to the high levels of background autofluorescence in tissues that precludes single-molecule detection and difficulty of probe access in thick specimens. Recently, several signal amplification approaches and tissue clearing methods have been applied to overcome these problems [13–15]. Here, we focus on signal amplification, as clearing methods are more complicated and highly tissue-specific. The hybridization chain reaction (HCR) is a simple and effective method for RNA FISH signal amplification that has been used by multiple labs to visualize single mRNAs in mouse myofibers [16, 17]. In HCR FISH, an “initiator” sequence is appended to primary probes. After initial hybridization of primary probes to target mRNAs in situ, two fluor-labeled hairpin readout probes are added in an amplification step. The initiator sequence opens the hairpins and starts the chain reaction, which results in a linear accumulation of labeled readout probes at the target transcript. In the recent HCR v3.0 strategy, pairs of primary probes with a split initiator sequence suppress off-target amplification events by requiring adjacent binding of two primary probes on the target transcript to nucleate HCR [15]. While the unique physiology of skeletal muscle cells and the large variety of myopathies associated with mRNA misprocessing make skeletal muscle an attractive target for studies of mRNA localization, the lack of robust protocols for mRNA visualization in skeletal muscle cells has hindered work in this area. Skeletal muscle tissue is an especially challenging context for RNA imaging. Muscle tissue is a complex organ comprised of bundles of skeletal

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muscle cells (myofibers) surrounded by a various connective tissues, nerves, and blood vessels. The myofibers themselves are large, multinucleated syncytia hundreds of microns in diameter and centimeters in length. All these features make RNA imaging in muscle a challenge. Here, we present a protocol for smFISH in mouse skeletal muscle, the extensor digitorum longus (EDL), that combines single myofiber isolation with HCR-based signal amplification. Isolating myofibers away from the complex muscle tissue improves interpretability of images and reduces background, while HCR amplification of FISH signal facilitates single-molecule mRNA detection (Fig. 1). Finally, we incorporate

Fig. 1 Myofiber RNA FISH workflow. Mice are euthanized according to institutional protocols and EDL muscles are dissected from each leg. Muscles are digested with collagenase and flushed with warm media to dissociate myofibers. Myofibers are collected into culture dishes and either transferred to spot plates for immediate fixation or kept in ex vivo culture for up to 3 days. IF staining is performed in spot plates if desired followed by post-IF fixation. Overnight hybridization of HCR RNA FISH probes followed by signal amplification is performed in spot plates. Myofibers are then mounted on slides for confocal imaging

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immunofluorescence staining into the protocol, which allows RNA positions to be evaluated relative to subcellular structures.

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Materials

2.1 Isolation and Culture of Mouse Myofibers

1. Myofiber culture media: 20% fetal bovine serum (FBS) in DMEM (high glucose, GlutaMAX Supplement, pyruvate), 1% Pen/Strep. Filter sterilize with 0.2 μm PES filter unit before use. Store at 4 °C and aliquot before use. 2. Myofiber digestion solution: 0.2% collagenase type I in DMEM. Filter sterilize with 0.2 μm PES filter unit after dissolving collagenase. 1–2 mL required per EDL, prewarm to 37 °C before use. Extra solution can be aliquoted and stored at -20 °C. 3. Horse serum-coated dishes: Coat all dishes that will be used for an experiment with horse serum to prevent myofibers sticking to dishes. For one EDL, prepare a 6 cm dish for digestion, a 6 cm dish for dissociation, a 6 cm dish for collection, and a 6 well dish if myofibers will be cultured or treated in downstream experiments. Add 3 mL horse serum per 6 cm dish, incubate at 37 °C for at least 15 min, aspirate serum, and replace with digestion or culture media. Keep plates with media in the incubator prior to use. 4. Mice: At least two mice per experiment to account for failed dissection or myofiber damage. We use an even split of male and female mice for all published experiments. Use background strain appropriate to experimental goals. We have used C57BL/6 and FVB strains with similar results. Less than 1-year-old mice are desirable as muscle tissue is more difficult to dissociate and more auto-fluorescent in aged mice. 5. Sterile disposable filter units, 250 mL PES 0.2 μm. 6. Collagenase type I. 7. Horse serum. 8. Standard dissection tools (scissors, forceps, pins, pinboard). 9. Stereoscopic dissection microscope. 10. 37 °C, 5% CO2 sterile tissue culture incubator. 11. Tissue culture-treated dishes (35 mM, 6 cM, 6-well, or similar).

2.2 RNA FISH and Immunofluorescence

1. RNase Zap. 2. Pyrex spot plates. 3. Sterile PBS, pH 7.4. 4. Paraformaldehyde (PFA), 16% ampules.

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Fig. 2 HCR strategy for RNA FISH signal amplification. Primary probes are hybridized to the transcript of interest. These probes contain an initiator sequence at the 3′ end that is complementary to the HCR amplifiers. The HCR amplifiers consist of two fluor-labeled oligonucleotide hairpins. Hairpin 1 binds the initiator sequence, which opens the hairpin and allows binding to hairpin 2 to begin the hybridization chain reaction. This results in linear accumulation of fluorescent signal at the mRNA of interest

5. Ultrapure 20× SSC. 6. Probe hybridization buffer (Molecular Instruments) (see Note 1). 7. Probe wash buffer (Molecular Instruments) (see Note 1). 8. Amplification buffer (Molecular Instruments) (see Note 1). 9. Primary oligonucleotide HCR probes (Molecular Instruments) (see Note 2, Fig. 2). 10. HCR amplifiers (Molecular Instruments) (see Note 2, Fig. 2). 11. Antibodies for protein of interest (see Note 3). 12. NxGen RNase inhibitor. 13. Ultrapure RNase-free BSA (see Note 4). 14. Glass slides. 15. Coverslips. 16. ProLong Diamond Antifade Mountant. 2.3

Staining Buffers

1. Fixation buffer: 4% PFA in 1× PBS. 2. IF wash buffer (PBST): 0.1% Tween-20 in 1× PBS. 3. Permeabilization: 1.0% Triton X-100 in 1× PBS. 4. Blocking buffer: 0.1% Tween-20, 1% Ultrapure BSA, 1 U/μL RNase inhibitor in 1× PBS. 5. FISH wash buffer 1: (SSCT): 5× SSC, 0.1% Tween-20. 6. FISH wash buffer 2: 2× SSC. 7. DAPI staining solution: 1 μg/mL DAPI in 1× PBS.

2.4 Imaging and Image Analysis

1. Confocal microscope with appropriate laser lines, detectors, and high NA (1.4) oil objective at least 40× magnification. 2. Computer with image analysis software installed (FIJI, Imaris, Python).

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Methods Perform all steps at room temperature unless otherwise noted.

3.1 Myofiber Isolation

1. Prepare media, dissociation solution, and plates for myofiber isolation (see Note 5). 2. Euthanize animal according to institutional regulations. 3. Spray down mouse hindlimbs of euthanized animal with 70% EtOH and pin animal face up (see Note 6). 4. Cut the skin at the ankle and peel toward knee to reveal the underlying hindlimb muscles. 5. Expose the tendons connected to the tibialis anterior (TA) and EDL at the base of each hindlimb. The two tendons will be on top of one another, and after revealing the tendons, the forceps should be able to slide in between them. 6. Snip the overlying TA tendon with small scissors and lift the TA to reveal the EDL beneath it. Take care not to disrupt the EDL tendons. If the EDL tendon is cut, the muscle can still be isolated if there is no damage to the muscle body. Carefully grab the EDL tendon with forceps and peel the EDL from the underside of the TA. 7. Cut the TA at the proximal end. Because the proximal TA tendon is buried in the knee, it will be difficult to remove the TA at the tendon. Get as close as possible to the knee and cut the muscle with scissors, taking care not to cut the EDL beneath it. By removing the TA, it will be easier to access the proximal tendon of the EDL. 8. Snip the distal EDL tendon as close as possible to the insertion if it is not already cut, and then snip the proximal EDL tendon as close as possible to the insertion. 9. Carefully transfer the EDL via the tendons to the dish containing digestion solution. Take care not to touch the muscle body or the myofibers will be damaged and hyper-contract during isolation.

3.2 Dissociation and Isolation of Myofibers from EDL Muscle (see Note 6)

1. Incubate the EDL in digestion solution at 37 °C for 20 min. Begin checking the tissue every 10 min for dissociation. Incubation times may vary depending on collagenase activity, the age of the mouse, and the muscle used. When dissociation is complete, the tissue should appear looser with space between bundles of myofibers becoming visible and individual fiber ends separating from the tissue. 2. Gently transfer the tissue to the dissociation dish containing myofiber culture media with a large bore pipette tip, taking care not to damage the tissue. Begin gently flushing the media over

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the tissue with a 1 mL pipette. Use enough force to dislodge individual fibers from the tissue but try to avoid causing damage and hyper-contraction. Do not triturate the tissue and aspirate it into the pipette. If fibers are not easily separating, return the tissue to the dish containing digestion media and incubate another 10–20 min. Return to the dissociation dish and resume flushing. Repeat as many times as is necessary depending on the myofiber yield required. 3. As fibers separate from the tissue, transfer them immediately to the collection dish containing myofiber culture media with the 1 mL pipette. Place this dish in the 37 °C incubator as soon as possible. If many tissues are being processed, prepare a dish for each to avoid prolonged incubation of collected myofibers at room temperature. 4. If drug treatments or pulse-chase experiments are to be performed, myofibers can be transferred to a horse serum coated 6 well (or other) dish. 3.3 Immunofluorescence and Single-Molecule RNA FISH Staining of Myofibers (see Note 7)

1. Transfer myofiber in media into a well of a spot plate. Remove media and replace with fixation buffer. Incubate for 10 min. Then remove PFA and wash thrice with PBS. 2. Replace PBS with permeabilization buffer. Incubate for 20 min. Wash thrice with PBS. 3. If performing IF, replace PBS with blocking buffer. Incubate for 30 min. If not performing IF, skip to step 11. 4. Remove blocking buffer and replace with primary antibody diluted into blocking buffer (consult antibody manufacturer for dilution). Incubate for 2 h to overnight at room temperature. We found that incubation at 4 °C does not allow sufficient antibody penetration throughout the myofiber. We obtain even staining throughout the fiber with overnight incubations at room temperature. However, for some antibodies or if only near-surface staining is required, a shorter incubation is sufficient (see Note 3). 5. Wash thrice 10 min with PBST at room temperature. 6. Remove PBST and replace with appropriate secondary antibody diluted into blocking buffer (consult manufacturer for dilution). Incubate for 2 h to overnight at room temperature. Typically, 2 h is sufficient for staining, but calibrate this for each new antibody used if full fiber staining is required. 7. Wash thrice 10 min with PBST. 8. Wash twice 10 min with PBS. 9. Fix the antibody-stained myofiber with fixation buffer. 10. Wash thrice 5 min with PBS. 11. Replace final wash with 2× SSC and incubate for 5 min.

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12. Pre-hybridize samples with prewarmed (37 °C) probe hybridization buffer in oven at 37 °C for 30 min (see Note 8). 13. Dilute probes to 1 nM in prewarmed probe hybridization buffer (1:1000 from Molecular Instruments stock tubes) (see Note 9). 14. Incubate samples overnight in probe solution in humidified oven at 37 °C. 15. Remove hybridization solution and wash once with 400 uL prewarmed probe wash buffer. Vigorously flush the wash buffer over the myofibers to dissolve all the leftover viscous hybridization buffer. 16. Wash additional four times for 10 min in prewarmed probe hybridization buffer at 37 °C. 17. Wash twice for 5 min with SSCT. 18. Incubate in 200 μL amplification buffer for 30 min. 19. During incubation in amplification buffer, prepare the HCR amplifier oligos. Aliquot desired amount of HCR amplifiers into separate PCR tubes, heat to 95 °C for 90 s, and allow to cool to room temperature protected from light. Use 2 μL of each amplifier per 100 μL of amplification buffer. Use 200 μL amplification buffer per well of spot plate (see Note 10). 20. Dilute amplifiers into amplification buffer immediately before adding to samples, vortex, and add to samples. 21. Amplify at room temperature in humidified chamber. Amplification time depends on application. For smFISH in myofibers, amplify for 4 h (see Note 11). 22. Wash five times for 10 min in SSCT. 23. Wash once with DAPI staining buffer for 20 min. 24. Mount on slide using ProLong Diamond (or similar) and appropriate coverslip for imaging system (see Note 12). 3.4 Imaging and Image Analysis

1. Mount slides on confocal microscope and locate myofibers using brightfield or DAPI channel with low magnification objective. 2. Switch to high NA oil objective >40× and focus on myofiber region of interest (see Note 13). 3. Acquire Z-stacks spaced > Python 3. The following actions can be used as well: Put the file in the Jupyter Notebook working directory, and then create a new .ipynb file in Jupyter Notebook; type %load xxxx.py in the newly created file (“xxx” is the file name). 3. “mRNA on a surface” mainly points to the study of mitochondria-localized mRNA. We defined a plane where the beads are not formed under it to model the mRNA conformation on a mitochondrial surface. To conduct the simulation, the x-y plane was set up as the surface so that the z coordinate would never be less than zero. 4. When running all the visualization-related code (see Subheading 3.2), there may be a problem where the generated 3D model cannot be dragged. Please add the line “%!matplotlib widget” after the “import copy” code line. You can also add this line of code to the code in Subheading 3.1 as well so that more coordinates can be displayed. 12.import copy 13.%matplotlib widget

5. “Visualization mRNA with or without ribosomes” is a simulation code that generates one strand of input nucleotide length of RNA onto 3D Cartesian coordinates. Choosing a different number of ribosomes will yield different results. 6. “mRNA on a surface” and “Visualization mRNA on a surface” work the same, except the latter simulates RNA that is localized onto a surface. 7. “Visualization of mRNA on surface with plane” is the visualization of mRNA on a surface displayed along with its reference plane z = 0. 8. Please set the number of iterations to be greater than 200.

Acknowledgments This work was supported by startup funds from Tsinghua SIGS (to T.T.). T.T. also acknowledges support from the Jilin Fuyuan Guan Food Group Co., Ltd., the Science, Technology, and Innovation Commission of Shenzhen Municipality (WDZC20220811144737001).

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References 1. Jackson RW, Smathers CM, Robart AR (2023) General strategies for RNA X-ray crystallography. Molecules 28(5):2111 2. Furtig B, Richter C, Wohnert J, Schwalbe H (2003) NMR spectroscopy of RNA. Chembiochem 4(10):936–962 3. Vallina NS, McRae EKS, Hansen BK et al (2023) RNA origami scaffolds facilitate cryoEM characterization of a broccoli-pepper aptamer FRET pair. Nucleic Acid Res 51(9): 4613–4624 4. Zuker M (2003) Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acid Res 31(13):3406–3415 5. Lorenz R, Bernhart SH et al (2011) ViennaRNA package 2.0. Algorithms Mol Biol 6(1):26 6. Knudsen B, Hein J (2003) Pfold: RNA secondary structure prediction using stochastic context-free grammars. Nucleic Acid Res 31(13):3423–3428 7. Do CB, Woods DA, Batzoglou S (2006) CONTRAfold: RNA secondary structure prediction without physics-based models. Bioinformatics 22(14):E90–E98 8. Wang LY, Liu YN, Zhong XD et al (2019) DMfold: a novel method to predict RNA secondary structure with pseudoknots based on deep learning and improved base pair maximization principle. Front Genet 10:143 9. Zhang H, Zhang CH, Li Z et al (2019) A new method of RNA secondary structure prediction based on convolutional neural network and dynamic programming. Front Genet 10 10. Fu LY, Cao YX, Wu J et al (2022) UFold: fast and accurate RNA secondary structure prediction with deep learning. Nucleic Acid Res 50(3):e14–e14 11. Townshend RJL, Eismann S, Watkins et al (2021) Geometric deep learning of RNA structure. Science 373(6558):1047–1051 12. Mao YH, Liu HL, Liu YL et al (2014) Deciphering the rules by which dynamics of mRNA secondary structure affect translation efficiency in Saccharomyces cerevisiae. Nucleic Acid Res 42(8):4813–4822 13. Kawaguchi D, Shimizu S, Abe N et al (2020) Translational control by secondary-structure

formation in mRNA in a eukaryotic system. Nucleosides Nucleotides Nucleic Acids 39(1–3):195–203 14. Lin JA, Chen Y, Zhang YP et al (2022) Deciphering the role of RNA structure in translation efficiency. BMC Bioinformatics 23(3): 1–15 15. Mustoe AM, Busan S, Rice GM et al (2018) Pervasive regulatory functions of mRNA structure revealed by high-resolution SHAPE probing. Cell 173(1):181–195 16. Guo T, Modi OL, Hirano J et al (2022) Singlechain models illustrate the 3D RNA folding shape during translation. Biophys Rep 2(3) 17. Imataka H, Gradi A, Sonenberg N (1998) A newly identified N-terminal amino acid sequence of human eIF4G binds poly(A)binding protein and functions in poly(A)dependent translation. EMBO J 17(24): 7480–7489 18. Wells SE, Hillner PE, Vale RD et al (1998) Circularization of mRNA by eukaryotic translation initiation factors. Mol Cell 2(1): 135–140 19. Adivarahan S, Livingston N, Nicholson B et al (2018) Spatial organization of single mRNPs at different stages of the gene expression pathway. Mol Cell 72(4):727–738 20. Koch A, Aguilera L, Morisaki T et al (2020) Quantifying the dynamics of IRES and cap translation with single-molecule resolution in live cells. Nat Struct Mol Biol 27(12): 1095–1104 21. Khong A, Parker R (2018) mRNP architecture in translating and stress conditions reveals an ordered pathway of mRNP compaction. J Cell Biol 217(12):4124–4140 22. Tsuboi T, Viana MP, Xu F et al (2020) Mitochondrial volume fraction and translation duration impact mitochondrial mRNA localization and protein synthesis. elife 9:e57814 23. Jourdren L, Delaveau T, Marquenet E et al (2010) CORSEN, a new software dedicated to microscope-based 3D distance measurements: mRNA-mitochondria distance, from single-cell to population analyses. RNA 16(7): 1301–1307

Part II DNA FISH

Chapter 14 Combined 3D DNA FISH, Single-Molecule RNA FISH, and Immunofluorescence Souvik Sen, Shivnarayan Dhuppar, and Aprotim Mazumder Abstract Nuclear architecture is a potential regulator of gene expression in eukaryotic cells. Studies connecting nuclear architecture to gene expression are often population-averaged and do not report on the cell-level heterogeneity in genome organization and associated gene expression. In this report we present a simple way to combine fluorescence in situ hybridization (FISH)-based detection of DNA, with single-molecule RNA FISH (smFISH) and immunofluorescence (IF), while also preserving the three-dimensional (3D) nuclear architecture of a cell. Recently developed smFISH techniques enable the detection of individual RNA molecules; while using 3D DNA FISH, copy numbers and positions of genes inside the nucleus can be interrogated without interfering with 3D nuclear architecture. Our method to combine 3D DNA FISH with smFISH and IF enables a unique quantitative handle on the central dogma of molecular biology. Key words Fluorescence in situ hybridization (FISH), 3D DNA FISH, Single-molecule RNA FISH (smFISH), Immunofluorescence (IF)

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Introduction Nuclear architecture is the non-random, reproducible organization of the genome in terms of its inter- and intrachromosomal contacts and also its interactions with different nuclear compartments such as the nuclear lamina or nucleoli. The majority of our knowledge on nuclear architecture and associated gene regulation comes from bulk assays such as chromosome conformation capture [1] and PCR-based assays. These assays, while reporting accurately on population level means, fail to report on the cell-to-cell heterogeneity of genome organization or gene expression. According to a recent report [2], there is extensive heterogeneity in spatial genome organization within a population to the extent that at a time the average interactions captured at the population level are harbored by only a small percentage of cells within that population. This makes it all the more important to study nuclear architecture-dependent gene

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regulation at a single-cell and single-allele level. Other studies have correlated fluorescence in situ hybridization (FISH)-based detection of gene positions with bulk changes in mRNA expression [3]. In this report, we present a simple and reliable way to combine FISH-based detection of target DNA, with single-molecule RNA FISH (smFISH) and immunofluorescence (IF), while also preserving the 3D nuclear architecture of a cell. It should be noted that ours is not the first method for combining DNA and RNA FISH and IF [4–9], but we perform single-molecule level detection of absolute RNA counts at a single cell resolution along with 3D DNA FISH and IF, as described previously [10]. In the past two decades, smFISH for RNA has emerged as a highly sensitive technique to detect the gene expression by yielding absolute mRNA counts in cells and tissues instead of relative intensities as done in standard in situ hybridization (ISH) techniques [11–16]. In the version of this method used in our laboratory, 40 or more short (20 nt) singly labeled oligonucleotide probes are used against specific mRNA to make each mRNA molecule stand out as an identifiable diffraction-limited fluorescence spot [12–15]. Although there are methods that have been developed to combine DNA FISH and RNA FISH [4, 5, 17], fewer attempts have been made to combine 3D DNA FISH and smFISH. Conventional DNA FISH protocols often involve extensive ethanol dehydration steps that lead to destruction of 3D architecture of the cell by flattening them out. The nucleus is most affected by such dehydration as its water content can be as high as 85% [18]. High formamide- or acid-based treatments for DNA denaturation, and steps like nitrogen freeze-thaw cycles, also may be employed in existing DNA FISH protocols, which can adversely interfere with immunofluorescence or smFISH signals, thus making the methods difficult to combine [7, 19–21]. Here we describe a simple protocol (Fig. 1) to combine 3D DNA FISH, smFISH, and immunofluorescence. We perform immunostaining first and refix the cells with 4% paraformaldehyde and then proceed for 3D DNA FISH finally followed by smFISH. The individual protocols remain largely the same. Our protocol excludes extensive ethanol dehydration steps to keep the 3D nuclear architecture intact. The lesser efficiency of probe penetration can be compensated by longer incubation times (> = 24 h). Unlike other 3D DNA FISH protocols, our protocol does not have nitrogen freeze-thaw cycles either. The specified sequence of the three techniques in the protocol is necessary to preserve the immunofluorescence and smFISH signals. If not performed in the correct order, the high formamide concentration in DNA FISH buffers can adversely affect smFISH RNA signals. Thus, we can get DNA FISH, RNA FISH, and IF signals in the same experiment at a single cell and single allele resolution (Figs. 2 and 3). The positions of each allele can be determined with respect to nuclear landmarks like

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Fig. 1 Workflow for the protocol. Fixation, permeabilization, denaturation, and washing steps are done in a 35 mm dish. Antibody incubations and probe hybridizations steps are done on a glass slide inside a humidified chamber. The humidified chamber is a 90 mm culture plate. The lid should be closed (aluminum foil may be attached on the lid to make the chamber dark) and sealed with parafilm before incubation. Silicone vial stoppers act as spacers to separate the slide from the soaked tissues

nuclear envelope or the nucleoli – this information would be largely lost in dehydrated cells, which are flattened against the coverslip. Further, beyond transcript counts on a cell-by-cell basis, it may be possible to determine the transcriptional status of each allele from the colocalization of the DNA FISH (gene) and single-molecule RNA FISH (transcript) signals (see Figs. 2 and 3 for examples).

2 Materials Prepare all general solutions and buffers using nuclease-free (not DEPC-treated) water and store these at 4 °C. Always warm formamide-containing buffers to room temperature before use. Clean the work benches and pipettes with RNaseZAP or RNase AWAY. Make sure that all plasticware like pipette tips and centrifuge tubes are DNase-/RNase-free. Pipette tips with a filter barrier to reduce chances of RNase contamination.

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Fig. 2 Combined 3D DNA FISH, smFISH, and immunofluorescence for CCNA2/Cyclin A2 in HeLa cells. (a) Nuclei are stained with DAPI (blue). (b) 3D DNA FISH signal for the CCNA2 gene position (red). Three copies of CCNA2 genes are observed in HeLa cells. (c) smFISH signal for CCNA2 mRNA (yellow). Each yellow spot represents a single CCNA2 mRNA molecule. (d) Cyclin A2 protein expression in HeLa cells (green). (e) Merged image for DNA FISH, smFISH and DAPI; IF image is not used in the merge. Cell to cell variability in mRNA and protein expression is clearly visible between two cells, and expression of mRNA and protein is correlated for CCNA2. Scale bar: 10 μm 2.1

Cell Culture

1. 22 × 22 mm cover slips. 2. 1 mg/mL poly-D-Lysine. 3. 35 mm plastic bottom dish. 4. Cell culture media: McCoy 5A, DMEM/F12 or appropriate culture medium supplemented with 10% Fetal Bovine Serum (FBS) and 1% Pen Strep Glutamine. 5. 1× PBS: Made in Milli-Q water and autoclaved. 6. 0.25% Trypsin EDTA. 7. Cell Lines: We have used U2OS and HeLa cell lines as examples for our experiments. 8. Forceps.

2.2 Immunofluorescence

1. 200 mM vanadyl ribonucleoside complex (VRC) (used as a RNase inhibitor). 2. 50 mg/mL Ultrapure Bovine Serum Albumin (BSA).

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Fig. 3 Combined 3D DNA FISH, smFISH, and immunofluorescence for TP53/p53 in U2OS cells. (a) Nuclei are stained with DAPI (blue). (b) 3D DNA FISH signal for the TP53 gene position. Two copies of TP53 genes are observed in U2OS cells (red). (c) smFISH for p53 mRNA (yellow). Each yellow spot represents a single TP53 mRNA molecule. (d) p53 Protein expression in U2OS cell (green). p53 protein levels are usually low in the absence of DNA damage – contrast has been adjusted to reveal nuclear p53 signal. (e) Merged image for DNA FISH, smFISH and DAPI; IF image is not used in the merge. Scale bar: 10 μm

3. 1× PBS: Add 5 mL of 10× nuclease-free PBS from stock in 45 mL of nuclease-free water in a 50 mL centrifuge tube to make 50 mL 1× PBS. Store at 4 °C. 4. 4% PFA in 1× PBS: Weigh 0.6 g of paraformaldehyde (PFA). Add it in 1.5 mL 10× nuclease-free PBS and nuclease-free water to make the final volume 15 mL. Heat and stir the solution with a magnetic stirrer to dissolve the PFA completely. Cool, and store at 4 °C. 5. 0.3% Triton X-100 in 1× PBS: Make a 10% intermediate stock of Triton X-100 by adding 1.5 mL Triton X-100, 1.5 mL 10× PBS, and nuclease-free water to make the final volume of 15 mL. Store at 4 °C. From this stock make 0.3% Triton X-100 in 1× PBS fresh before experiment. 6. Blocking solution: 1 mg/mL Ultrapure BSA, 2 mM VRC in 1× PBS (make in nuclease-free water). Always make it fresh. The blocking step may be redundant for some antibodies [22], but all antibody staining steps are performed in this blocking solution. VRC may be redundant but may be used if degradation of RNA signals is observed.

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7. Primary antibodies: Appropriate primary antibodies may be used (here, Anti-p53 mouse antibody and Anti-Cyclin A2 Rabbit antibody are used as examples). 8. Secondary antibodies: Appropriate fluorophore-labeled antibodies may be used (e.g., Goat Anti-mouse antibody tagged with Alexa Fluor 594 and Goat anti-rabbit antibody tagged with Alexa Fluor 488). 9. Humidified Chamber: Take a 90 mm petri dish. Attach aluminum foil on the lid to make the chamber dark. Put a Kimwipe inside the petri dish and soak it with water and keep two rubber silicone vial stoppers (or other spacers that can keep the glass slide slightly above the base of the petri dish). Place the glass slide on these. 10. Forceps and a 26G 0.5 inch needle. 2.3

DNA FISH

1. 20× SSC (nuclease-free) (main stock). 2. 100% deionized-formamide (main stock). 3. 2× SSC: dilute 20× SSC 1:10 in nuclease-free water. Store at 4 °C. 4. Denaturation Solution: 70% formamide, 2× SSC. 5. Hybridization buffer: We use Empire Genomics Hybridization buffer. 6. DNA FISH probe against gene of interest (Empire Genomics) (e.g., TP53 or CCNA2). 7. DNA FISH wash buffer: 50% formamide in 2× SSC. 8. Block heater. 9. Incubator at 80 °C. 10. Humidified Chamber: Take a 90 mm petri dish. Attach aluminum foil on the lid to make the chamber dark. Put a Kimwipe inside the petri dish and soak it with DNA FISH wash buffer and keep two rubber silicone vial stoppers (or other spacers that can keep the glass slide slightly above the base of the petri dish). Place a glass slide on these. Seal the petri dish with parafilm. Design of the humidified chamber can vary, but it is important to ensure that the cells do not dry out.

2.4

RNA FISH

1. Stellaris hybridization buffer (LGC Biosearch Technologies). 2. Fluorophore-labeled smFISH probes against mRNA of interest (LGC Biosearch Technologies) (e.g., TP53 or CCNA2. We use CAL Fluor 610 or Quasar 570 labeled probes). 3. RNA FISH wash buffer: 10% formamide in 2× SSC. Store in 4 °C. 4. 1 mg/mL DAPI in nuclease-free water (stock solution for staining DNA). Used at a final concentration of 1 μg/mL.

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5. Humidified Chamber: Make the humidified chamber similar to the DNA FISH humidified chamber, just soak the Kimwipe with RNA FISH wash buffer instead of DNA FISH wash buffer. 6. Vectashield (Vector Laboratories) (for mounting). 2.5 Imaging and Image Analysis

1. Microscope: A widefield epifluorescence microscope with a 60× oil immersion objective (1.42 NA) and a Retiga 6000 CCD camera is used for imaging. 2. Image analysis: Custom-written MATLAB codes are used for image and data analysis.

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Methods The complete protocol for combined DNA FISH, RNA FISH, and immunofluorescence can be executed in a total of 3 days. On the first day, fixation, permeabilization, and the whole immunostaining protocol can be done in 3 h. Then the cells have to be kept for 24 h for DNA FISH hybridization. On the second day, after 24 h (or appropriate duration depending on the probes) of DNA FISH hybridization, the RNA FISH protocol can be started. On the last day after two washes, the coverslip can be mounted for imaging. The protocol is described for adherent cells grown in culture (Fig. 1). The cells are grown on coverslips to minimize volumes of buffers and antibodies/probes required. If these are not limiting, the protocol can be modified for glass-bottom confocal dishes too by scaling up volumes appropriately. Single-molecule RNA FISH probes are designed in Stellaris probe designer (LGC Biosearch Technologies). Each probe is 20 nucleotides long and carries a single fluorophore conjugated through a 3′ amine modification and is HPLC-purified. At least 40 such probes are used against each target. Five nanomoles of the probes are resuspended in nuclease-free water to achieve a stock concentration of 10 μM for RNA FISH probes. These are typically used at a 1:20 to 1:30 dilution. Ready-made DNA FISH probes were procured from Empire Genomics. The stock probes (the concentration of which can vary slightly from batch to batch) were used at a 1:10 dilution.

3.1

Cell Culture

1. Coat 22 × 22 mm coverslips with 0.1 mg/mL Poly-D-Lysine (diluted in autoclaved Milli-Q) for 6–8 h. Then wash them thoroughly five to six times with autoclaved Milli-Q water to remove extra poly D-Lysine. Let it dry and after that keep it inside a 35 mm plastic bottom dish (see Note 1). 2. Pour appropriate cell culture medium in the 35 mm dish. Seed trypsinized cells (e.g., HeLa (Fig. 2) or U2OS (Fig. 3) cells) on the coverslip that is sitting inside the 35 mm dish. Make sure the coverslip is at the bottom of the dish and not floating on the media. 3. Allow the cells to grow for at least 24 h before fixation.

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3.2 Immunofluorescence

1. Wash the coverslip twice with 1× PBS. 2. Fix the cells with 4% PFA in 1× PBS for 15 min at room temperature (see Note 2). 3. After fixation, wash the coverslip twice with 1× PBS (see Note 3). 4. Permeabilize the cells with 0.3% Triton X-100 in 1× PBS for 10 min at room temperature (see Note 2). 5. Wash twice with 1× PBS. 6. Prepare 30 μL blocking solution fresh before use. Clean a glass slide properly with RNaseZap and let it dry for some time. Put the blocking solution on the glass slide. Then gently place the coverslip over it with a pair of forceps (cleaned with RNaseZap). Make sure that the side of the coverslip with cells faces the blocking solution. Keep it at room temperature inside a heavily humidified chamber for 30 min (see Note 4). 7. Prepare 30 μL of primary antibody mixture in a 1.5 mL centrifuge tube with appropriate primary antibody in 1:300 dilution in blocking solution. Vortex the primary antibody solution to mix it properly. Clean a glass slide properly with RNaseZap and let it dry for some time. Put the primary antibody solution on the glass slide. Then gently place the coverslip over it with a pair of forceps (cleaned with RNaseZap). Make sure that the side of the coverslip with cells faces the antibody mix. Keep it at room temperature inside a heavily humidified chamber for 60 min (see Note 5). 8. After 60 min remove the coverslip with forceps and a needle (both cleaned with RNaseZap), and keep it inside the 35 mm plastic bottom dish. Make sure the side of the coverslip with cells is facing upward. Give two washes with 1× PBS for 5 min each at room temperature (see Note 2). 9. Now make the secondary antibody mix using the same procedure by adding appropriate secondary antibody in 1:300 dilution in blocking solution. Clean a glass slide properly with RNaseZap and let it dry for some time. Put the secondary antibody solution on the glass slide. Then gently place the coverslip over it with forceps (cleaned with RNaseZap). Make sure that the side of the coverslip with cells faces the antibody mix. Keep it at room temperature inside a heavily humidified chamber and incubate for 60 min (see Note 5). 10. After 60 min, remove the coverslip with forceps and a needle (both cleaned with RNaseZap) and keep it inside the 35 mm plastic bottom dish. Make sure the side of the coverslip with cells is facing upward. Wash twice with 1× PBS for 5 min each at room temperature (see Note 2).

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11. Refix the cells with 4% PFA in 1× PBS for 15 min at room temperature. 12. Wash the cells twice with 1× PBS. 3.3

DNA FISH

1. Wash the cells once with 2× SSC. 2. Prepare 1.2 mL denaturation solution fresh. Make sure formamide is at room temperature before use. Add denaturation solution on the coverslip and keep it in 80 °C for 15 min. 3. In the meantime, make hybridization mixture for DNA FISH with 9 μL of Empire Genomics hybridization buffer and 1 μL appropriate DNA FISH probe in a microcentrifuge tube. Vortex it properly. Heat the hybridization mix at 80 °C for 3 min using a block heater. 4. After a 15-min incubation of the cells in denaturation solution, give it a brief wash with DNA FISH wash buffer. 5. Clean a glass slide properly with RNaseZap and let it dry for some time. Put the hybridization mixture on the glass slide. Then gently place the coverslip over it with forceps (cleaned with RNaseZap). Make sure that the side of the coverslip with cells faces the hybridization mix. Keep it at 37 °C for 24 h inside a heavily humidified chamber (see Notes 6 and 7). 6. The next day, remove the coverslip from the slide and put it in the 35 mm dish. Make sure that the side of the coverslip with cells is facing upward. Wash it twice with DNA FISH wash buffer for 10 min each at 37 °C (see Note 2).

3.4

RNA FISH

1. Wash the cells twice with RNA FISH wash buffer briefly. 2. Prepare 15 μL RNA FISH hybridization mixture with 13 μL Stellaris hybridization buffer, 1.5 μL 100% formamide, and 0.5 μL appropriate smFISH probe (e.g., TP53 or CCNA2). Vortex the hybridization mix. Clean a glass slide properly with RNaseZap, and let it dry for some time. Put the hybridization mix on the glass slide. Then gently place the coverslip over it with forceps (cleaned with RNaseZap). Make sure that the side of the coverslip with cells faces the hybridization mix. Keep it inside a heavily humidified chamber and incubate for 12–16 h at 37 °C (see Note 6). 3. The next day, remove the coverslip from the slide using forceps and a needle and put it in the 35 mm dish. 4. Wash the cells twice with RNA FISH wash buffer for 15 min each at 37 °C. In the second wash, add 1 μg/mL DAPI to the wash buffer (see Note 2). 5. Give the cells a brief wash with 2× SSC.

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6. Prepare a glass slide by washing it with RNaseZap and let it dry for some time. Then put 13 μL Vectashield on it. Take the coverslip with forceps and gently place it on the slide containing Vectashield. Make sure that the side of the coverslip with cells faces Vectashield. 7. Keep the slide for at least 1 h in 4 °C before imaging. 3.5 Imaging and Image Analysis

1. A 60×, 1.42 NA objective on a motorized widefield epifluorescence microscope (e.g., Olympus BX63 upright or IX83 inverted microscope) is used for imaging. A Retiga 6000 CCD camera is used without binning to resolve individual transcripts. 2. Filter cubes should be chosen such that there is no spectral bleed-through between the channels. Typically, 31 z-slices for HeLa cells and 23 z-slices for U2OS cells are routinely acquired with a constant step size of 300 nm. Typical exposure times used for CCNA2 are as follows: 250 ms for CCNA2 DNA FISH, 700 ms for CCNA2 RNA FISH, 1 s for CCNA2 protein, and 500 μs for DAPI. Typical exposure times for TP53 are as follows: 300 ms for TP53 DNA FISH, 1 s for TP53 RNA FISH, 2 s for TP53 protein, and 600 μs for DAPI. Hundreds of cells are analyzed in a typical experiment; details of analysis performed are available in reference [10]. 3. Immunofluorescence analysis: Segmentation is done using image processing toolbox of MATLAB using custom-written codes. 4. RNA FISH data analysis: RNA FISH spot counting is done starting from a MATLAB Code developed in a seminal study [15]. This code counts spots in 3D and not on Z-projections. More evolved versions of this tool may be found at https:// rajlab.seas.upenn.edu/resources too. 5. MATLAB codes and user-defined functions for analysis of nuclear architecture-dependent gene expression were customwritten [10] and are available at https://github.com/shuppar.

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Notes 1. Poly-D-Lysine (PDL) coating may be necessary for cells to adhere properly on the coverslip, depending on the cell type used. If cells adhere without PDL coating, then this step can be avoided. If PDL coating is done, then it needs to be washed thoroughly; otherwise, higher concentration of PDL can cause cytotoxicity.

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2. For the washing, fixation, and permeabilization steps, try to add more than 1 mL of the respective buffers so that the whole coverslip is submerged in the buffer. Drying out of cells in any steps should be avoided. 3. In some cases after fixation, the coverslip can be stored in 70% molecular biology-grade ethanol (made in nuclease-free water) at -20 °C for a few days. But caution should be exercised to ensure that the primary antibody still recognizes the protein against which it is directed and also that there is not dehydration induced change in nuclear architecture. 4. In our experience, the blocking step can be avoided if the primary antibody is highly specific. Recent reports have shown that blocking step is not critical for antibody staining [22]. But we use the blocking solution also for incubation with primary and secondary antibodies. 5. VRC is used just to make sure that RNases are inactivated in the solutions used in immunofluorescence. Adding VRC can be avoided generally, but if there is loss of smFISH signals, VRC may be used. 6. Always clean the forceps and needles with RNaseZap or RNase AWAY before use. Before placing the coverslip on the slide in the hybridization steps, there might be some remaining wash buffer present on the coverslip. Try to remove the wash buffer from one end slightly with a Kimwipe (do not spend much time in this step as the cells can dry out). Do not touch the cells while doing this. Touch the end of the coverslip with the glass slide first, then bring the end in contact with the hybridization mix and keep it around 30 seconds in the slanting position so that the hybridization mixture can mix with the remaining wash buffer present in the coverslip. Place the coverslip carefully on the slide and make sure there is no bubble. This procedure is necessary for even spread out of the DNA FISH/smFISH probes; else, all the cells might not get equal exposure to the probes. 7. The incubation time for DNA FISH hybridization can be increased if the signal is not observed after 24 h of hybridization. It can be more than 40 h depending on the probes [10]. For us the TP53 probes worked in 24 h, while CCNA2 probes took 40 h.

Acknowledgments This work was supported by intramural funds at TIFR Hyderabad from the Department of Atomic Energy, Government of India (Project Identification No. RTI 4007).

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References 1. Dekker J, Rippe K, Dekker M, Kleckner N (2002) Capturing chromosome conformation. Science 295(5558):1306–1311 2. Finn EH, Pegoraro G, Branda˜o HB, Valton A-L, Oomen ME, Dekker J, Mirny L, Misteli T (2019) Extensive heterogeneity and intrinsic variation in spatial genome organization. Cell 176:1502–1515.e10 3. Chambeyron S, Bickmore WA (2004) Chromatin decondensation and nuclear reorganization of the HoxB locus upon induction of transcription. Genes Dev 18:1119–1130 4. Clemson CM, McNeil JA, Willard HF, Lawrence JB (1996) XIST RNA paints the inactive X chromosome at interphase: evidence for a novel RNA involved in nuclear/chromosome structure. J Cell Biol 132:259–275 5. Clemson CM, Hutchinson JN, Sara SA, Ensminger AW, Fox AH, Chess A, Lawrence JB (2009) An architectural role for a nuclear noncoding RNA: NEAT1 RNA is essential for the structure of paraspeckles. Mol Cell 33:717– 726 6. Morey C, Kress C, Bickmore WA (2009) Lack of bystander activation shows that localization exterior to chromosome territories is not sufficient to upregulate gene expression. Genome Res 19:1184–1194 7. Zhang L-F, Huynh KD, Lee JT (2007) Perinucleolar targeting of the inactive X during S phase: evidence for a role in the maintenance of silencing. Cell 129:693–706 8. Okamoto I (2018) Combined immunofluorescence, RNA FISH, DNA FISH in preimplantation mouse embryos. Methods MoBiol 1861: 149–159 9. Chaumeil J, Augui S, Chow JC, Heard E (2008) Combined immunofluorescence, RNA fluorescent in situ hybridization, and DNA fluorescent in situ hybridization to study chromatin changes, transcriptional activity, nuclear organization, and X-chromosome inactivation. Methods Mol Biol 463:297–308 10. Dhuppar S, Mazumder A (2020) Investigating cell cycle-dependent gene expression in the context of nuclear architecture at single-allele resolution. J Cell Sci 133(12):jcs246330 11. Zenklusen D, Larson DR, Singer RH (2008) Single-RNA counting reveals alternative modes of gene expression in yeast. Nat Struct Mol Biol 15:1263–1271

12. Dhuppar S, Mazumder A (2018) Measuring cell cycle-dependent DNA damage responses and p53 regulation on a cell-by-cell basis from image analysis. Cell Cycle 17:1358–1371 13. Pasnuri N, Jaiswal M, Ray K, Mazumder A (2023) Buffered EGFR signaling regulated by spitzto-argos expression ratio is a critical factor for patterning the Drosophila eye. PLoS Genet 19(2):e1010622 14. Pasnuri N, Khuntia P, Mazumder A (2018) Single transcript imaging to assay gene expression in wholemount Drosophila melanogaster tissues. Mech Dev 153:10–16 15. Raj A, van den Bogaard P, Rifkin SA, van Oudenaarden A, Tyagi S (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5:877–879 16. Raj A, Peskin CS, Tranchina D, Vargas DY, Tyagi S (2006) Stochastic mRNA synthesis in mammalian cells. PLoS Biol 4:1707–1719 17. Jiang J, Jing Y, Cost GJ, Chiang J-C, Kolpa HJ, Cotton AM, Carone DM, Carone BR, Shivak DA, Guschin DY et al (2013) Translating dosage compensation to trisomy 21. Nature 500: 296–300 18. Century TJ, Fenichel IR, Horowitz SB (1970) The concentrations of water, sodium and potassium in the nucleus and cytoplasm of amphibian oocytes. J Cell Sci 7:5–13 19. Bienko M, Crosetto N, Teytelman L, Klemm S, Itzkovitz S, Van Oudenaarden A (2013) A versatile genome-scale PCR-based pipeline for high definition DNA FISH. Nat Methods 10: 122–124 20. Bolland DJ, King MR, Reik W, Corcoran AE, Krueger C (2013) Robust 3D DNA FISH using directly labeled probes. J Vis Exp 78: e50587 21. Morey C, Da Silva NR, Perry P, Bickmore WA, Kanno M, Taniguchi M, Vidal M, Alkema M, Berns A, Koseki H (2007) Nuclear reorganisation and chromatin decondensation are conserved, but distinct, mechanisms linked to Hox gene activation. Development 134:909– 919 22. Buchwalow I, Samoilova V, Boecker W, Tiemann M (2011) Non-specific binding of antibodies in immunohistochemistry: fallacies and facts. Sci Rep 1:28

Chapter 15 Determining the Compaction State of Genes Using DNA FISH Masako Narita, Ioana Olan, and Masashi Narita Abstract DNA fluorescence in situ hybridization (FISH) enables the visualization of chromatin architecture and the interactions between genomic loci at a single-cell level, complementary to genome-wide methods such as Hi-C. DNA FISH uses fluorescent-labeled DNA probes targeted to the loci of interest, allowing for the analysis of their spatial positioning and proximity with microscopy. Here, we describe an optimized experimental procedure for DNA FISH, from probe design and sample preparation through imaging and image quantification. This protocol can be readily applied to querying the spatial positioning of genomic loci of interest. Key words DNA fluorescence in situ hybridization (FISH), Senescence, Heterochromatic foci, Chromatin architecture, Spatial positioning, Image quantification

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Introduction The three-dimensional organization of chromatin within the nucleus plays a crucial role in gene regulation and cellular states [1, 2]. High-order chromatin structure has traditionally been studied using microscopic imaging techniques such as fluorescence in situ hybridization (FISH) and genomic methods, including chromosome conformation capture (3C) and its extensions like Hi-C. Within the chromatin architecture field, there is a long-standing debate on how to reconcile the physical distance between genomic loci observed from microscopy experiments, in a small number of cells, with the contact frequencies and relative positioning of genomic loci inferred from genomic experiments performed on millions of cells (e.g., Hi-C) [3]. Historically, researchers focused on capturing representative images, but more recently, the focus has shifted toward simultaneously capturing representative images as well as quantification of a reasonably sized sample of cells. While DNA-FISH is widely used for assessing the physical position of genomic loci within the cell nucleus [4], many modifications exist to the FISH protocol that require optimization at various steps.

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Moreover, the comparison between different biological conditions is often of interest. The reconciliation of the relative positioning of different loci with observations derived from chromatin capture assays (e.g., Hi-C) requires careful interpretation [5]. In this chapter, we will outline the steps required to successfully design probes for regions of interest, perform DNA-FISH, capture images, and quantify the relative positioning of multiple genomic locations. This protocol was optimized for cultured human diploid fibroblasts, both normal and senescence (a state of stable cell cycle arrest with distinct cellular and nuclear morphology) [6, 7]. While further optimization might be required, the principle can be applied to other cell types. As FISH probes, this protocol uses bacterial artificial clones (BACs), which carry large fragments of, for example, the human genome. The size typically ranges from 100 to 300 kilobases (kb) with a median length of 157 kb.

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Materials

2.1 Cell Culture on Coverslips

1. 12-well cell-culture treated multi-dish. 2. Round glass coverslips: 16 mm in diameter, No. 1.5 thickness. Cleaned with 2M NaOH and stored in 70% ethanol (see Note 1). 3. 0.1% gelatin solution.

2.2 Fixation and Permeabilization

1. Phosphate buffered saline (PBS, pH 7.4). 2. Fixation buffer: 4% paraformaldehyde (PFA) EM grade in PBS pH 7.4. Aliquot and store at -20 °C. 3. 1M hydrochloric acid. 4. Triton X-100 solution: 10% in H2O. 5. 20× SSC (saline sodium citrate) buffer: 3M NaCl, 0.3M sodium citrate, pH 7.0. 6. Permeabilization solution: freshly made ice-cold 0.1M HCl, 0.7% Triton X-100 or 70% ethanol in distilled water. 7. 2× SSC.

2.3 Probe Preparation

1. Bacterial artificial chromosome (BAC) clones as fluorescently labeled FISH probes (Empire Genomics (EG)) (see Subheading 3.1 on probe design). 2. Hybridization buffer (Hyb B) supplied with the probes (EG). 3. 0.2 mL PCR tubes. 4. Thermal cycler. 5. Foil, to cover and keep reagents/samples from direct light.

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Dehydration

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1. Formamide, stored at 4 °C after opening to keep deionized. 2. Wash buffer: 2× SSC, 10% formamide. Prepare fresh. 3. 70% ethanol in distilled water. Prepare fresh. 4. 85% ethanol in distilled water. Prepare fresh. 5. 100% ethanol, molecular grade.

2.5 Probe Codenaturation and In Situ Hybridization

1. Rubber cement. 2. Heat block (73 °C). 3. Slide glasses: SuperFrost Plus Adhesion slides, 25 × 75 × 1 mm. 4. Humidified hybridization chamber (see Note 2).

2.6 Wash, Counterstain, and Mounting

1. 10% Igepal solution (see Note 3). 2. Wash solution 1 (WS1): 0.3% Igepal or NP-40/0.4× SSC at 73 °C. Prepare fresh. 3. Wash solution 2 (WS2): 0.1% Igepal or NP-40/2× SSC Prepare fresh. 4. DAPI (4′,6-diamidino-2-phenylindole, dihydrochloride) stock solution: 1 mg/mL in H2O. Store frozen and protected from light. 5. Counterstain DAPI solution: 0.2–0.125 ug/mL DAPI in 2× SSC. Prepare fresh. 6. Antifade mounting medium. 7. Coverslip sealant or nail polish.

2.7

Imaging

1. Wide-field or confocal fluorescence microscope. 2. 63× or 100× oil immersion objectives. 3. Appropriate fluorescent filters for each probe dye. 4. Scikit-image software.

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Methods There are numerous methods and variations for probe preparation and DNA FISH [4]. The following protocol is optimized for human diploid fibroblasts using pre-labeled ready-to-use BAC probes, but the same BAC clones can also be labeled using standard methods, such as nick translation using fluorescent nucleotides [4].

3.1

Probe Design

Selecting regions of interest: Choosing genomic locations of interest is highly context-dependent and can include enhancerpromoter pairs, loop anchors, or regions attached and detached from their local chromatin environment in a condition-dependent manner. For instance, based on Hi-C experiments, the genome is

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organized in insulated topological-associating domains (TAD), which can be detected using BAC-based probes. As an example, in our condition of interest (cellular senescence), some genes lose interactions with the rest of the TAD they belong to and become “detached” (Fig. 1a). This detachment can be tested and visualized by choosing FISH probes corresponding to the gene and a nearby location representing the remaining genomic region within the same TAD (Fig. 1b). Potential probe candidates can be found by cross-checking the coordinates of BAC clones with the regions of interest (gene bodies/promoters). Candidates are then selected based on good overlap, as well as membership of the BAC clone to a set of validated clones as suitable for FISH: 1. Go to UCSC Genome Browser. https://genome.ucsc.edu/ 2. Go to “Genome” and select, e.g., “Human GRCh37/hg19” (Fig. 2a). To identify genomic regions of interest, enter, e.g., gene names in the search box (Fig. 2b). 3. Expand “Mapping and Sequencing.” At the “BAC End Pairs” pull-down menu, choose “squash” or “pack” and “FISH clones” pull-down menu to “pack” or “full.” Then “refresh” (Fig. 2c). 4. Select two or more BAC clones with ≥50% overlap with each target region. Prioritize clones validated for FISH (Fig. 2d). 5. Click the chosen BAC clones to obtain more information (see Note 4). 6. The hg19 coordinates of all BAC clones can be downloaded from UCSC: https://shorturl.at/bev29 7. From this link, you can find clones mapping to regions of interest. https://bacpacresources.org/mapped-clones. htm#find (see FISH Mapped Clones V1.3 Download). 8. Some BAC library clones (RPCI-11 and others) can be ordered from BACPAC Genomics, RIKEN, or possibly from other companies. We typically order custom FISH probes from Empire Genomics (EG) (see Note 5). Their Custom FISH probes can contain up to four/five fluorescent dyes (see Note 6). 3.2

DNA FISH

Oncogenic HRASG12V-induced senescence (RIS) can be achieved by the overexpression of HRASG12V in IMR90 cells, human female diploid fetal lung fibroblasts. HRASG12V can be introduced to the cells through retroviral gene transfer. This procedure was detailed previously [8]. Senescence is typically established around 7–9 days after infection of the HRASG12V-expressing retrovirus [8].

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Fig. 1 Assessing dynamic alterations to chromatin structure during senescence using Hi-C and DNA-FISH. (a) Example of probe design at the NRG1 gene locus: Hi-C interaction maps of proliferating (top) and oncogenic RAS-induced senescent (RIS) (bottom) IMR90 human fibroblasts at 40 kb resolution, showing the detachment of the NRG1 gene from its associated TAD (topologically associated domain), marked by global loss of interactions between the gene body and the rest of the TAD. Hi-C data were reanalyzed from the previous study [6]. The genomic coordinate is based on the GRCh37/hg19 human reference genome assembly. Probe 1 (BAC clone RP11-57I3) targeted the gene body of NRG1, and Probe 2 (BAC clone RP11-451O18) targeted to a nearby region contained within the same TAD defined in the proliferating condition. Relative genomic locations of the TAD (proliferating cells), gene body, and probes are magnified below. (b) Representative DNA-FISH images, corresponding to the same condition as the Hi-C map (top, proliferating; bottom RIS), using Probe 1 and Probe 2, as marked in A. In the RIS condition, green fluorescence signals (Probe 1: NRG1) show an extended pattern, while magenta signals (Probe 2: TAD border) remain punctate

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Fig. 2 Screenshots of probe design steps using UCSC genome browser. (a) Select “Genome” (arrow) and choose “Human GRCh37/hg19” (ellipse) as the reference genome. (b) In the search box, enter the name of the gene of interest (e.g., NRG1) (arrow). (c) Under the “Mapping and Sequencing” section (arrow), expand the options. Unhide “BAC End Pairs” and “FISH Clones” (ellipses) from the pull-down menus. (d) Select BAC clones (arrow) that exhibit a minimum of 50% overlap with each target region (rounded rectangle)

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1. Place sterilized coverslips in a 12-well plate (one per well). 2. Aspirate any remaining ethanol and leave to dry. 3. Coat the coverslips with a few drops of 0.1% gelatin solution at room temperature for a few minutes (see Note 7). 4. Aspirate the Gelatin and seed around 250,000 cells (for IMR90) onto the coverslips. Incubate the cultures at 37 °C; 5% CO2, and preferred % of O2, overnight.

3.2.2 Fixation and Permeabilization (Day 2)

It is recommended to process fresh cells without stopping from here to mounting. 1. Aspirate media, wash with PBS once. Remove PBS by decantation. 2. Fix cells with 500 μL of fixation buffer, tilting gently (e.g., on a motorized platform) for 5 min at room temperature. 3. Wash with PBS ×3 (total 5–10 min). 4. Permeabilize the cells in ice-cold 0.1M HCl, 0.7% Triton X-100 for 10 min on ice (see Note 8). 5. Wash with 2× SSC twice, for 5 min each at room temperature.

3.2.3

Probe Preparation

1. Start by thawing the Empire Genomics probes and its supplied hybridization buffer (Hyb B) at room temperature for 15 min. Vortex and spin down briefly. 2. Mix the probes and Hyb B well in 0.2 mL PCR tube(s) by pipetting vigorously and briefly spinning down (see Note 9 for the volume of each). As a guide to the amount of probe mixture needed, allow 4.5–5 μL total per ø16 mm (200 mm2), which is equivalent to 10 μL mixture per 22 × 22 mm (484 mm2) coverslip. 3. Start to denature the probe mixture by heating for 5 min at 73 ° C while the cells are dehydrating (next section).

3.2.4

Dehydration

1. Wash with wash buffer and incubate at room temperature for 2–5 min. Aspirate wash buffer. 2. Apply 1 mL 70% ethanol. Leave for 2 min. Remove the ethanol by decantation. 3. Apply 1 mL 85% ethanol. Leave for 2 min. Remove the ethanol by decantation. 4. Apply 1 mL 100% ethanol. Leave for 2 min, no decantation. 5. Air dry the coverslips on a paper towel until the hybridization step.

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3.2.5 Probe Codenaturation and In Situ Hybridization

1. Immediately after denaturation of the probe mixture, apply the 4.5–5 μL of probe mixture onto a slide glass. Ensure air bubbles are removed. 2. Cover the DNA probe mixture with the cell coverslip, with the cells face down. 3. Seal all coverslip edges with rubber cement to prevent the probe from drying out. 4. Place the slide glass with probe mixture and coverslip on a 73 °C heat block to co-denature the probe and the genomic DNA. Incubate for 2 min. Use a plastic lid for heat retention. 5. Incubate the slides in a humidified hybridization chamber at 37 °C (see Note 2) overnight in the dark (at least 16–20 h).

3.2.6 Wash, Counterstain, and Mounting (Day 3)

1. Prewarm WS1 at 73 °C. 2. Carefully peel off the rubber cement with forceps and fingers. Slide the coverslip toward the proximal edge of the slide glass and remove the coverslip slowly from the edge. 3. Transfer the coverslips to a 12-well plate with 2× SSC. 4. Post-hybridization wash step 1: Wash in 1 mL of WS1 for 2–3 min at 73 °C. Agitate the coverslips for the first 10–15 s. Leave the plate on a 73 °C heat block. 5. Post-hybridization wash step 2: Wash in 1 mL of WS2 for 2 min at room temperature. Agitate the coverslips for the first 10–15 s. 6. Wash in 1 mL 2× SSC for 2 min. 7. Counterstain the coverslips by incubation in 500 μL of DAPI solution for 5 min. 8. Wash in 1 mL 2× SSC for 2 min. 9. Gently replace buffer with H2O (to remove salt from the buffer). 10. Pick up the coverslip with forceps and aspirate excess H2O (take care not to let the coverslip dry). 11. Mount the coverslips using 5 μL mounting solution on a glass slide. 12. Aspirate the excess mounting solution and seal the edge of the coverslip with CoverGrip or nail polish. Let it completely dry at room temperature. 13. Store the specimen at -20 °C.

3.3

Imaging

Capture images using a 63× or a 100× objective using a confocal fluorescence microscope with the appropriate detectors. Consider z-stacks if necessary. Compare spatial relationships between conditions.

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Quantification

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After confocal imaging, the 2D quantification of the FISH signal and the distance between two probes consists of using scikit-image [9] to apply filtering to the image to improve the signal-to-noise ratio and then image segmentation. In our case, each image consisted of three channels corresponding to the DAPI signal (ch00), FISH probe 1 (ch01), and FISH probe 2 (ch02) (see Note 10). 1. Test and choose an appropriate image filtering algorithm, e.g., the Li filter for the DAPI signal and the Yen filter for the FISH signal. 2. Apply Clear Border segmentation to the DAPI signal (ch00) to obtain objects corresponding to each cell nucleus. 3. Apply Clear Border segmentation or blob detection using the Laplacian of the Gaussian to the FISH signal to delimit the blobs corresponding to each probe (ch01 and ch02). 4. Calculate the distances between each ch01 blob and ch02 blob residing within the same cell nucleus and pair the closest ones, as the two probes correspond to genomic locations which are very close to each other on the linear genome. 5. Use this “nearest” distance as the estimate of the distance between the two regions in 3D space. 6. To compare the relative positioning of the two FISH probes across conditions, calculate the distance between the probes in at least 200 cells in each condition and apply statistical testing (Student’s t-test or Wilcox). In most cases, there were two pairs of probes in each nucleus corresponding to the two copies of the chromosome.

4

Notes 1. It is imperative that the coverslips used in fluorescence microscopy procedures be very clean. There may be a thin film coating them that will not allow cultured cells to adhere well. Therefore, it is recommended to wash the coverslips with acid or base solutions to rid them of this coating. We usually drop our coverslips into 2M NaOH individually in succession. Allow the coverslips to sit for 2 h with occasional swirling. Then rinse extensively in dH2O until the pH of the wash water is back to ~pH 7. Store the coverslips in a covered container submerged in 70% ethanol. 2. Prepare a humidified hybridization chamber by filling a smaller plastic container (e.g., 10 × 10 cm), and placing it with watersoaked tissues within a plastic storage container with snapclosed lids (e.g., 18 × 13 cm). Place the slides in a smaller plastic container. Snap-close the outer lid to prevent evaporation.

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3. Make the 10% IGEPAL® CA-630 solution in distilled water by stirring overnight. 4. Clones referred to as RP11 would be ordered from library RPCI-11; clones referred to as CH230- would be ordered from library CHORI-230; clones referred to as RP98- or BACR would be ordered from library RPCI-98. 5. Empire Genomics accesses the RPCI-11 human BAC library and RPCI-23 mouse BAC libraries. 6. These fluorophores are FITC (Ex/Em = 491/515 nm), Rhodamine 6G (525/551 nm), 5-TAMRA (548/573 nm), 5-ROX (580/599 nm), and Aqua (418/467 nm). But Aqua is relatively weak and cannot be counter-stained with DAPI. 7. Gelatin subbing enhances the adherence of cells to coverslips. 8. This method preserves heterochromatic foci (e.g., senescenceassociated heterochromatic foci (SAHFs) [10]), condensed chromatin that appears as dense spots in the nucleus when the cells are stained for DNA (e.g., DAPI) and viewed microscopically. Alternatively, immerse the cells in ice-cold 70% ethanol for at least 1 h, tilting gently in a cold room. 9. It is possible to stain up to four EG FISH probes at the same time to visualize a particular structure of chromatin. Below is the mixture ratio with the supplied hybridization buffer (Hyb B) for each ø16 mm coverslip. It is not recommended to use lower volumes. For 1 Probe: 1 μL probe with 4 μL Hyb B For 2 Probes: 1 μL probe each with 3 μL Hyb B For 4 Probes: 0.5 μL probe each with 3 μL Hyb B 10. Scikit-image is an image processing program in Python [9]. The recommendations provided are intended as general guidance; users should test different filters and segmentation algorithms to find the optimal combination for their specific images. 11. Here are some tips for preparing samples to compare multiple conditions using DNA FISH: Culture the cells for each condition on coverslips in parallel. Keep seeding density and culture conditions consistent. Acquire images using identical microscope settings: objective, exposure time, z-stack parameters, etc. Capture sufficient cells to allow statistical comparisons. Use automated image analysis tools whenever possible. Apply appropriate statistical tests to compare probe distances or radial positions between conditions and determine significance. Increase biological replicates to ensure robustness.

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Acknowledgments The authors thank Andrew Young for critical reading. This work is supported by a Cancer Research UK Cambridge Institute core grant (no. C9545/A29580) to the Narita laboratory. Masashi N is also supported by BBSRC (BB/S013466/1 and BB/T013486/ 1) and Diabetes UK via BIRAX and the British Council (65BX18MNIB). IO is also supported by a Cancer Research UK Pioneer Award (C63389/A30462). References 1. Olan I, Handa T, Narita M (2023) Beyond SAHF: an integrative view of chromatin compartmentalization during senescence. Curr Opin Cell Biol 83:102206 2. Olan I, Narita M (2022) Senescence: an identity crisis originating from deep within the nucleus. Annu Rev Cell Dev Biol 38:219–239 3. Fudenberg G, Imakaev M (2017) FISH-ing for captured contacts: towards reconciling FISH and 3C. Nat Methods 14:673–678 4. Chaumeil J, Augui S, Chow JC et al (2008) The nucleus, volume 1: nuclei and subnuclear components. Methods Mol Biol 463:297–308 5. Dekker J (2016) Mapping the 3D genome: aiming for consilience. Nat Rev Mol Cell Biol 17:741–742 6. Olan I, Parry AJ, Schoenfelder S et al (2020) cohesin Transcription-dependent

repositioning rewires chromatin loops in cellular senescence. Nat Commun 11:6049 7. Tomimatsu K, Bihary D, Olan I et al (2022) Locus-specific induction of gene expression from heterochromatin loci during cellular senescence. Nat Aging 2:31–45 8. Narita M, Narita M (2016) Oncogene-induced senescence, Methods and Protocols, Presented at the November 4 9. van der Walt S, Scho¨nberger JL, NunezIglesias J et al (2014) Scikit-image: image processing in python. PeerJ 2:e453 ˜ ez S, Heard E et al (2003) 10. Narita M, Nun Rb-mediated heterochromatin formation and silencing of E2F target genes during cellular senescence. Cell 113:703–716

Chapter 16 Hi-M: A Multiplex Oligopaint FISH Method to Capture Chromatin Conformations In Situ and Accompanying Open-Source Acquisition Software Jean-Bernard Fiche, Marie Schaeffer, Christophe Houbron, Christel Elkhoury Youhanna, Olivier Messina, Franziska Barho, and Marcelo Nollmann Abstract The simultaneous observation of three-dimensional (3D) chromatin structure and transcription in single cells is critical to understand how DNA is organized inside cells and how this organization influences or is affected by other processes, such as transcription. We have recently introduced an innovative technology known as Hi-M, which enables the sequential tagging, 3D visualization, and precise localization of multiple genomic DNA regions alongside RNA expression within individual cells. In this chapter, we present a comprehensive guide outlining the creation of probes, as well as sample preparation and labeling. Finally, we provide a step-by-step guide to conduct a complete Hi-M acquisition using our open-source software package, Qudi-HiM, which controls the robotic microscope handling the entire acquisition procedure. Key words Fluorescence microscopy, Fluorescent in situ hybridization, 3D genome architecture, Transcription

1

Introduction Genomes are folded in a hierarchical organization that contributes to regulating transcription and other processes [1, 2]. In the last decade, chromosome conformation capture (3C and derivatives) has revolutionized our understanding of chromatin architecture and has provided important insight into the molecular mechanisms involved [3, 4]. Microscopy-based strategies based on fluorescence in situ hybridization (FISH) have more recently been developed to image chromatin organization in single cells, unveiling a large degree of heterogeneity [5, 6]. These approaches, however, could

Co-first authors: Jean-Bernard Fiche, Marie Schaeffer, Christophe Houbron. Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Outline of Hi-M experiment. Schematic description of the main steps in the Hi-M protocol. Samples are immobilized in a microfluidics chamber (gray circle). A pump is used to deliver the FISH probes and buffers into the chamber. Barcode solutions (tubes with colored caps) are sequentially injected using a needle mounted on a XYZ translation stage. Images are acquired using a widefield or a confocal microscope (microscope icon). DNA-FISH spots (dots of different colors on the chromosome) are imaged and sequentially bleached to enable 3D chromosome tracing in single cells

only detect a small number of genomic locations at once (typically 3–4), limiting their ability to dissect structure-function relationships or to detect levels of chromatin organization involving multiple loci. We and others have recently overcome this limitation by combining DNA-FISH with robotized microfluidics devices. This new technology that we termed Hi-M (high-throughput, highresolution, high-coverage microscopy-based technology; see below) employs high-throughput synthesis of short oligonucleotide (oligo) probes combined with RNA labeling and multiple rounds of hybridization in a sequential imaging scheme to enable the localization of tens of different genomic loci alongside the transcriptional state of the cell [7] (Fig. 1). Similar approaches have also been employed in concurrent work [8–11] and build upon pioneering work from Wang and colleagues [12]. More recently, combinatorial encoding was used to acquire highly multiplexed datasets [13, 14]. Hi-M is built upon two fundamental concepts: the recent advancement in FISH probe design known as Oligopaints, which utilizes high-throughput microarray oligo synthesis [15, 16], and the groundbreaking development of multiplexing strategies for imaging numerous RNA species [17, 18]. In the Hi-M technique, a microarray library containing thousands of bioinformatically designed and commercially synthesized oligonucleotides (referred to as an oligopool) is employed to target multiple genomic locations. These genomic loci, typically spanning 3–25 kb, are labeled using unique sets of ~40–250 tiled oligos, forming what is termed “barcodes.” Each oligo within a barcode comprises three segments: a genomic homology region, a barcode-specific readout sequence, and a priming region for PCR amplification (Fig. 2a). The labeling process encompasses four steps. Initially, the oligopool library undergoes enzymatic amplification and purification (Fig. 2b–e).

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Fig. 2 Design and amplification of Oligopaints. (a) Upper panel, schematic structure of oligos: (1) a forward and reverse 20-nt universal priming region in blue-green, (2) a 32-nt readout sequence in blue, (3) a 42-nt genome homology variable region in black. Lower panel, schematic description of the steps to design an oligopool. (b) Upper panel, oligopool amplification scheme. Blue-green represents the universal priming region common to all barcodes whereas blue, amber and burgundy represent barcode-specific readout sequences. Lower panel, schematic description of main steps involved. .(c) Example of an agarose gel electrophoresis result for the small scale limited-cycle PCR step. “L” is the DNA ladder or molecular weight size marker. A band of the expected size (166-nt in this case) is observed between the 100- and 200-nt bands of the ladder. In cycles 13 and 14 a second, non-specific band of ~300-nt begins to appear. Therefore, 11 amplification cycles (at PCR cycle 12) were chosen for this specific amplification reaction. (d). Example of an agarose gel electrophoresis result for the large scale limited-cycle PCR step. “L” is the ladder as in panel Lane 1 and 2 correspond to a PCR performed without or with a template, respectively. dNTPs are observed at the bottom. Lane 3 corresponds to column-purified PCR products. (e) Example of an Urea PAGE result. “L” is the low range ssRNA ladder. Bands from lanes 1–4 appear close to the height of the 150-nt band from the ladder. Lane 1 corresponds to 200 ng of emulsion PCR break, lane 2 to 200 ng of RNA product from in vitro transcription (note the higher size due to the presence of the T7-promoter region), lanes 3 and 4 to 200 ng of ssDNA before and after precipitation, respectively. (Figure panels were adapted from Ref. [19])

Subsequently, samples are harvested and fixed. Finally, the oligopool is hybridized to the genomic DNA. Following labeling, the samples are affixed to a microfluidic chamber connected to a microfluidics pump system and inserted into an automated widefield fluorescence microscope. In the initial stage, DAPI and RNA signals are captured in various fields of view (FOVs) (also referred to as regions of interest (ROIs) in the acquisition software Qudi-HiM described below) (Fig. 3). The subsequent imaging of each barcode involves: (1) labeling the sample with fluorescently labeled readout

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Fig. 3 Pipeline for Hi-M experiment. .(a) A primary oligopool library (blue) is hybridized in the bench to genomic DNA prior to mounting to the fluidics device. A readout oligo (red), bearing a fluorophore (red star), specifically binds to the primary oligopool library. (b) 3D, multi-color images are taken for a user-specified number of FOVs. (c) The fluorophore on the readout oligo is chemically cleaved or alternatively, photobleached. (Figure adapted from Ref. [22]).

oligonucleotides specific to the barcode; (2) washing away unbound readout oligos; (3) capturing 3D, two-color images of all FOVs; and (4) photobleaching. This sequence is iterated for each barcode (Fig. 3). Throughout the sequential rounds, an additional fluorescent oligo with a distinct spectrum serves as the fiducial barcode to correct for any drift during post-acquisition analysis. Since a typical Hi-M experiment requires several days of automated acquisition, we developed Qudi-HiM, an open-source Python software suite that allows unsupervised and robust data acquisition and handling. Qudi-HiM controls and coordinates all hardware components needed for fluid handling (e.g., pumps, valves, etc.) and fluorescence 3D imaging (e.g., camera, piezo stage, acousto-optic tunable filter, etc.). Therefore, human intervention is only required to define the acquisition parameters (e.g., number of barcodes to inject, injection routines for barcode labeling and washing, laser powers, FOV positions, etc.), and QudiHiM automatically performs the Hi-M experiment for efficient and reproducible data acquisition.

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In this chapter, we provide a complete list of materials and stepby-step protocols for designing and amplifying Hi-M libraries using Oligopaints. We explain the procedures for sample preparation and DNA-FISH labeling in detail and provide examples of expected outcomes. Additionally, we outline the setup of a Hi-M experiment using our homemade widefield microscope, covering equipment and setup instructions. Lastly, we introduce Qudi-HiM and describe how to use it to acquire a full Hi-M dataset.

2 2.1

Materials Reagents

1. 20X saline-sodium citrate buffer (SSC): 3M NaCl, 0.3M sodium citrate. 2. 30% (w/w) hydrogen peroxide solution. 3. 16% formaldehyde solution (w/v). 4. 4′,6-Diamidine-2′-phenylindole dihydrochloride (DAPI). 5. Acetone. 6. Agarose standard DNA grade. 7. Alexa Fluor 488 Tyramide Reagent. 8. Atto-550 readout probes used for the fiducials. 9. Alexa-647 readout probes. 10. Ammonium persulfate (APS). 11. Ammonium acetate 5M. 12. Anti-digoxigenin-POD, AB_514500).

Fab

fragments

(Roche,

RRID:

13. BSA. 14. CHAPS. 15. Cetyl PEG/PPG-10/1 dimethicone. 16. Clorox ultra germicidal liquid bleach. 17. D(+) Glucose anhydrous. 18. DNA Clean & Concentrator kit: 100 μg capacity (Zymo). 19. DNA Clean & Concentrator kit: 25 μg capacity (Zymo). 20. Dextran sulfate. 21. Diethyl ether. 22. Dulbecco’s phosphate-buffered saline (PBS). 23. Dry fine yeast. 24. Ethyl acetate. 25. GeneRuler 100 bp DNA Ladder. 26. Glycogen 5 mg/mL.

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27. Glucose oxidase. 28. Heparin. 29. HiScribe T7 High Yield RNA Synthesis Kit (New England Biolabs). 30. KAPA Taq Kit with dNTPs (CliniSciences). 31. Low Range ssRNA Ladder (New England Biolabs). 32. Maxima H Minus Reverse transcriptase kit (Fisher Scientific). 33. Methanol. 34. Mineral oil. 35. Oligo Clean & Concentrator kit (Zymo). 36. Poly-L-lysine solution. 37. RNase A. 38. RNA loading dye 2X (New England Biolabs). 39. RNA Ribonuclease Inhibitors (Promega). 40. SYBR Gold nucleic acid gel stain (Fisher Scientific). 41. SYBR Safe nucleic acid gel stain (Invitrogen). 42. Sodium chloride (99,5%). 43. Sodium dihydrogen phosphate, dihydrate. 44. TAE Buffer 50X (Tris-acetate-EDTA). 45. TCEP (tris(2-carboxyethyl)phosphine). 46. TEMED. 47. Tris Base, BM grade. 48. Triton X-100. 49. Tween-20. 50. dNTP Set 100 mM Solution (Fisher Scientific). 51. Deionized formamide. 52. Dry fine yeast. 2.2

Equipment List

1. PCR Machine. 2. Positive displacement micropipette M250. 3. 1.8 mL plastic cryotube (Thermo Fisher Scientific). 4. Magnetic stirring bar. 5. Magnetic stirrer (10 mm) 6. NanoDrop spectrophotometer (Thermo Fisher Scientific, model no. ND-1000UV/Vis). 7. Vortex, standard mini vortex. 8. Falcon 15-mL conical centrifuge tubes. 9. Falcon 50-mL conical centrifuge tubes.

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10. Tabletop centrifuge. 11. Syringe 30 mL. 12. Syringe 20 mL. 13. Embryo collection cage (8.75 cm × 14.8 cm; Flystuff.com). 14. Nylon Filter. 15. Water bath Grant Instruments JBN5. 16. Thermomixer-AccuTherm (Labnet).

Microtube

Shaking Incubator

17. Gas burner. 18. Disposable scalpel. 19. Plastic petri dishes. 20. Disposable glass Pasteur pipette. 21. Glass vial for embryo collection (DWK Life Sciences). 22. Rotating wheel. 23. Wide-field epifluorescence microscope (see the wide-field epifluorescence microscope section). 24. Laser protective google (Thorlabs). 25. Microscope coverslips (Bioptechs Inc.). 26. Microfluidics FCS2 chamber, no heat and low dead volume (Bioptechs Inc.). 27. PEEK tubing 1/16″ × 0.75 mm green (CIL). 28. TEFLON tubing 1/16″ × 1 mm (CIL). 29. MFCS-EZ negative pressure pump (Fluigent). 30. Micro-perfusion peristaltic pump (Instech). 31. Flow unit L (Fluigent). 32. HVXM8-5 & HVMXM2-5 injection valves and controllers (Hamilto). 33. Huygens deconvolution software (Scientific Volume Imaging, https://svi.nl/HuygensSoftware). 34. Server running Linux with 256 threads, a GeForce GTX 2080ti GPU card, and 512GB of RAM. 2.2.1

Reagent Setup

1. PCR oil phase: 95.95% mineral oil, 4% ABIL EM-90, and 0.05% Triton X-100 (v/v/v). If you have access to a positive displacement pipette, you can conveniently pipette 2 mL of ABIL EM90, 65 μL of Triton X-100, and 47.975 mL of mineral oil into a 50-mL Falcon tube. When adding the mineral oil, do so in two steps, vortexing in between. In case a positive displacement pipette is not available, you can accurately measure the volume by weight. To prepare 50 mL of the PCR oil phase, weigh 20.3 g of mineral oil (approximately

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24 mL) directly into a 50 mL Falcon tube. Then, add 2 mL of ABIL EM90 and 65 μL of Triton X-100, vortex the mixture thoroughly, and allow it to sit for 5 min. Next, add 20 g of mineral oil and homogenize the mixture by gently inverting the tube. Finally, create 20 mL aliquots of the PCR oil phase and store them at 4 °C indefinitely. 2. Water-saturated diethyl ether: 3 mL of diethyl ether, 3 mL of ddH2O. Vortex for 30 s. Allow the mixture to settle and use the organic upper phase. Prepare freshly. 3. Water-saturated ethyl acetate: 2 mL of ethyl acetate, 2 mL of ddH2O. Vortex for 30 s. Allow the mixture to settle and use the organic upper phase. Prepare freshly. 4. 10% Tween 20 (v/v) solution: 50 μL of Tween 20, 450 μL of ddH2O. Vortex until the solution becomes homogeneous. Store at 4 °C for up to 2 weeks. 5. PBS–Tween 20 (PBT) solution: 49.5 mL of PBS, 500 μL of 10% Tween 20 (v/v). 6. PBS-Triton X-100 (PBS-Tr) solution: 50 μL of Triton X-100, 10 mL of PBS. Vortex until the solution becomes homogeneous. 7. 4% (w/v) formaldehyde in PBS (8 mL): 2 mL of 16% formaldehyde (w/v), 6 mL of PBS. The solution can be stored at 20 °C for several months. 8. 5% (w/v) formaldehyde in PBT: 3.1 mL of 16% formaldehyde solution, 6.9 mL of PBT. 9. TBE 10X solution: 60.55 g of Tris base, 30.9 g Boric acid, and 3.7 g of EDTA. Adjust volume to 500 mL with ddH2O. Store at RT indefinitely. 10. Gel for denaturing urea polyacrylamide gel electrophoresis (Urea PAGE). Mix 6 g urea, 1.25 mL TBE 10X, and 3.5 mL of ddH2O. Heat the solution at 60 °C in a water bath until the urea is dissolved. Add 3.125 mL of acrylamide/bisacrylamide, 75 μL of APS 10% and 15 μL of TEMED. Cast the polyacrylamide gel in 0.75 mm thick spacers. Prepare freshly. 11. RNA hybridization solution (RHS, 250 mL): 125 mL of formamide, 62.5 mL of 20X SSC, 1.25 mL of 10 mg/mL heparin, 2.5 mL 10% Tween-20 (v/v), 2.5 mL of 10 mg/mL of salmon sperm and 56.5 mL ddH2O. Prepare 50 mL aliquots and stock at -20 °C for several months. 12. RNA probe preparation: 2 μL of RNA probe in 250 μL of RHS, incubate at 85 °C on a dry bath incubator for 2.5 min. Freshly prepare the RNA probe, keeping it on ice no more than an hour before its use.

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13. Maleic acid buffer (200 mL): 2.3 g of maleic acid, 1.7 g of NaCl, 1.2 g of NaOH and 100 mL of ddH2O. Measure pH and adjust to pH = 7.5 with 5 M NaOH. Make up the volume to 200 mL with ddH2O and filter. Solution can be stored at RT for up to 6 months. 14. 5X blocking solution: 10 g of blocking reagent, 50 mL of maleic acid buffer. Agitate and heat until complete dissolution. Complete to 100 mL with a maleic acid buffer. Autoclave and make 10 mL aliquots. 15. 1X blocking solution: dilute one fifth the 5X Blocking solution with PBT. 16. RNAse A solution: 10 mg vial in 1 mL of ddH2O (100X). Make small aliquots and store at -20 °C for up to a year. 17. 50% (w/v) Dextran sulfate: 25 g of dextran sulfate, 40 mL of ddH2O. Heat to 37 °C until it fully dissolves and then add ddH2O to a final volume of 50 mL. The solution can be stored at 4 °C for several months. 18. 100 mM NaH2PO4 pH = 7 (50 mL): 0.78 g of NaH2PO4, 30 mL of ddH2O. Adjust to pH = 7, and complete with ddH2O to 50 mL. Pass it through a 0.22 μm filter. The solution can be stored at 4 °C for several months. 19. Pre-hybridization mixture (pHM) 50% formamide: 4× SSC, 100 mM NaH2PO4, pH = 7, 0.1% Tween 20. Prepare freshly. 20. DNA hybridization solution (DHS): 5 mL of formamide, 2 mL of 50% (v/v) dextran sulfate, 1 mL of 20X SSC, 500 μL of salmon sperm (10 mg/mL), and 1.5 mL of ddH2O. Store at -20 °C for up to several months. Prewarm at 37 °C before use. 21. 50% (w/v) glucose (40 mL): 20 g of glucose, 30 mL ddH2O. Heat to 60 °C until it dissolves and then add ddH2O to 40 mL. The solution can be stored at RT for several months. 22. 1 M NaCl solution (50 mL): 2.92 g of NaCl, 30 mL of ddH2O. Complete to 50 mL with ddH2O and pass it through a 0.22 μm filter. The solution can be stored at RT for several months. 23. 1 M Tris-HCl pH = 8 solution (50 mL): 6 g of Tris base, 30 mL of ddH2O. Using a pH meter, slowly add HCl using a glass Pasteur pipette to reach the desired pH. Complete to 50 mL with ddH2O, and pass it through a 0.22 μm filter. The solution can be stored at RT for several months. 24. 55 mM NaCl in 11 mM Tris-HCl pH = 8 solution (50 mL): mix 2.75 mL of 1 M NaCl solution, 0.55 mL of 1 M Tris-HCl pH = 8 solution, and 46.7 mL of ddH2O. Prepare freshly.

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25. Gloxy solution (1 mL): 50 mg glucose oxidase, 100 μL catalase, and 900 μL of 55 mM NaCl in 11 mM Tris-HCl pH = 8. Make 60 μL aliquots and store at -20 °C. Solution is stable for several months. Defrost on the day of the experiment and keep it on ice until use. If there is a precipitate, spin it down and employ the supernatant. Once defrosted, the aliquot should be used within 1 week. 26. Hi-M wash buffer (100 mL): 10 mL of 20X SSC, 40 mL of formamide and make up to 100 mL with ddH2O (final concentration of 40% v/v formamide) and filter. 27. Imaging buffer: 1.1 mL of 50% (w/v) glucose, 9.9 mL of PBS, and 110 μL of Gloxy. Add Gloxy solution just before using the solution and mix. Once the tubing is introduced, add a layer of mineral oil to prevent contact with oxygen from the ambient. Replace after 12–15 h. 28. Chemical bleaching solution (10 mL): 0.5 mL of TCEP, 9.5 mL of 2X SSC. Prepare the solution right before its use and discard any remaining solution. 2.3 Wide-Field Epifluorescence Microscope Setup

We have implemented Hi-M in both a widefield and a confocal microscope with very similar hardware configurations. Both setups are controlled by the same software interface (Qudi-HiM) with minor adaptations. For simplicity, we will describe in this chapter our home-made widefield microscope system (Fig. 4a–c), built around a RAMM chassis (Applied Scientific Instrument), equipped with a 60x water-immersion objective (Plan-achromat NA = 1.2) and a sCMOS camera (Orca Flash 4.0v3, Hamamatsu). Our setup uses a standard 200 mm Nikon tube lens (C60-TUBE-B) to achieve a 106 nm effective pixel size, leading to a ~ 217 × 217 μm2 field of view. To avoid water evaporation during Hi-M experiments, we use immersion oil for imaging (Zeiss, Immersol™ W). Wide-field epifluorescence illumination is achieved using 405/488/561 nm (OBIS-405/488, Sapphire 561, Coherent) and 641 nm (VFL-P-1000-642-OEM1-B1, MPB) lasers combined with an acousto-optic tunable filter (AOTFnC-400.650, AA optoelectronics). To avoid the use of a mechanical filter wheel, separation between laser excitation and fluorescence emission is achieved using a four-band dichroic mirror (zt405/488/561/640rpc-UF2, Chroma) combined with a four-band emission filter (ZET405/ 488/561/640 m, Chroma). Sample displacement and FOVs/ROIs selection are performed using a three-axis motorized stage (MS2000, Applied Scientific Instrumentation). Finally, the objective lens is mounted on a single-axis piezo-stage (Nano-F100, Mad City Labs Inc.), allowing for a nm-precision control of the objective axial position during z-scan and focus stabilization.

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Fig. 4 The microscope design. (a) Our homemade Hi-M setup was constructed using a RAMM microscope (gray curved boxes). The lasers are combined by dichroic mirrors (DM) and redirected to the microscope by mirrors (M). The beams are expanded by a telescope and focused at the back focal plane of a 60X objective by lenses L1 and L2. An acousto-optic filter (AOTF) is used to switch on/off the different laser lines and to change their intensity. The emitted light is filtered and focused on the sCMOS camera using a tube lens. The sample is placed in an FCS2 fluidic chamber and the buffer flow is controlled by a pump (vacuum or peristaltic) and an online flow unit that constantly monitors the flow rate at the chamber outlet. Valves are used to select the type of liquid to be injected into the chamber (e.g. buffer, fluorescent probes, etc.). Liquid is drawn from the inlet of a valve, passes through the FCS2 flow chamber, the online flow unit, and is discarded into a waste bottle. A custom-built delivery robot with an XYZ translation stage is used to guide a needle into different tubes (see rainbow tubes organized on a tray), allowing selection of the liquid to be injected. The microfluidic tube is shown in light blue. (b) Illustrative picture of the custom-made delivery robot for readout probe injections. (c) Picture of our home-made epi-fluorescence setup. (d) Overview of hardware devices that are controlled and synchronized by Qudi-HiM. The software controls all hardware required for imaging (e.g. camera, filters wheel), sample positioning (motorized and piezo stages) and fluidic injection (pumps, valves and motorized stages). Acquisition boards (DAQ and FPGA) are used to optimize the speed and reproducibility of specific tasks, such as the acquisition of large 3D image stacks. (Figure adapted from Ref. [22]).

A home-made focus stabilization system is used to compensate for the axial drift in real time. A 785 nm laser beam (OBIS-785, Coherent) is focused on the back-focal plane of the objective, reaching the coverslip/sample interface in near-TIRF illumination conditions. The position of the reflected beam is then measured on a position-sensitive detector (OBP-A-4H, Newport), and any variation in the objective-sample distance above 100 nm is automatically compensated by repositioning the objective lens using the single-axis piezo stage.

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Image acquisition, sample positioning, and liquid handling are controlled by a custom-made software package (Qudi-HiM) described in a dedicated section below. 2.4 Hi-M Sequential Hybridization

The fluid handling circuit design was implemented as described in Cardozo Gizzi et al. [7, 19]. The sample is mounted in an FCS2 chamber and flow is generated using a pump (either negative pressure or peristaltic). An online flow unit is used to continuously monitor the flow rate. This allows precise control of the injected volumes and maintains a steady flow in the chamber. The complete injection sequence (including buffer type, volume, flow-rate, incubation time, etc.) is defined by the user using Qudi-HiM (Fig. 5). For each sequence, readout probes are first selected using a custom-built delivery device consisting of three single-axis motorized stages (VT-80, PI) that allow X-Y selection of individual Eppendorf tubes and Z translation of a needle to pump the barcode solution into the FCS2 chamber (Fig. 4a, b). Up to 100 different readout probes can be sequentially injected during a Hi-M experiment using this system. All other buffers are then injected sequentially by selecting a specific outlet from the eightway HVMX8-5 valves.

2.5

Software packages needed:

Software

1. Library design: https://github.com/HiM-public-resources/ oligopaint-design 2. Microscope control: https://github.com/NollmannLab/ qudi-HiM 3. Data analysis. (a) MATLAB code: https://data.mendeley.com/datasets/ 5f5hd9yj3z/1#folder-26d1f8c0-fc58-4b87-8c4f-cd8a2 94a555 (b) Python code (pyHiM): https://github.com/marcnol/ pyHiM

3

Methods

3.1 Design of Oligopaint Libraries

1. Clone the repository from https://github.com/HiM-publicresources/oligopaint-design, and download the .bed files containing the sequences of previously mined oligos covering the Drosophila non-repetitive genome (https://oligopaints.hms. harvard.edu/genome-files) [15, 20, 21]. 2. In the input_parameters.json file, enter the various parameters required, such as the chromosome of interest, the location of the .bed files, the start-end coordinates of the target region, the desired number of loci, their genomic size, and the minimum number of primary probes per locus.

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Fig. 5 Modules in Qudi-HiM. (a) Basic imaging module interface that allows real-time imaging of the sample. The user can adjust camera, illumination (laser & brightfield), and filter settings to optimize imaging conditions. Images can also be saved as needed. (b) The ROI selector module interface is used to manually or automatically select the ROIs/FOVs that will be acquired during a Hi-M experiment. (c) The Focus module is used to initialize and calibrate the autofocus procedure. Any axial drift is compensated during a Hi-M experiment to ensure reproducibility of data acquisition. (d) The Fluidics Handling Module controls all valves and pumps for buffer/probe injections. It also controls the custom-built delivery robot for the readout probes. The Z-positions of the injection needle and the X-Y-positions of the tubes are specified by the user. (e) The

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3. The Library_Design.py script will (i) calculate the genomic coordinates for each barcode (start, end), (ii) select primary probe sequences for each locus, (iii) concatenate primary sequences with readout sequences and with universal primers, and (iiii) check the homogeneity of the size of the different barcodes. 4. Several output text files are created after running Library_Design.py. (a) Library_Summary.csv contains a table summarizing the information for each barcode (number, start-end position, readout probe, primer forward, primer reverse, number of probes). (b) outputParameters.json is a JSON dictionary file containing the parameters used to generate the library. (c) Full_sequence_only.txt contains the raw primary probe sequences for the oligos used to order the microarray. 5. It is possible to embed multiple libraries within one oligopool by using different sets of universal primers. 6. Order the microarray from an oligopool synthesizer company. 3.2 Amplification of Oligopaint Libraries

1. Emulsion PCR: This step is performed to amplify the starting oligopool (which can be limiting) in a non-biased manner. Set up a PCR Master Mix for each library as indicated in Table 1 and keep it on ice until needed. Pre-chill a 1.8 mL cryotube in the freezer, place it on the center of a controlled stirring plate, and then add a pre-cooled stirring bar to the cryotube. Transfer 600 μL of PCR oil phase to the tube with a positive displacement pipette. Stir at 1000 rpm for at least 1 min. While the stirring bar is still spinning, add 100 μL of PCR master mix in steps of 20 μL increments using a P20 pipette (i.e., dispense 20 μL 5 times). Stir at 1000 rpm for 10 minutes, the emulsion should appear milky white and foamy. Transfer the emulsion to a PCR strip tube (~8 × 75 μL) with a positive displacement pipette. Forward primer is the 5′ = > 3′ whereas reverse primer is the reverse complement of the reverse universal priming sequence. Emulsion preparation must be performed in a cold room at 4 °C. All the equipment must be put there in advance to cool it down before use. It will not be possible to transfer the whole emulsion volume to the PCR strip tube, quality over quantity here.

ä Fig. 5 (continued) Injection Configurator interface is used to define all injection parameters. In particular, the lists of available buffers and readout probe names. And the injection sequences for the hybridization and photobleaching steps of a Hi-M experiment. (Figure adapted from Ref. [22])

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Table 1 Emulsion PCR master mix Reagent

Quantity (μL)

ddH2O

79

10X Taq buffer

10

BSA (10 μg/μL)

5

dNTPs (10 mM)

2

Forward primer (100 μM)

1

Reverse primer (100 μM)

1

Kapa polymerase enzyme (5U/μL)

1

ssDNA library (10–30 ng/μL)

1

2. Perform the PCR using the following cycling conditions: 95 ° C, 2 min, 30× (95 °C, 15 s, 60 °C, 15 s, 72 °C, 20 s), 72 °C, 5 min. PCR products can be stored at 4 °C for a few days. 3. Small-scale emulsion PCR breaking. Pool the emulsion PCR reactions in a 1.5-mL Eppendorf tube. Add 1 μL of gel loading buffer to visualize the aqueous phase. Add 200 μL of mineral oil and vortex for 30 s. Centrifuge at maximum speed for 10 min and remove the upper organic phase. 4. Add 1 mL of water-saturated diethyl ether and vortex for 1 min. Centrifuge at maximum speed for 1 min and remove the diethyl ether upper phase. 5. Add 1 mL of water-saturated ethyl acetate and vortex 1 for min. Centrifuge at maximum speed for 1 min and remove the ethyl acetate upper phase. 6. Repeat step 4. Evaporate the residual diethyl ether by incubating the tube at 37 °C for 5 min with the cap open. The final volume should be around 80 μL. PCR products can be stored at 4 °C for a few days. 7. Purify the DNA by using Zymo Oligo Clean & Concentrator kit. Mix 80 μL of DNA from the emulsion PCR breaking, 160 μL of oligo binding buffer, and 320 μL of ethanol. Homogenize the solution by pipetting up and down ten times. Follow the manufacturer’s instructions up to the DNA elution. Repeat elution with an extra 15 μL of water. 8. Quantify DNA concentration using a NanoDrop by directly taking 2 μL of purified PCR product. Concentration should be between 20 and 40 ng/μL.

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Table 2 Small-scale limited-cycle PCR setup Product

Quantity per tube (μL)

Kapa buffer A

5

dNTP (10 mM)

1

Forward primer (100 uM)

0.5

Reverse primer + T7 promoter (100 uM)

0.5

Template emulsion PCR (1 ng/μL) obtained in step 7

2.5

Kapa polymerase enzyme (5U/μL)

0.5

ddH2O

Make up to a final volume of 50 μL

9. Run a gel electrophoresis to check for a single band amplification with 200 ng of PCR product in a 1.5% agarose TAE gel with 0.01% SYBR Safe at 100 V for 45 min. Purified products can be frozen at -20 °C for several months. 10. Perform the small-scale limited-cycle PCR by setting up the following reaction mix for eight tubes as indicated in Table 2. The limited number of cycles is performed to find the cycle number where the PCR is still at its exponential phase (Fig. 2c). Perform this step before proceeding to the largescale PCR. The T7 promoter sequence (5′- TAATACGACT CACTATAGGGT-3′) should be added to the reverse primer used for the emulsion PCR step to allow for the reverse transcription step. 11. Run the following PCR program: 95 °C, 5 min, 15× (95 °C, 30 s 60 °C, 45 s, 72 °C, 30 s). Pick up the corresponding tube after each of the cycles 8–15 just after the extension phase. To do so, quickly open the PCR machine, remove the corresponding tube, close the lid, and resume the program. PCR products can be left overnight (ON) at 4 °C or frozen for up to a month at -20 °C. 12. Run 20 μL of the PCR product in a 1.5% agarose gel with 0.01% SYBR Safe at 100 V for 45 min. Find the cycle with a single band of the expected size and the maximum intensity (Fig. 2c). 13. Perform a large scale limited-cycle PCR (Table 3) by running a reaction mix for 16 tubes as indicated in Table 2. This step will generate a big quantity of Oligopaints. The T7 promoter sequence should be added to the reverse primer to allow for the reverse transcription step.

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Table 3 Large-scale limited-cycle PCR mix Product

Quantity (μL)

Kapa buffer A

80

DNTP (10 mM)

16

Forward primer (100 μM)

8

Reverse primer + T7 promoter (100 μM)

8

Kapa enzyme (5U/μL)

8

ddH2O

600

Template emulsion PCR (1 ng/μL)

40

14. Split the volume of the mix in Table 2 into 16 × 50 μL PCR tubes and run the PCR program from step 11 using the optimized number of cycles determined in step 12. Add a last extension cycle of 5 min at 72 °C. PCR product can be safely stored for months at -20 °C. 15. Run 20 μL of the PCR product in an agarose gel as in step 12 to check that the PCR was successful. 16. Collect the 50 μL aliquots from the previous step in a 15 mL falcon tube and proceed to DNA column purification according to the manufacturer’s instructions. Use Zymo DNA purification kit with 25 μg capacity. Elute using 30 μL of DNAseand RNAse-free water. 17. Quantify product concentration with a NanoDrop using double-stranded DNA parameters. This typically requires a 1/10 dilution of a 2 μL aliquot of the purified product. Concentration should be between 30 and 50 ng/μL. 18. Run the remainder of the 1/10 stock dilution in a 1.5% agarose gel as in step 12 (Fig. 2d). Check for a single band of the expected size. 19. Perform in vitro transcription by setting up the reaction mix as indicated in Table 4. This step is a high-yield reaction that further amplifies the template molecules as well as converts them into RNA. It is necessary to keep RNAse-free conditions at all times. 20. Split the volume from the in vitro transcription solution into 3 × 20 μL PCR tubes and incubate at 37 °C for 12–16 h in a thermocycler. In vitro transcription products can be frozen for months at -80 °C. 21. Take 5 μL and purify with a Zymo Oligo Clean & Concentrator kit according to manufacturer’s instructions, using 15 μL of DNAse- and RNAse-free water to elute purified product. The

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Table 4 In vitro transcription solution mix Product

Quantity (μL)

Purified PCR product

6 μg template DNA

ATP (100 mM)

6

UTP (100 mM)

6

CTP (100 mM)

6

GTP (100 mM)

6

10X T7 buffer

6

Rnase inhibitor (40U/μL)

2.25

HiScribe T7 polymerase

6

ddH2O

Make up to a final volume of 60 μL

purification is only performed with a small aliquot to control if the in vitro transcription was successful and to estimate the RNA concentration in the non-purified RNA solution. Use Zymo DNA purification kit with 10 μg capacity. 22. Make a 1/10 dilution to perform a quantification of the purified RNA on NanoDrop using RNA parameters. Concentration should be between 0.5 and 2 μg/μL. The concentration obtained allows us to estimate concentration in non-purified RNA. For instance, a 2 μg/μL concentration in the purified RNA can be translated to an estimated concentration of 6 μg/μ L in the non-purified RNA considering a factor 3 dilution (from a 5 μL aliquot to a final volume of 15 μL). The total yield of the in vitro transcription step should be around 150–450 μg from a single transcription step (60 μL in total). 23. Check for the RNA quality by Urea PAGE (Fig. 2e). Perform a pre-run for 30 min in 1X TBE at 190 V to eliminate the excess of persulfate. When finished, wash the wells with the running buffer. Load 100 ng of purified RNA per lane. Heat the samples at 95 °C for 5 min and put it immediately on ice for 2 min. Perform the PAGE for 1 h at 190 V. For gel staining, incubate protected from light for 20 min in 30 mL of TBE and 3 μL of SyBR Gold. 24. Perform the reverse transcription reaction according to Maxima H Reverse Transcriptase kit by setting up the reaction mix indicated in Table 5. In this step, the non-purified RNA from step 20 is directly used. RNA should always be kept in ice to prevent degradation. Primer sequence is the same as for forward primer used in emulsion PCR or limited-cycle PCR.

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Table 5 Reverse transcription mix Product

Quantity (μL)

Non purified transcription product

150 μg

dNTP mix 100 mM

12

Forward primer (100 μM)

50

5X maxima buffer

240

RNAsin plus (40U/μL)

30

Maxima H reverse transcriptase (200U/μL)

30

ddH2O

Make up to a final volume of 1200 μL

25. Split the volume obtained in the previous step into two 1.5-mL tubes and incubate for 3 h at 50 °C in a water bath. Reverse transcription products can be frozen for months at -20 °C. 26. Perform the RNA degradation by adding into each tube 300 μL of 0.5 M EDTA and 300 μL of 1M NaOH and incubating at 95 °C for 15 min in a water bath. This step allows to selectively degrade the RNA while keeping single stranded DNA. 27. Take a 10 μL aliquot to control for DNA concentration and to perform a gel electrophoresis as in step 12. 28. DNA probe purification. Mix the two aliquots in a sterile 50 mL Falcon tube. Add 4.8 mL of oligo binding buffer and 19.2 mL of ethanol. Homogenize and evenly distribute across two columns. Follow the manufacturer’s instructions from this point on. Use the Zymo DNA purification kit with 100 μg capacity. 29. Take a 10 μL aliquot to measure DNA concentration and to perform a gel electrophoresis as in step 12. 30. Ethanol precipitation: Directly add to the 150 μL DNA elution, 24 μL of 5M ammonium acetate, 6 μL of glycogen, and 750 μL of 100% (v/v) ethanol at -20 °C. Vortex and incubate 1 h at -80 °C or overnight at -20 °C. Centrifuge at 13,000 G for 1 h at 4 °C. Discard the supernatant and wash the pellet with 1 mL of ice-cold 70% ethanol (v/v). Centrifuge at 13,000 G for 15 min at 4 °C. Discard the supernatant and add 20 μL of DNase- and RNase-free water. Let the singlestranded DNA (ssDNA) resuspend for 10 min at 37 °C with agitation. Keep on ice. 31. Quantify oligo concentration with a NanoDrop using ssDNA parameters. Total quantity of ssDNA should be in the order of 80–120 μg.

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32. Control the quality of ssDNA by Urea PAGE as in step 23 (Fig. 2e). This step allows to verify RNA degradation and the efficacy of reverse transcription step. Probes can be stored at -20 °C for months. 3.3 Sample Preparation and Fixation

1. Place 200–400 flies with a 2:1 female/male ratio into an egg-collection cage mounted with an apple juice plate containing a dollop of yeast paste and prewarmed to 25 °C (or the temperature required for the specific experiment). Perform an overnight (ON) prelaying step. 2. Replace the plate with a new one containing a dollop of yeast paste and prewarmed to 25 °C. Perform a laying step during 1.5 h at 25 °C. 3. Remove the plate, put the cover, and incubate 1 h (or the time required to obtain embryos in the desired developmental stage) at 25 °C. 4. Rinse the plates with ddH2O and carefully detach embryos using a paintbrush. Filter the liquid using a nylon filter. Embryos will remain on it. Rinse the nylon filter with water to remove yeast, and remove excess water filter by blotting dry on a paper towel. 5. Prepare one well plate containing bleach at 2.6% active chlorine in water. Put the filter with the embryos in the bleach containing well and incubate for 5 min. Use distilled water to thoroughly rinse the embryos. Ensure all embryos are in the medium by rinsing the walls of the nylon filter. 6. In a 20 mL glass vial, add 5 mL of 4% (v/v) formaldehyde in PBS and 5 mL of heptane. Formaldehyde is toxic and should be handled with protective gloves under a fume hood and discarded according to the relevant environmental and safety instructions. 7. Transfer the embryos to the vial using a paintbrush. 8. Close the vial and vigorously shake it manually for 30 s. You may cover the cap of the vial with parafilm to avoid leakage of the formaldehyde and heptane solution inside. Incubate the embryos for 20 min at RT. Embryos dechorionated will float between the two phases. 9. Using a glass Pasteur pipette, aspirate the lower aqueous phase from the bottom of the vial together with the embryos contained therein (non-dechorionated embryos). 10. Add 5 mL of methanol and vortex the glass vial for 15 s. Using a glass Pasteur pipet, transfer the embryos at the bottom of the glass vial to a 1.5-mL tube. Methanol is toxic and highly volatile and should be handled with protective gloves under a fume hood and discarded

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according to the relevant environmental and safety instructions. Glass pipet is used to avoid embryos attaching to the walls. Avoid using plastic tips. 11. Wash the embryos three times with 1 mL of methanol. Fixed embryos can be stored in methanol at -20 °C for months. 3.3.1 DNA In Situ Hybridization

1. Transfer 100–200 embryos from Subheading 3.3 step 11 to a 1.5 mL tube. Use a glass Pasteur pipette to prevent embryos sticking to the plastic tips. 2. Rehydrate the fixed embryos by incubating them with 1 mL of the following solutions (1) 90% methanol, 10% PBT; (2) 70% methanol, 30% PBT; (3) 50% methanol, 50% PBT; (4) 30% methanol, 70% PBT; (5) 100% PBT. Incubate 3–5 min at RT on a rotating wheel for each step. 3. Incubate the embryos with 1 mL of PBT, 100 μg/mL of RNAse for 2 h at RT or ON at 4 °C in a rotating wheel. 4. Permeabilize the embryos by incubating them with PBS-Tr for 1 h at RT in a rotating wheel. 5. Transfer tissues into pHM by passing embryos through 1 mL of the following freshly made solutions: (1) 80% PBS-Tr, 20% pHM; (2) 50% PBS-Tr, 50% pHM; (3) 20% PBS-Tr, 80% pHM; (4) 100% pHM. Incubate 20 min at RT on a rotating wheel for each step. Before exchanging solutions, allow the embryos to settle down 2–3 min. 6. Prepare a primary DNA probe by adding 45–225 pmol of Oligopaint probe to 25 μL of DHS. Keep the mix on ice. Denature primary DNA probe by incubating for 15 min at 80 °C in the Thermomixer. The amount of probe employed should be tested by quantifying the efficiency of labeling vs. increasing concentrations of DNA probe. 7. Carefully remove the pHM solution from the embryos tube and add 1 mL of fresh pHM. Denature embryonic DNA by incubating for 15 min at 80 °C in a water bath. 8. Carefully remove the pHM solution from the embryos tube and add 25 μL of the probes. Mix by gently flicking the tube with a finger. Carefully add 40 μL of mineral oil. Change the water bath temperature to 37 °C and incubate the embryos ON at 37 °C in the water bath. Mineral oil layer is added on top to prevent evaporation. Allowing the embryos to slowly cool down from 80 °C to 37 °C in the water bath greatly increases efficiency of labeling. 9. Carefully remove as much mineral oil as possible from the tube with a P20 pipette. Remaining oil dramatically affects embryo attachment to coverslips, as well as interfere with image acquisition.

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10. Add 500 μL of 50% formamide, 2× SSC, 0.3% CHAPS, let the embryos sink, and remove supernatant. This helps to immediately remove the mineral oil after hybridization. If the quantity of remaining oil is too big, this step will not be enough to prevent posterior issues of attachment and image acquisition. 11. Perform post-hybridization washes by passing embryos through 1 mL of the following freshly made solutions: (1) 50% formamide, 2× SSC, 0.3% CHAPS; repeat this wash once; (2) 40% formamide, 2× SSC, 0.3% CHAPS; (3) 30% formamide, 70% PBT; (4) 20% formamide, 80% PBT; (5) 10% formamide, 90% PBT; (6) 100% PBT; (7) 100% PBS-Tr. Perform washes (1)–(4) 20 min at 37 °C in a thermomixer with agitation (800–900 rpm), perform washes (5)– (7) 20 min at RT on a rotating wheel. 12. (Optional) Rinse the embryos with 1 mL of PBT. Crosslink primary library by incubating the embryos with 1 mL of 4% (w/v) paraformaldehyde in PBT for 10 min at RT in a rotating wheel. Although optional, in our hands cross linking the primary library improved the labeling efficiency. 13. Rinse the embryos with 1 mL of PBT. Incubate the embryos with DAPI for 20 min at RT in a rotating wheel. Afterward, remove the DAPI and rinse the embryos three times with 1 mL of PBS. Detergent-containing PBT can prevent embryos from attaching to the coverslip. DNA-labeled embryos can be stored for weeks at 4 °C before proceeding to mounting and imaging. 3.4 Hi-M Experiment Preparation and Data Acquisition with QudiHiM 3.4.1

Qudi-HiM Modules

Qudi-HiM [22] (https://github.com/NollmannLab/qudi-HiM) is a user/microscope interface written in Python 3 for automated Hi-M data acquisition. It is based on a software package called Qudi [23], a modular experiment management suite specifically designed for the development of custom user interfaces. Qudi provides a user-friendly framework with all the core functionality needed to build robust acquisition software, including error handling, hardware interfacing, and design of custom acquisition pipelines. QudiHiM was built on top of this framework by adding all the functionality needed to set up and run a Hi-M experiment. Qudi-HiM was designed as an open-source and flexible acquisition software that can be transferred to different Hi-M setups with a minimum of work. A list of all hardware already supported by Qudi-HiM can be found at https://github.com/NollmannLab/ qudi-HiM/blob/master/documentation/qudi-cbs%20documen tation/qudi-HiM_hardware/overview_hardware_qudi-him.md (Fig. 4d). It includes hardware used for image acquisition (lasers, filters, detectors, etc.), sample positioning (motorized and piezo stages), and sequential labeling (valves, pumps, and the injection robot). This list is not restrictive and any new device can be used as long as it is interfaced in Qudi-HiM.

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The Hi-M preparation procedure has five modules, each associated with a specific operation and a custom user interface (see Fig. 5): • The “Basic Imaging” module is dedicated to image acquisition. When setting up an experiment, this module is used to control the quality of the sample and tuning the acquisition parameters (laser power, acquisition time, etc.). • The “ROI selector,” together with the “Basic imaging,” module is used to select and save the ROIs on the sample (also referred as FOVs in the text). • A typical Hi-M experiment can last several days. In order to avoid any loss of focus due to mechanical and thermal drift during the acquisition, a “Focus” module was implemented to define an axial reference for the focus. • The “Fluidics Control” module is handling all the functionalities related to buffer injection and rinsing. It also controls the custom-built delivery device for injecting the readout probes (Fig. 4b). • Finally, the injection procedures (hybridization, photobleaching, number of probes, etc.) are defined in the “Injection configurator” module. Two other modules are available: • A module called “Task Manager” is used to launch custom acquisition scripts for Hi-M experiments. For instance, we use slightly different Hi-M acquisition conditions depending on sample types or microscopy setups. • For simplicity, each experiment is associated with a unique list of user-defined parameters such as laser wavelengths and powers, folder name, acquisition time, ROIs/FOVs list, etc. In the “Experiment Configurator” module, the user specifies all the acquisition parameters after selecting the type of experiment to be performed. The main steps needed for a full Hi-M acquisition are highlighted in Fig. 3 and described in detail below. Bear in mind that adaptations may be needed depending on your setup and sample. 3.4.2

Experiment Setup

1. The first step requires setting up the microscope and QudiHiM (Fig. 3, Experiment setup). Before switching on the lasers, check that all shutters are closed. When laser 785 nm emits, wear protection goggles. For reproducibility purposes, make sure laser powers are always the same at the beginning of each experiment and that the lasers are properly aligned. 2. Open Qudi-HiM and click on Load all modules. Wait for all modules to be loaded and check there are no errors (they will

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appear in red in the main Qudi-HiM window). In the event an error might occur, check that all hardware is properly connected and switched on. Repeat the previous step. 3. Between experiments, microfluidics tubing should remain filled with 50% ethanol to prevent bacterial growth and facilitate air bubble removal. Therefore, the first step for setting up an experiment consists of rinsing all the microfluidics tubing with ddH2O and then with filtered 2XSCC. This part can be done manually using a syringe or using the Fluidics handling module from Qudi-HiM. Similarly, at the end of the experiment, rinsing steps need to be repeated twice, once with ddH2O first, then once with 50% ethanol. 4. Place the sample coverslip in the FCS2 chamber. Ensure that all valves are closed during the process to avoid leaks and air bubbles: (a) Open the chamber and dry the Teflon spacer with a clean tissue. Dry the microfluidic chamber and sample coverslip as much as possible. (b) Replace the dried spacer and mount the coverslip with the attached sample on the inside. (c) Slowly fill the chamber with 2× SSC. Hold the chamber upright to avoid air bubbles. Before placing the sample on the microscope, check that the chamber is properly sealed by flowing 2× SSC at 150 μL/ min for 10 min and make sure that no leakage is detected and no air bubble enters the chamber. 5. Using the Fluidics module of Qudi-HiM: (a) Double check that all valves and pumps are working by checking the flow rate stability (no blockage or air bubble). (b) For sequential barcode injections, initialize the custommade delivery device by defining the reference position of the first barcode tube. Make sure the needle is properly centered in the tube and ~ 1 mm from the bottom. Using the motorized stages, adjust the XYZ positions of the needle if necessary. To avoid the formation of an air bubble in the needle, make sure it is always immersed in liquid. 6. Place a drop of immersion oil on the objective and focus on the coverslip surface using the Basic imaging module. We use Live Mode with Brightfield on to locate the sample. By changing the flow in the FCS2 chamber (e.g., manually with a syringe), check that the sample is stable and there is no risk of detachment during the experiment. If the sample detaches at this stage, then the experiment needs to be restarted.

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7. Using the ROI selector module, initialize a new list of FOVs/ ROIs by clicking Start new ROI list. ROIs are selected (typically 10–30) using a motorized X-Y stage (with a joystick for ease of use), and the stage position can be displayed in real time with tracking mode on. In our case, we select ROIs by imaging DAPI-stained nuclei and/or RNA expression patterns. By clicking on New ROI, the position of the X-Y stage is recorded in the ROI list and its position is displayed in the main window. When finished, change the file name and click Save ROI list. 8. If necessary, an optional photobleaching step can be performed to reduce the fluorescence background of the sample. In the Experiment configurator module, select Photobleaching, and put lasers 488 nm, 561 nm, 640 nm at 100% (unless you have some labeling you don’t wish to bleach). In Load ROI list, indicate the appropriate ROI file, set the bleaching time (e.g., 2 min per ROI), and press Save configuration. Go to Task runner in Qudi-HiM manager, click Photobleaching and press Run. 3.4.3

Mask Imaging

1. The second step in a Hi-M experiment is acquiring DAPI staining and their corresponding fiducial images (Fig. 3, mask imaging). In the Injection configurator module, indicate to which valves the different buffers are assigned by clicking on Add buffer. (a) For example: (i) Valve #7: probe (ii) Valve #1: SSC (for the 2× SSC) (iii) Valve #2: WB (for the Hi-M wash buffer) (iv) Valve #3: IB (for the imaging buffer) In our setup, valve #7 is connected to the custom injection robot with a tray that allows us to organize and inject up to 100 different fluorescent probes/reagents (Fig. 4b). (b) Prepare the fiducial in a 2 mL tube (see Reagent setup, readout probe solution). Place the tube on the tray, on position #1 (reference position). Using the Fluidics module, move the injection needle by clicking on Go to target. Make sure the needle goes to the first tube. (c) Prepare a DAPI staining solution (see Reagent setup, DAPI solution). Place the tube on the tray, on position #2. (d) In the Hybridization manager of the Injection configurator module, set up the injection protocol (see Table 6, steps 1–6). It should appear as follows:

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Table 6 DAPI-fiducial labeling Step

Solution

Volume (μL)

Flow rate (μL/min)

1

Fiducial probe

1500

150

2

Incubation time 900 s

3

Hi-M wash buffer

1500

150

4

2X SSC

1000

150

5

DAPI solution

1500

150

6

Incubation time 1800s

7

2X SSC

1000

150

8

Imaging buffer

900

150

(i) Probe, 1500 μL, 150 μL/min = injection of tube in position #1 (fiducials) containing fluorescently labeled imaging oligos complementary to the universal priming region. (ii) Incubation time: 900 s (iii) Hi-M WB, 1500 μ, 150 μ/min (iv) 2X SSC, 1000 μL, 150 μL/min (v) Probe, 1500 μL, 150 μL/min = injection of tube in position #2 (DAPI) (vi) Incubation time: 1800 s Save the fluidics settings by clicking on Save Injections. 2. Run the DAPI/fiducial staining procedure. Open the Experiment configurator and select the experiment handling only injections (e.g., Fluidics RAMM). This experiment requires only one parameter, the path to the fluidics settings saved in step 9. Click on Save configuration and launch the experiment from the Task runner module. 3. Using the Basic imaging module, briefly check the fluorescence intensity of the DAPI and fiducial markers, without photobleaching the signal. If the signal is too low, repeat step 9. Otherwise, perform the following injections manually using the Fluidics control module (Table 6, steps 7 and 8): (a) 1000 μL of 2× SSC (b) 900 μL of imaging buffer This step is critical because the fiducial signal is used to correct for sample drift during postprocessing. The imaging buffer is used to minimize photobleaching and ensure that the fluorescence intensity of the fiducial probe remains stable throughout the experiment.

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4. The autofocus must be calibrated before the acquisition can begin. Using the Basic Imaging module in live mode, select the plane where the acquisition will start. As a rule of thumb, we usually define this reference plane ~1 μm below the z-position where the first fluorescence signal is detected. This is to avoid any variability in sample position between ROIs and to ensure that we acquire enough planes to perform accurate 3D localization of the fluorescent spots during post-processing. Put on protective goggles and open the shutter for the 785 nm laser. In the Focus module, click on Start Live Display and verify that a clean and bright reflection of the 785 nm laser is detected. Perform the autofocus calibration by clicking Start Calibration. The calibration curve should appear as a straight line and the Precision should indicate a value lower than 50 nm. Finally, click Define Offset for Focus Search to complete the autofocus calibration procedure. 5. Image acquisition of fiducial and DAPI. Open the Experiment configurator and select the ROI multicolor scan experiment. This experiment was designed to acquire a 3D stack of images for each ROI selected by the user. Tick Dapi to automatically add “DAPI” to the names of the saved images. Indicate the name of the experiment as well as the exposure time (e.g., 0.05 s). Compose the imaging sequence to be performed. For instance, for DAPI and fiducial imaging, we use the following settings: (a) 405 nm to 10% (b) 561 or 488 nm to 20–60% (depending on the fluorescent label used for the fiducial imaging oligo) This means that two images are acquired sequentially for all planes of the 3D stack: the first with the 405 nm laser (DAPI) and the second with the 561 nm laser (fiducial). Specify the number of planes and the spacing required for the 3D stack (e.g., 65 planes separated by 0.25 μm), as well as the path to the file where the ROI positions were saved (see step 7). Example DAPI-stained nuclei are shown in Fig. 6a. Finally, save these parameters and launch the acquisition from the Task runner module. Before moving to the next part, check the quality of the DAPI and fiducial signals. Make sure the quality of both images is high enough to enable mask segmentation and correct fiducial registration. 3.4.4

Cycle Imaging

1. Make sure there is enough buffer for the entire experiment, depending on the number of cycles (typically a few mL per cycle). 2. Make sure there is enough space on the local computer disk to store the data. In our case, a typical experiment is 0.5–1 TB.

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Fig. 6 Typical raw images of DAPI, barcodes, fiducials. Image of the outcome of a typical Hi-M experiment. (a) Raw images of DAPI-labeled nuclei from different embryos in six FOVs. Images correspond to single z-planes. Image size: 217 × 217 μm. (b) Representative images of fiducial labeling in the same FOV, acquired in three different cycles. Images correspond to single z-planes. Image size: 217 × 217 μm. (c) Representative images of DNA-FISH spots for different barcodes in the same FOV, acquired in different cycles. Images correspond to single z-planes. Image size: 217 × 217 μm.

3. Place the 2 mL tubes with the different readout probes on the injection robot tray. 4. In the Injection configurator module, in the Probe Name section, enter the names of each readout probe (RT) (e.g., RT01, position 1; RT02, position 2) in the same order as the tubes are organized on the tray. In the Hybridization Manager section, enter the injection sequence as described in Table 7. In the

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Table 7 Readout probe labeling for Hi-M experiment Step

Solution

Volume (μL)

Flow rate (μL/min)

1

Readout probe solution (RT)

1500

150

2

Incubation time 900s

3

Hi-M wash buffer

1500

150

4

2X SSC

1000

150

5

Imaging buffer

900

150

Table 8 Photobleaching for Hi-M experiment Step

Solution

Volume (μL)

Flow rate (μL/min)

1

TCEP

1000

150

2

2X SSC

1000

150

Photobleaching Manager section, indicate the injection sequence used to photobleach the readout probes (see Table 8). Save these parameters in a separate file by clicking on Save Injections. 5. In the Experiment configurator module, select Hi-M experiment and fill in all the required parameters: (a) The number of planes and the spacing should be the same as those used for DAPI/Fiducial imaging. The ROI list should also be the same. (b) The laser imaging sequence should be adapted according to the fluorescent dyes used. (c) Indicate the name of the injection parameters file saved in step 17. (d) Check the Transfer data automatically option if you also want the data to be stored on a remote server. If selected, the transfer will be done as a background task between each cycle, during the photobleaching and hybridization injections. In our case, we use this option to store the data on a server and start the deconvolution while the experiment is still running. This allows us to save time on postprocessing. 6. From the Task Manager module, launch the Hi-M experiment. 7. At the end of the experiment, proceed to cleaning as described in step 3, first in ddH2O and then in 50% ethanol for storage.

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Image Analysis

Image analysis was previously described in detail in Refs. [19, 24]. Briefly, these include the following steps: 1. Demix channels and deconvolve 3D images using Huygens. 2. Calculate and apply global registrations to images from all cycles using a reference cycle. 3. Segment DAPI and fiducial masks in 3D. Typical examples of DAPI and fiducial images for different hybridization cycles are provided in Fig. 6a, b. 4. Segment DNA-FISH spots and localize them using 3D Gaussian fitting routines. Typical examples of DNA-FISH images for different hybridization cycles are provided in Fig. 6c. 5. Calculate local registrations when necessary. Apply to 3D DNA-FISH spot localizations. 6. Match DNA-FISH localizations to DAPI/ fiducial masks. 7. Calculate pairwise distance matrices for each combination of barcodes and for each mask. 8. Calculate ensemble pairwise distance and proximity matrices.

Acknowledgments This project was funded by the European Union’s Horizon 2020 Research and Innovation Program (EpiScope, Grant ID 724429, M.N.). We acknowledge the Bettencourt-Schueller Foundation for their prize “Coup d’e´lan pour la recherche Franc¸aise,” and the France-BioImaging infrastructure supported by the French National Research Agency (grant ID ANR-10-INBS-04, “Investments for the Future”). References 1. Nollmann M, Bennabi I, Go¨tz M et al (2022) The impact of space and time on the functional output of the genome, vol 14. Cold Spring Harb Perspect Biol, p a040378 2. Szabo Q, Bantignies F, Cavalli G (2019) Principles of genome folding into topologically associating domains. Sci Adv 5:eaaw1668 3. Mirny LA, Imakaev M, Abdennur N (2019) Two major mechanisms of chromosome organization. Curr Opin Cell Biol 58:142–152 4. Davidson IF, Peters J-M (2021) Genome folding through loop extrusion by SMC complexes. Nat Rev Mol Cell Biol 22:445–464 5. Finn EH, Pegoraro G, Branda˜o HB et al (2019) Extensive heterogeneity and intrinsic variation in spatial genome organization. Cell 176:1502–1515.e10

6. Cattoni DI, Cardozo Gizzi AM, Georgieva M et al (2017) Single-cell absolute contact probability detection reveals chromosomes are organized by multiple low-frequency yet specific interactions. Nat Commun 8:1753 7. Cardozo Gizzi AM, Cattoni DI, Fiche J-B et al (2019) Microscopy-based chromosome conformation capture enables simultaneous visualization of genome organization and transcription in intact organisms. Mol Cell 74: 212–222.e5 8. Bintu B, Mateo LJ, Su J-H et al (2018) Superresolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362:eaau1783 9. Nir G, Farabella I, Pe´rez Estrada C et al (2018) Walking along chromosomes with super-

Chromatin Tracing with Hi-M resolution imaging, contact maps, and integrative modeling. PLoS Genet 14:e1007872 10. Mateo LJ, Murphy SE, Hafner A et al (2019) Visualizing DNA folding and RNA in embryos at single-cell resolution. Nature 568:49–54 11. Liu M, Lu Y, Yang B et al (2020) Multiplexed imaging of nucleome architectures in single cells of mammalian tissue. Nat Commun 11: 2907 12. Wang S, Su J-H, Beliveau BJ et al (2016) Spatial organization of chromatin domains and compartments in single chromosomes, vol 353. Science, pp 598–602 13. Su J-H, Zheng P, Kinrot SS et al (2020) Genome-scale imaging of the 3D organization and transcriptional activity of chromatin. Cell 182:1641–1659.e26 14. Takei Y, Zheng S, Yun J et al (2021) Single-cell nuclear architecture across cell types in the mouse brain. Science 374:586–594 15. Beliveau BJ, Joyce EF, Apostolopoulos N et al (2012) Versatile design and synthesis platform for visualizing genomes with Oligopaint FISH probes. Proc Natl Acad Sci U S A 109:21301– 21306 ˜ o MS et al 16. Beliveau BJ, Boettiger AN, Avendan (2015) Single-molecule super-resolution imaging of chromosomes and in situ haplotype visualization using Oligopaint FISH probes. Nat Commun 6:7147. https://doi.org/10.1038/ ncomms8147 17. Lubeck E, Coskun AF, Zhiyentayev T et al (2014) Single-cell in situ RNA profiling by sequential hybridization. Nat Methods 11: 360–361

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18. Chen KH, Boettiger AN, Moffitt JR et al (2015) RNA imaging. Spatially resolved, highly multiplexed RNA profiling in single cells. Science 348:aaa6090 19. Cardozo Gizzi AM, Espinola SM, Gurgo J et al (2020) Direct and simultaneous observation of transcription and chromosome architecture in single cells with Hi-M. Nat Protoc 15:840–876 20. Beliveau BJ, Boettiger AN, Nir G et al (2017) In situ super-resolution imaging of genomic DNA with OligoSTORM and OligoDNAPAINT. Methods Mol Biol 1663:231–252 21. Beliveau BJ, Kishi JY, Nir G et al (2018) OligoMiner provides a rapid, flexible environment for the design of genome-scale oligonucleotide in situ hybridization probes. Proc Natl Acad Sci U S A 115:E2183–E2192 22. Barho F, Fiche J-B, Bardou M et al (2022) Qudi-HiM: an open-source acquisition software package for highly multiplexed sequential and combinatorial optical imaging. Open Res Eur 2:46 23. Binder JM, Stark A, Tomek N et al (2017) Qudi: a modular python suite for experiment control and data processing. SoftwareX 6:85–90 24. Xavier D, Jean-Bernard F, Marion B, et al (2023) pyHiM, a new open-source, multi-platform software package for spatial genomics based on multiplexed DNA-FISH imaging. h t t p s : // w w w . b i o r x i v . o r g / c o n t e n t / 1 0.1 10 1/ 20 23 . 0 9. 1 9.5 5 84 12 v 1. abstract

Chapter 17 Rapid DNA-FISH in Arabidopsis thaliana Somatic Cells Olga Raskina and Ofir Hakim Abstract Fluorescence in situ hybridization (FISH) technique has been widely used to detect and localize specific DNA and RNA sequences in interphase nuclei and chromosomes in animals and plants. Here, we present a protocol for localization of genomic loci in nuclei of the model plant Arabidopsis thaliana. This protocol includes several advances and adaptations to A. thaliana, including preparation of nuclei and chromosomes without the use of liquid nitrogen, and an in situ hybridization procedure that preserves chromatin structure without the use of paraformaldehyde and formamide. Simultaneous denaturation of the BAC (bacterial artificial chromosome) probe and nuclei followed by annealing at high temperature allows hybridization in less than an hour. These hybridization conditions also provide high signal to noise ratio by a small number of washes. Thus, this simplified in situ hybridization procedure is completed in one working day. Key words Arabidopsis thaliana, BAC-FISH, Chromosome preparation, Interphase nuclei, Fluorescence in situ hybridization, Molecular cytogenetics, Plant

1 Introduction The use of fluorescence in situ hybridization (FISH) and other microscopy-based methods has played a crucial role in unraveling the relationship between the spatial organization of plant genomes and their function and evolution. Recent advances in plant 3D genomics, specifically chromosome conformation capture (3C) techniques, have provided detailed maps of chromosomal spatial associations. These genome-wide maps of averaged snapshot of chromosomal conformations across multiple cells have led to the generation of new hypotheses and models that can be further explored using DNA-FISH in single cells to reveal heterogeneity and variability within the cell population.

Rapid DNA-FISH in Arabidopsis Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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To perform DNA-FISH, the DNA probe is fluorescently labeled using various methods such as nick translation, random primed labeling, or PCR labeling [1]. The labeled probe and target cell DNA are then denatured separately or simultaneously, and both methods are widely used. Combining the denatured probe and target allows the annealing of complementary DNA sequences, and the hybrid DNA locus is detected using a fluorescent microscope. FISH protocols in both animal and plant research typically involve denaturation of DNA probes and cells in formamide and overnight annealing at 37 °C. However, FISH can also be performed in a formamide-free buffer system, where denaturation of DNA probe and target occur simultaneously and subsequent hybridization is conducted at high temperature in a very short time, in the scale of minutes rather than hours [2]. In plants, fast in situ hybridization protocols involving hybridization of DNA probes and target at 63–65 °C during 2 h have been successfully applied to both mitotic and meiotic chromosomes [3–5]. Here, we present a simple and rapid DNA-FISH protocol adapted for model plant Arabidopsis thaliana (Figs. 1 and 2) [6], based on a method described by Shams and Raskina [7]. The protocol involves preparing DNA probes from BAC clones obtained from the Arabidopsis Biological Resource Center (ABRC) (as an example in this chapter, F1H21; Chr5:9,635,755–9,730,934 and F10O11; Chr1: 491,600–603,132), simultaneous denaturation of DNA probes and cells under a coverslip followed by hybridization at 63 °C for 50 min. The entire process, including pre- and post-hybridization treatments and washes, can be completed within a single working day. Notably, this protocol does not require the use of formamide and paraformaldehyde. Additionally, we present an air-drying

Fig. 1 Preparation of cytological slides. (a) Cell suspension in a DDW droplet observed using an upright light microscope at low (×10) magnification, with the iris diaphragm closed and the condenser lowered. The remaining non-meristematic tissues must be removed (red arrows). (b) Assessing the quality of cytological slides after DAPI staining using fluorescent microscope. (b) Low (×20) and (c) high (×100) magnification. Single nucleus is shown in the boxes at low (×20) and high (×100) magnification

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Fig. 2 DNA fluorescence in situ hybridization (FISH) on somatic nuclei (a, c) and chromosomes (b, d) of Arabidopsis thaliana WT col-0 (a–b) and ADF8p:NTF/ACT2p:BirA transgenic plants (c–d). F1H21 probe on chromosome 5 labeled in green, F10O11 probe on chromosome 1 labeled in red, DNA counterstaining with DAPI labeled in blue. The proximity of the two foci in the nucleus and chromosomes of transgenic plants is a result of their linear proximity in the rearranged genome [6]. Chromosomes at anaphase and prometaphase stages are shown for WT (b) and ADF8p:NTF/ACT2p:BirA (d) plants, respectively. FISH foci are indicated with arrows. Enlarged translocated chromosome is shown in the small box (d). (e) Normal chromosomes 1 and 5 (WT col-0 plants) and rearranged chromosomes (ADF8p:NTF plants) are shown in the scheme. The breakpoints for chromosomal rearrangements are indicated with black arrows for WT col-0 plants

method for somatic nuclei and chromosomes preparation in a drop of acetic acid without the use of liquid nitrogen. These steps are based on previously published protocols for A. thaliana [8] and cereals [5, 7] and include significant modifications. It is important to note that preparation of interphase nuclei and chromosome spreads for cytogenetic analysis typically require enzymatic digestion of the cellulose cell wall. The optimization of digestion conditions and enzymes combination is determined empirically for specific plant species and tissues, resulting in a lack of a uniform method. Consequently, a wide range of enzymes and different digestion conditions are used for plants. Here we describe a complete DNA-FISH protocol optimized for A. thaliana, which ensures the preservation of the genomic architecture of cytoplasm-free nuclei and chromosome spreads, resulting in high FISH resolution and reproducible results.

2

Materials Prepare all solutions using double-distilled water (DDW) and analytical grade reagents. Carefully follow all waste disposal regulations when disposing of waste.

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Plant Material

1. Arabidopsis thaliana seeds. As an example, here we show for wild-type (WT) col-0 and ADF8p:NTF/ACT2p:BirA transgenic plant [9]. 2. Filter paper. 3. Petri dishes.

2.2

Slide Preparation

1. Carnoy’s fixative (3:1) solution: 3 parts of ethanol absolute and 1 part of acetic acid glacial in a 50-mL centrifuge tube with screw-top cap. Prepare fresh when needed. 2. 60% acetic acid: 600 μL of glacial acetic acid and 400 μL of double-distilled water (DDW) in 1.7 mL or 2.0 mL microcentrifuge tube. Prepare fresh when needed. 3. 0.1 M citrate buffer, pH 6.0: Prepare 200 mL of DDW in a suitable glass container (conical flask). Weigh 6.07 g sodium citrate dihydrate and transfer to the water. Add 0.84 g citric acid to the solution. Mix to dissolve. Add DDW until the volume is 250 mL. Adjust solution to desired pH using 0.1 N HCl. Store in aliquots in 50-mL centrifuge tubes with screwtop caps at -20 °C. 4. 10% cellulase: 1 g cellulase in 10 mL DDW. Vortex well to completely dissolve the powder. 5. Enzyme mixture: Mix 0.5 g cellulase “Onozuka” R10 and 1.0 g pectinase in a 50-mL centrifuge tube. Add DDW until the volume is 10 mL. Vortex well to completely dissolve the powder. Add 100 μL of 10% cellulase to the enzyme mixture. Vortex well. To precipitate the foam, centrifuge briefly or store overnight at +4 °C. Store 10% cellulase and enzyme mixture in aliquots in 1.7-mL microcentrifuge tubes at -20 °C (see Note 1). 6. Excavated glass block (staining block) with concave bottom and cover glass. 7. Binocular stereomicroscope. 8. Upright light microscope (see Note 2).

2.3 DNA Probe Labeling, in Situ Hybridization

1. BAC clones (Arabidopsis Biological Resource Center (ABRC) (https://abrc.osu.edu/). 2. Salmon sperm DNA: 1500 ng/μL salmon sperm DNA dissolved in DDW. Dissolve 1 mL of 10 mg/mL stock solution (commercially available) in 5.7 mL of DDW. Store in aliquots at -20 °C. 3. SSC (×20) buffer, pH 7.0: Prepare 200 ml of DDW in a suitable glass container (conical flask). Weigh 43.83 g sodium chloride and transfer to the DDW. Add 88.2 g tri-sodium citrate dihydrate to the solution. Mix to dissolve. Add DDW until the volume is 250 mL. Adjust solution to desired pH using 0.1 N HCl. Autoclave and store at room temperature.

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4. 7.5M ammonium acetate: Dissolve 57.81 g ammonium acetate in DDW to a final volume of 100 mL (pH 5.5). Sterilize the solution by passing it through a 0.22-μm filter and store in aliquots at -20 °C. 5. TBE (×5) stock buffer: Weigh 27 g Tris base and 13.75 g boric acid and transfer to the glass beaker. Add water to a volume of 200 mL. Add 10 mL of 0.5 M EDTA. Add DDW until the volume is 500 mL. Mix to dissolve. Store at room temperature. Working solution is 0.5× TBE buffer. 6. 1% agarose gel: 0.5 g agarose in 50 mL of 0.5× TBE and 250 mL of 0.5× TBE buffer in gel tray (7 × 10 cm). 7. Alexa Fluor 488 Nick translation Labeling Kit. 8. Cy3 Nick translation Labeling Kit (see Note 3). 9. DAPI (4′,6-diamidino-2-phenylindole) stock solution: 100 mg/mL DAPI in DDW. Store at -20 °C in aliquots protected from light. 10. DAPI working solution: dilute DAPI stock solution 1:40 in DDW. Store at -20 °C in aliquots protected from light. 11. Rubber cement glue. 12. Coplin jars. 13. ThermoBrite StatSpin System (for in situ denaturation/hybridization procedures). 14. Fluorescent microscope (see Note 4).

3

Methods

3.1 Seed Germination and Fixation of Seedlings

1. Germinate seeds on moist filter paper in a Petri dish at 23–25 ° C in the dark. 2. Fix young seedlings, 9–14 days post-germination in a freshly prepared Carnoy fixative (see Note 5). The ratio of plant material and fixative solution should be at least 1:10. The microcentrifuge tubes with a volume of 1.7–2.0 mL are convenient to use for fixing 10–20 seedlings per tube. 3. The next day, replace the fixative with a fresh portion. Keep the fixed seedlings at room temperature (RT) for 3–5 days before the preparation of cytological slides. For longer storage, keep seedlings in a fixative at +4 °C or - 20 °C until use up to a year.

3.2 Tissue Processing and Cellulose Enzymatic Digestion

This part is performed under a binocular microscope. 1. Transfer a seedling to a microscope slide. Cut off the intact leaves (or root tips) with a razor blade or two sharp tweezers. Remove other non-meristematic tissues. Perform this process for each seedling separately to prevent the tissues from drying out. Collect the leaves (or root tips) of 10–15 seedlings in a

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glass staining block containing 300–500 μL DDW. Dip the leaves (or root tips) to the bottom using stainless steel needles (see Note 6). 2. Remove water by pipetting and add 300–400 μL fresh DDW. Wash tissues with DDW 2 × 5 min. Remember to ensure that the plant tissue is placed at the bottom of glass staining block and covered with liquid in all the subsequent steps. 3. Remove DDW by pipetting and incubate the tissues in 200–300 μL of 0.1M citrate buffer 2 × 5 min. 4. Remove citrate buffer by pipetting and collect leaves (or root tips) in the center of a glass staining block using stainless steel needles. Carefully remove the remaining liquid surrounding the tissues using a dry filter paper. 5. Add 150–200 μL of enzyme mixture. Submerge the leaves (or root tips) at the bottom with stainless steel needles. Do not allow tissues to float to the surface of the drop; otherwise, the enzymatic treatment will not be successful. Cover a glass staining block with a glass to prevent evaporation and carefully transfer to an incubator at 37 °C for 20 min (see Note 7). 3.3 Preparation of Cytological Slides

1. Carefully transfer the glass staining block from the incubator to a binocular. Carefully remove the enzyme mixture by pipetting without touching the soft tissues. To keep the floating leaves (or root tips) in the center of the drop, make a few gentle clockwise (or counterclockwise) circular motions with the glass staining block, but don’t let the leaves stick together. Leave the tissue in a minimal amount of liquid to avoid sticking to the glass. 2. Wash in citrate buffer for 5 min by carefully adding 200–300 μL of buffer drop by drop from the side so that the liquid slowly flows down the wall toward the center. 3. Carefully remove the buffer by pipetting and wash in DDW for 5 min twice as above, and leave the plant tissues in the water. 4. Transfer the leaves of two to three seedlings to the center of a pre-cleaned (ready-to-use; commercially available) microscope slide in a drop of 0.5 μL DDW. Spread the cells with a stainless steel needle to form a suspension. Remove the rest of non-meristematic tissues. The concentration of cells in the drop should be high (Fig. 1a). 5. Add 3.5 μL of 60% acetic acid and spread the cells with a stainless steel needle to form a fine suspension. 6. At this stage, it is recommended to monitor the quality of the cell suspension in an upright light microscope at a low (×20) magnification, with the aperture iris diaphragm closed and the condenser lowered. The cytoplasm must be transparent, and the dark-gray/black nuclei must be clearly visible.

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7. Let the drop to partially evaporate at room temperature by gently tilting the microscope slide left and right and up and down, while keeping the drop in place (see Note 8). This droplet size is suitable for subsequent in situ hybridization under an 18 × 18 mm coverslip. 8. If the periphery of the circle dries quickly, then carefully spread the drop over the entire area with a stainless steel needle, but without touching the glass. The needle should be held at the smallest angle of inclination, that is, almost parallel to the microscope slide. This procedure also avoids a high concentration of precipitating cells at the periphery of the droplet circle. 9. When the cells adhere to the slide and the drop has largely, but not completely evaporated (about 3–5 min), that is, all cells are stuck to the glass, but have not yet dried completely, add ~600 μL of fixative by pipetting around the cells, tilting the slide slightly from left to right and up and down for 10–15 s, as described above, so that the fixative solution covers the entire surface. 10. Discard the fixative solution and add another ~600 μL of fixative around the cells adhering to the glass while continuing to rotate the slide. 11. Discard the fixative and rinse the slide twice with ~600 μL of 100% ethanol for 3–5 s. 12. Discard the ethanol, place the slide vertically on the filter paper, and allow it to air dry completely. 13. To assess the quality of cell spreads, use ×20 and ×40 air objectives on an upright light microscope with a closed aperture iris diaphragm and a lowered condenser. Dark gray-black nuclei should be clearly visible, completely free of cytoplasm, and separated from each other without forming clumps (see Note 9). 14. Recommended: in the next day, for quality assessment, slides can be stained with 30–40 μL of DAPI, mounted in 10 μL of VECTASHIELD Antifade Mounting Medium and covered with 24 × 24 mm coverslip (see Subheading 3.9 for details) and examined under coverslips at low and high (×100) magnification using a fluorescent microscope (Fig. 1b, c). Then remove the coverslips and VECTASHIELD in DDW (in a Coplin jar), air dry the slides, and fix again with Carnoy’s fixative for 10–15 s. Allow slides to air dry at room temperature. The slides can be used for FISH in the next day. The slides can be stored at RT for 1–1.5 weeks. For longer storage, immerse slides in 100% ethanol and keep at -20 °C.

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3.4 DNA Probe Labelling by Nick Translation Using Alexa Fluor 488 and Cy3 NT Labeling Kits

DNA probes for BAC-FISH are prepared using nick translation kits, according to the manufacturer’s instructions (https://www. jenabioscience.com/probes-epigenetics/dna-cdna-labeling/fluo rescent-dna-cdna-labeling/fluorescent-nick-translation-labelingkits/pp-305-cy3-cy3-nt-labeling-kit) with some modifications. 1. Prepare 20 μL Nick translation labeling assay in a sterile 0.2mL PCR tube: 2 μL of 10× NT labeling buffer, 2 μL of Cy3 or AF488 labeling mix, 1 μg of BAC DNA, 1 μL of enzyme mix (instead of 2 μL recommended by manufacturer), and PCR-grade water, and fill up to 20 μL. Vortex the mix gently and centrifuge briefly to collect the reaction mixture at the bottom of the tube. 2. Place the tube in a precooled PCR thermocycler at 15 °C. Incubate for 35–40 min to generate DNA fragments in a size range between 200 and 500 bp. If the DNA fragments are too large or too small, it is necessary to increase or decrease the reaction time by 5–10 min, respectively. The optimal time is determined empirically. 2. To control the length of the DNA fragments, load 2 μL of the assay on 1% agarose gel. Place the reaction tube at -20 °C while running the gel at 80 V/53 mA for 30 min. 3. To stop the reaction, add 5 μL of stop buffer. Place the tube on ice. Proceed to purification or store at -20 °C.

3.5 Purification of the DNA Probes

1. Add to the tube: 2.4 μL of salmon sperm DNA, 25 μL of 7.5 M ammonium acetate, 150 μL of 100% ethanol (precooled at 20 °C). Mix well by pipetting and put at -80 °C for 2–3 h. 2. Spin at 14462 ×g for 30 min. Discard supernatant. 3. Add 700 μL of 70% ethanol. Spin at 14462 ×g for 5 min. Discard supernatant. 4. Air-dry the pellet for 1 h or more in the hood until the ethanol has evaporated. Keep the microcentrifuge tubes horizontal. 5. Dissolve the pellet in 20 μL of molecular biology-grade water (MW): Incubate the tube at 60 °C for 10 min in a water bath. Mix well by pipetting. Incubate again at 60 °C for 10 min. Mix well by pipetting. The precipitate must completely dissolve. Otherwise, put the tube back at 60 °C for additional 10 min and mix by pipetting until it fully dissolves. 6. Vortex briefly. Centrifuge briefly. The pellet must completely dissolve. Store at -20 °C.

3.6 PreHybridization Slide Treatments

1. The day before hybridization, dehydrate the slides in a series of increasing concentrations of ethanol. To do this, prepare 40–50 mL of 70%, 90%, and 100% ethanol in three Coplin jars. 2. Immerse slides sequentially in 70%, 90%, and 100% ethanol for 5 min each.

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3. Transfer slides from 100% ethanol to a glass Coplin jar (or any convenient glass slide holder) and place in an incubator at 37 °C overnight. 4. Next day, treat the cells with 1 μg/mL of DNase-free RNase A in 2× SSC buffer for 60 min at 37 °C in a Coplin jar. 5. Wash 3 × 3 min with 2× SSC at 37 °C (preheat 2× SSC buffer to 37 °C overnight along with slides) in a Coplin jar. 6. Cool to room temperature (approximately 15–20 min). 7. Dehydrate the slides in increasing ethanol series (70%, 90%, and 100%, 3 min each) at room temperature. 8. Immerse in 2× SSC buffer for 2 min. 9. Allow slides to air dry at room temperature for 1 h. 3.7

Hybridization

1. Prepare 20 μL of hybridization mix for each slide in 1.7 mL microcentrifuge tube: 8 μL of MW, 2 μL of 20× SSC, 4 μL of 50% dextran sulfate (cut off the end of the tip to make it easier to pipette the dextran). Vortex very well, centrifuge briefly. Add 2 μL of DNA probe F1H21 labeled with AF488 (700–800 ng) and 2 μL of DNA probe F10O11 labeled with Cy3 (700–800 ng). Vortex briefly, spin briefly, and place on ice (see Note 10). 2. Mark the center of the dry drop on the side edge of the microscope slide with a marker. Place 20 μL of hybridization mix onto slide and cover with 18 × 18 mm coverslip. Be sure that the coverslip is central, the liquid is well distributed, and there are no air bubbles. Apply rubber cement glue around the coverslip. Allow to air-dry at room temperature (approximately 15–20 min). Transfer slide to hybridization chamber in ThermoBrite StatSpin System. 3. Denature DNA probes and cells at 93 °C for 3 min and then hybridize at 63 °C for 50 min.

3.8 Posthybridization Washes

1. Remove carefully the rubber cement using tweezer. Hold the corner of the coverslip so that it does not move. 2. Remove the coverslips by immersing the slides in a Coplin jar containing 2× SSC preheated to 63 °C in water bath. 3. Transfer slides to another Coplin jar containing 2× SSC preheated to 63 °C in a water bath, wash twice for 5 min. 4. Transfer Coplin jar from the water bath to the working desk and cool to 37 °C (approximately 20 min). 5. Wash slides in 0.1× SSC (preheated to 37 °C), twice for 5 min, at 37 °C in incubator.

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6. Cool to room temperature (approximately 15 min). 7. Wash slides in DDW for 1 min. 8. Let the slides air dry in the dark (approximately 15–20 min). Shake the slides slightly several times to remove the drops more quickly. 3.9

DAPI Staining

1. Add 30–40 μL of 2.5 mg/mL DAPI on the slide and cover with 24 × 24 mm coverslip or Parafilm (cut into squares ~24 × 24 mm) and stain for 10 min in the dark at room temperature. 2. Remove the Parafilm (or coverslip) under a gentle stream of running water and wash slides in DDW for 1 min in a Coplin jar. 3. Air dry the slides in the dark (approximately 15–20 min). Shaking the slides gently several times may help to remove excess liquid and promote evaporation. 4. Place a drop of VECTASHIELD Antifade Mounting Medium (approximately 10 μL) on the slide and cover with the 24 × 24 mm (or 24 × 32 mm) coverslip. 5. To remove excess medium, use a filter paper and apply gentle pressure. Applying excessive pressure may damage the nuclei/ chromosomes. 6. Apply rubber cement around the coverslip and allow to air dry in the dark. Store the slides at +4 °C protected from light. 7. The next day, examine the slides using a fluorescent microscope. To stabilize the fluorescence before analysis, keep the slides at +4 °C at least overnight.

3.10 Microscopy and Image Analysis

1. We examined the slides on a Leica DMi8 fluorescent inverted microscope with an HC PL APO 100×/1.40–0.70 oil objective and the filter sets for DAPI, Cy3, and Alexa Fluor 488 using Leica Application Suite X (LAS X) software for image capture and adjustment. 2. Images of interphase nuclei and chromosomes were processed using the PaintShop Pro8 software.

4

Notes 1. A wide range of enzymes and different digestion conditions are used for plants. We use the following enzymes: Cellulase “Onozuka” R-10 (Yakult Honsha CO., Ltd.), Pectinase ex Aspergillus (NBS Biologicals), Cellulase ex Aspergillus niger (NBS Biologicals) [5].

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2. We use Leica DME upright light microscope. This microscope is simple and small, which makes it convenient to use on the same working desktop along with a binocular microscope. 3. We use AF488 and Cy3 Nick Translation Labeling Kits of Jena Bioscience GmbH. 4. We use Leica DMi8 fluorescent inverted microscope with Leica Application Suite X (LAS X) software. 5. To accumulate metaphase chromosomes, transfer the 9–14-day post-germination seedlings in a 1.7-mL microcentrifuge tube containing few drops of ice-cold water and place on crushed ice for 20–24 hours. Then, remove excess water by placing the seedlings onto a piece of absorbent filter paper. Then, immerse seedlings in a freshly prepared fixative 3:1 (v/v) 100% ethanolacetic acid glacial. 6. For successful washing and subsequent enzymatic treatment, the tissues should lie at the bottom of the staining block and not float on the drop surface. 7. The longer the material is stored in the fixative (several months), the faster the enzymatic digestion occurs. To avoid over-digestion, reduce the time or concentration of enzymes empirically. Conversely, if there is insufficient enzymatic digestion, the cell wall is preserved and/or the chromosomes and nuclei are in the cytoplasm, which will cause FISH to fail. 8. It is important to note that maceration in 60% acetic acid is conducted at room temperature. Maceration at 45–50 °C has been applied for Arabidopsis using other enzymatic digestion conditions [8]. In developing this protocol, we found that maceration in the hot drop of acetic acid cause chromatin destruction during denaturation at 93 °C. 9. Damaged or pale light-gray nuclei indicate enzymatic overdigestion. In this case, there is a high probability of chromatin destruction during denaturation at 93 °C causing dim DAPI and DNA-probe fluorescence after hybridization. 10. For hybridization under a 22 × 22 mm coverslip, increase the total volume of Hybridization Mix to 25 μL per slide.

Acknowledgments This protocol is adapted from the previous published paper [6] supported by the Israel Science Foundation (grant 748/14) and I-CORE Program of the Planning and Budgeting Committee and the Israel Science Foundation, grant no. 41/11 for OH.

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References 1. Schwarzacher T, Heslop-Harrison P (eds) (2000) Practical in situ hybridization. Bios Scientific Pub Ltd, Oxford 2. Celeda D, Aldinger K, Haar F-M et al (1994) Rapid fluorescence in situ hybridization with repetitive DNA probes: quantification by digital image analysis. Cytometry 17(1):13–25 3. Anamthawat-Jonsson K, Reader SM (1995) Pre-annealing of total genomic DNA probes for simultaneous genomic in situ hybridization. Genome 38(4):814–816 4. Reader SM, Abbo S, Purdie KA et al (1994) Technical tips. Trends Genet 10(8):265–266 5. Raskina O, Belyayev A, Nevo E (2004) Activity of the En/Spm-like transposons in meiosis as a base for chromosome repatterning in a small, isolated, peripheral population of Aegilops speltoides Tausch. Chromosom Res 12(2):153–161

6. Krispil R, Tannenbaum M, Sarusi-Portuguez A et al (2020) The position and complex genomic architecture of plant T-DNA insertions revealed by 4SEE. Int J Mol Sci 21(7):2373. https://doi. org/10.3390/ijms21072373 7. Shams I, Raskina O (2018) Intraspecific and intraorganismal copy number dynamics of retrotransposons and tandem repeat in Aegilops speltoides Tausch (Poaceae, Triticeae). Protoplasma 255(4):1023–1038. https://doi.org/10.1007/ s00709-018-1212-6 8. Lysak M, Fransz P, Schubert I (2006) Cytogenetic analyses of Arabidopsis. Methods Mol Biol 323:173–186 9. Deal RB, Henikoff S (2011) The INTACT method for cell type–specific gene expression and chromatin profiling in Arabidopsis thaliana. Nat Protoc 6(1):56–68. https://doi.org/10. 1038/nprot.2010.175

Chapter 18 DBD-FISH Using Specific Chromosomal Region Probes for the Study of Cervical Carcinoma Progression Catalina Garcı´a-Vielma, Elva I. Corte´s-Gutie´rrez , Jose´ L. Ferna´ndez, Martha I. Da´vila-Rodrı´guez, and Jaime Gosa´lvez Abstract Genomic instability is an important biomarker in the progression of cervical carcinoma. DBD-FISH (DNA breakage detection-fluorescence in situ hybridization) is a sensitive method that detects strand breaks, alkali-labile sites, and incomplete DNA excision repair in cells of the cervical epithelium. This technique integrates the microgel immersion of cells from a vaginal lesion scraping and the DNA unwinding treatment with the capacity of FISH integrated into digital image analysis. Cells captured within an agarose matrix are lysed and submerged in an alkaline unwinding solution that generates single-stranded DNA motifs at the ends of internal DNA strand breaks. After neutralization, the microgel is dehydrated and the cells are incubated with DNA-labeled probes. The quantity of a hybridized probe at a target sequence corresponds to the measure of the single-stranded DNA produced during the unwinding step, which is equivalent to the degree of local DNA breakage. DNA damage does not show uniformly throughout the entire DNA of a cell; rather, it is confined to specific chromosomal sites. In this chapter, an overview of the technique is supplied, focusing on its ability for assessing the association between DNA damage in specific sequences and in the progressive stages of cervical carcinoma. Key words DBD-FISH, DNA breakage detection-fluorescence in situ hybridization, DNA damage, Microgel embedding, Cervical carcinoma

1

Introduction Genomic instability is defined as the increase in genetic alterations that cause heterogeneity in the cell and that comprises a particular feature prior to the development of cancer [1]. Cervico-uterine cancer (CaCU) ranks fourth at the worldwide level among the most frequent cancers in women and is associated with human papilloma virus (HVP) infection [2]. Once the HPV is introduced into the cell, it gives rise to instability in the cell, manifesting itself in alterations at the molecular and chromosomal level. Genomic instability is an important biomarker in the progression of low-grade squamous intraepithelial lesion (LG-SIL) and high-grade

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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squamous intraepithelial lesion (HG-SIL) in CaCU [3, 4]. These alterations appear in specific sites of the chromosomes, where the genes that predispose to CaCU are located [5, 6]. Tandemly repeated DNA makes up a significant part of the human genome. Satellite DNA, which consists of very large arrays of tandemly repeated, noncoding DNA, is the principal element of functional centromeres and forms the main structural essential of heterochromatin [7]. Alpha satellite DNA is formed of monomers of 171 bp in length that are assembled in higher-order repeated units and are located in the centromeric and telomeric regions of all chromosomes. Satellite II and III families include microsatellites of 5 bp derived from variants of the core sequence 5’-TTCCA-3′ and contain sequences based on tandem repetitions. These sequences are rich in the juxtacentromeric heterochromatin of human chromosomes 1, 9, 16, and in the long arm of the Y chromosome [8]. The tandem organization of these sequences produces special functional characteristics in centromeric DNA organization, replication, and response to stress as cancer [9]. Chromosome-1 plays an important role in the development of genomic instability in CaCU, for example, the aneusomy of this chromosome is necessarily associated with high-risk HPV infection in HG-SIL [4, 10], although structural alterations have also been observed [11–13], frequently involving the 1p36 region [14]. Changes in other chromosomes, such as the gain of genetic material in 3q [15] and the loss of 11q, have been reported as early events in progression to CaCU [16]. Genomic instability can be analyzed with the use of molecular cytogenetic methodologies, such as the micronucleus technique, sister chromatid exchange, and the comet assay [17–20]. The DBD-FISH is a procedure that allows cell-by-cell detection and the quantification of DNA breakage in the whole genome or within specific DNA sequences using whole genome or fluorescence-labeled specific chromosomal regions [21]. DBD-FISH permits the detection of simple DNA breaks at the alkaline-labile sites of individual cells; at specific chromosomal sites, DNA damage can be quantified using image analysis software, such as ImageJ [22]. The results are interpreted by the amount of fluorescence generated, that is, a cell with more damage will generate more breaks in the DNA. Thus, a larger amount of probe will hybridize, generating a greater fluorescence signal that will be expressed in numerical values [4]. This chapter proposes the application of the DBD-FISH technique using DNA probes from specific chromosomal regions (satellite DNA alphoid, 5 bp classical satellite III, telomeric regions, and unique sequences (1p36 and 3q26)), as well as support of the monitoring of these chromosomal biomarkers in the study of the malignant progression of cervical carcinoma.

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273

Materials

2.1 Materials and Reagents (See Note 1)

1. Agarose is used to prepare the slides (see Subheading 3.1, step 1 Slide preparation section). 2. 0.4% Trypan Blue (see Note 2). 3. Buffer to calibrate potentiometer. 4. Horizontal staining dishes (glass). 5. Cytological brushes (cytobrushes). 6. 22 mm × 22 mm glass coverslips. 7. Plastic coverslips (parafilm paper). 8. DAPI (4′,6-diamidino-2-phenylindole) II counterstain. 9. Diamond Tip Pencil. 10. 1 N NaOH. 11. Phosphate buffered saline (PBS) 1X. 12. Tweezers. 13. Plastic Pasteur pipettes. 14. Slides. 15. Graduated cylinder. 16. Silicone or rubber adhesive. 17. Dako Total Telomere PNA Probe. 18. Centromeric probes: Chromosome 1 (CEP 1-D1Z1) satellite III, Chromosome 3 (CEP 3-D3Z1), and Chromosome 8 (CEP 8-D8Z2), Vysis, Abbott Molecular. 19. Specific probes for chromosomal regions 1p36 and 3q26, Vysis, Abbott Molecular.

2.2

Solutions

1. 20X SSC buffer: 3.0 M NaCl and 0.3 M Trisodium Citrate Buffer at pH 5.3. 2. 0.4X SSC/0.3% NP-40: Mix 2 ml of 20X SSC and 0.3 mL of NP-40 to make up to 100 mL with double-distilled water and adjust to pH 7–7.5. 3. 1% agarose: Dissolve 1 g of low-melting-point agarose in 100 mL of water. Apply heat to melt, and add 50-μL aliquot into microtubes. 4. 2X SSC wash buffer: Dilute 20X SCC 1:10 in double-distilled water. 5. Hybridization wash buffer: 50% formamide in 2X SSC (v/v). 6. 0.5 M EDTA: Dissolve 18.6 gr of EDTA in 100 mL of doubledistilled water and adjust to pH 7.5.

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7. 70%, 80%, and 96% ethanol: Place 70, 80, and 96 mL of ethanol, respectively, into a volumetric flask and make up to 100 ml with double-distilled water. 8. Cell lysis buffer: 2 M NaCl, 0.05 M EDTA, 0.4 M Tris Base, and 1% SDS. Adjust to pH 7.5. Store at 4 °C. 9. 0.9% NaCl: Dissolve 0.9 gr of NaCl in 100 mL of doubledistilled water. 10. Strong alkaline solution: 0.03 M NaOH and 1 M NaCl. Adjust to pH 12.5. Prepare before use and store at 4 °C until use. 11. Neutralizing solution: 0.4 M Tris-HCl, pH 7.5 in doubledistilled water to 100 mL and store at 4 °C. 12. 1X TBE: 0.09 M Tris-borate-EDTA buffer and 2 mM Trisborate. 13. Propidium iodide 1:100 in antifade mounting medium. 14. 0.4% NP40/2X SSC: Mix 2 mL of 20X SSC, 0.3 mL of NP-40. Adjust to pH 7–7.5 and dilute to 100 mL with distilled water. 15. 2X SSC/0.1% NP-40: Mix 10 mL of 20X SSC with 0.1 mL of NP-40. Adjust to pH 7–7.5. and dilute to 100 mL with distilled water. 16. Slide degreasing solution: Mix 210 mL of 96% ethanol, 110 mL of methanol, and 150 mL of acetic acid. Prepare when it is going to be used. 2.3 Laboratory Equipment and Instruments

1. Hybridizer or humid chamber. 2. Constant temperature oven (45–50 °C). 3. Bright-field microscope and phase contrast. 4. Fluorescence microscope (see Note 3).

3

Method

3.1 Preparation of Agarose Slides

1. Dissolve 0.65 g of low-melting-point agarose in 100 mL of distilled water. 2. Mix at room temperature for 10 min. 3. Apply heat and continue stirring until dissolved. 4. Pour the agarose into a vertical flask in a water bath at 80 °C. 5. Clean the slides with slide defatting solution and label them with a diamond pencil, marking the side of the slide where the agarose layer will remain. 6. One by one, slowly dip the slides into the jar with the hot agarose. 7. Remove the slide and clean the back with gauze.

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8. Place the slides in a metal tray previously cold at 4 °C. 9. Incubate at 4 °C for 30 min. 10. Store at room temperature in boxes protected from dust and light. 3.2

Sampling

1. Use a soft-bristled brush (cytobrush) to perform moderate-tointense scraping of the lesion area (see Note 4). 2. The brush should be rotated 360° repeatedly to collect the largest number of cells and to ensure that the sample derives from the endocervix. 3. The brush with the sample is placed in a sterile 15-mL tube with 1X PBS, duly labeled, and transported at room temperature until processing. 4. No more than 1 h should elapse between sample collection and processing.

3.3

Viability

1. Centrifuge the tube with the cytobrush with the patient’s sample at 1000 ×g for 10 min. Remove the cytobrush and centrifuge again under the same conditions. Decant the supernatant to 1 mL and resuspend the cell button. 2. Take 10 μL of the cell button and place it in a micro-tube. 3. Add Trypan Blue to the sample tube to a final concentration of 0.04%. 4. Mix slowly with a Pasteur pipet and incubate at room temperature for 30 min. 5. Place an aliquot of the mixture on a glass slide, cover with a coverslip, and observe under a bright-field microscope. Dead cells will stain blue and live cells will appear transparent. 6. Determine percentage of live cells. 7. An optimal sample for analysis is >85% live cells (see Note 5).

3.4 DBD-FISH in Cervical-Scraping Cells

1. Prepare previously microtubes with 50 microliters of 1% lowmelting-point agarose and leave at 37 °C until use. 2. Take 30 μL of the epithelial cell button and dilute it in a microtube with 50 μL of 1% low-melting-point agarose. Mix well. 3. Place approximately 30 μL of the mixture on glass slides previously prepared with a layer of 0.65% agarose. Embed in two wells along the slide. 4. Cover with 22 × 22 mm glass coverslips (Fig. 1). 5. Incubate at 4 °C for 15 min on a pre-chilled aluminum tray. 6. Carefully remove the coverslip by sliding it to one side of the slide.

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Fig. 1 Preparation of slides for DBD-FISH. (1 and 2) Glass slides prepared with a layer of 0.65% agarose. (3) Sample; 30 μL of the epithelial cell +50 μL of 1% low-melting-point agarose. (4) Glass coverslips

7. Immerse the slides in horizontal staining boxes, at room temperature, containing cell lysis buffer. Incubate for 30 min. 8. Replace the cell lysis buffer with 1X TBE buffer and incubate for 10 min. 9. Wash with 0.9% NaCl for 2 min. 10. Incubate with strong alkaline for 2.5 min. 11. Incubate with neutralizing solution for 5 min. 12. Incubate with 1X TBE buffer for 2 min. 13. Incubate with 70%, 90%, and 100% ethanol for 2 min each. 14. Let the slides dry, protected from dust and light, overnight. 15. Check the slide to verify the formation of the DBD halo (see Note 6) (Fig. 2). 3.5

Hybridization

The conditions for hybridization can slightly vary according to the type of probe. The specific conditions of hybridization of satellite DNA alphoid (CEP 3 and CEP 8) 5 bp classical satellite III of chromosome-1 (see Note 7) and unique sequence (1p36 and 3q26) (see Notes 8–9) are presented in the Table 1. Generally, perform the following steps:

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Fig. 2 Chromatin dispersion halo produced after treatments with lysis solution (bottom panel); failed (a) and successful halo (b) for DBD-FISH. Representative scheme (up panel)

1. Prepare a mix of probes and hybridization buffer as indicated in Table 1. 2. Denature probes at the indicated temperature for 5 min. Apply 7–10 μL of probe mix on each sample and incubate at the indicated temperature overnight or for the indicated time as listed in Table 1. Keep samples in the dark. 3.6 Posthybridization Washes

Post-hybridization washing after FISH hybridization is necessary to aid the removal of nonspecific interactions between the probe and undesirable regions of the genome, thus allowing greater probe specificity. Washes are performed in horizontal staining boxes at room temperature.

3.6.1 Regular Washing

1. Formamide 50%/2X SSC for 10 min (perform this step three times with fresh solution each time). 2. 2X SSC for 10 min. 3. 0.4% NP40/2X SSC for 5 min. 4. Allow to dry (see Note 10).

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Table 1 Differences in the hybridization procedure of specific chromosome probes in DBD-FISH

Probe

Probe mark Preparation ®

Denaturation probe

Temperature and hybridization time

CEP 3 (D3Z1)

Vysis Abbott molecular

7 μl hybridization buffer, 1 μL 73 ± 1 °C/5 min of each probe and complete to 10 μL with ultra-pure water

42 °C/ overnight

CEP 8 (D8Z2)

Vysis® Abbott molecular

7 μL direct probe

The probe is already denatured; it is applied directly on the sample

42 °C/ overnight

Vysis® 5-bp classical satellite III of Abbott chromosomemolecular 1 (D1Z1)

7 μL direct probe

The probe is already denatured; it is applied directly on the sample

42 °C/ overnight

Unique sequence 1p36

Vysis® Abbott molecular

7 μL hybridization buffer, 1 μL 73 ± 1 °C/5 min of each probe and complete to 10 μL with ultra-pure water

37 °C/ overnight

Unique sequence 3q26

Vysis® Abbott molecular

7 μL hybridization buffer, 1 μL 73 ± 1 °C/5 min of each probe and complete to 10 μL with ultra-pure water

37 °C/ overnight

Total telomere PNA probe, human Pantelomeric

DAKO®

7 μL direct probe

37 °C/ 30 min

3.6.2

Quick Wash

80 °C/5 min

1. 0.4 X SSC/0.3% NP-40 at 73 °C for 1–3 s. 2. 2X SSC/0.1% NP-40 at room temperature for 1–3 s (see Note 11). 3. Let dry. 4. After hybridization, carefully remove the coverslip and perform post-hybridization washes, add 10 μL of DAPI antifade medium, and place the solution on the glass coverslips, incubate at room temperature for 15 min in the dark, and observe under a fluorescence microscope with specific filters for the fluorochrome with which the probe is labeled.

3.7 Fluorescence Microscope Analysis

1. Observe the hybridized slides in a fluorescence microscope with the specific filter for the fluorochrome with which the probe is labeled.

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2. Analyze a minimum of 50 cells per patient (see Note 12). 3. Take photographs for further analysis with image analysis software. 3.8

Image Analysis

The photographs are analyzed with ImageJ (image analysis software) as follows (19): 1. Download the Image J program, at https://imagej.nih.gov/ij/ download.html. 2. When opening the program, the toolbar is displayed (Fig. 3, Step 1). 3. Select “File” and then “Open” to open the photo you want to analyze. 4. Select the “Oval” tool to outline the fluorescence signal in the cell. Select “Analyze” and then “Set measurements” and select the following measurements: Area, Integrated Density, Standard Deviation, and Mean Gray Value. 5. Subsequently, select “Image,” “Adjust,” and “Threshold” to adjust the upper and lower limit at which the measurements will be made. The area delimited with the yellow oval is “filled in” simulating the amount and signal of fluorescence observed. All images are analyzed under the same measurement threshold (Fig. 3, Step 2). 6. Finally, select “Analyze” and then “Measure.” The values of the selected measurements of each cell and each selected area of fluorescence are displayed (Fig. 3, Steps 3 and 4). The software calculates the integrated density (ID) of each sample, which is the signal intensity multiplied by the signal area and expresses this in numerical values (Table 2). Each line represents a measurement on a cell. Analyze 50 cells and obtain average value per patient.

3.9 Statistical Analysis

1. The numerical values obtained from the software are analyzed by means of the Kruskal–Wallis test to investigate any differences in ID between the different specific DNA sequences analyzed (see Note 13). 2. A value of P < 0.05 is considered significant. With regard to specific instability, a significant increase in the hybridization signal of satellite DNA alphoid (CEP 3), 5 bp classical satellite DNA sequences from chromosome-1, unique sequences (1p36 and 3q26), and telomeric regions according to neoplastic development is observed. CEP-8 was used as control (Fig. 4 and Table 2).

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Fig. 3 Analysis of microphotographs by Digital Image Analysis using Image J software (19) Table 2 Comparison of integrity density (ID) in satellite DNA alphoid (CEP 3 and CEP 8), 5 bp classical satellite of chromosome-1, and unique sequence (1p36 and 3q26) in cervical epithelium of control women, women with LG-SIL or with HG-SIL Integrity density (ID) (pixels) (median) Control

LG-SIL

HG-SIL

CEP 3

4.1 × 103 c

4.6 × 103 c

11.2 × 103 a,b

CEP 8

0.3 × 10 3

0.3 × 10 3

0.2 × 10 3

16.5 × 103 b,c

30.0 × 103 a, c

87.0 × 103 a,b

1p36

4.8 × 103 c

2.7 × 103 c

11.2 × 103 a,b

3q26

2.4 × 103 b.c

7.2 × 103 a

7.1 × 103 b

Probes (chromosomal region) Satellite DNA alphoid

5 bp classical satellite D1Z1 (1q12 satellite III) Unique sequence

a

Differ from control; bDiffer from LG-SIL; cDiffer from HG-SIL, P < 0.05

4 Notes 1. Prepare all reagents with ultra-pure or double-distilled water. The reagents should be molecular biology grade. 2. It is recommended to add 0.01 mL of 0.4% Trypan Blue stock solution to 0.1 mL of cells in order to achieve a final concentration of 0.04%.

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Fig. 4 Cervical epithelial cells after DBD-FISH using specific probes for the centromeric region of chromosome-3 (a) and chromosome-8 (c), the 5 bp classical satellite III of chromosome 1 (D1Z1) (b), the unique sequences: 1p36 (d) and 3q26 (e), and the telomeric regions (f). Probes were labeled with red (a–c and f), green (e), and red/green (d) fluorophores. Nucleus was contrasted with DAPI (blue)

3. It is recommended to use a fluorescence microscope that is equipped with the main filters, such as DAPI (detects DNA, blue 358–461 nm), FITC (probes marked in green, 320–492 nm), and Texas Red (probes marked in red, 561–594 nm). The use of 63X and 100X “Plan Neo-fluor” objectives and specific fluorescence immersion oil is essential. To take photographs, the microscope must be connected to a

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camera with good resolution (Axiocam CCD camera, Zeiss®, Carl Zeiss, Germany) in 16-bit black and white in JPEG format in order to distinguish small fluorescence dots and software (Zen Zeiss®) appropriate for analysis. Another important point is the lighting source; currently, there are LED sources that have greater power and the hours do not require counting, as with the mercury lamp. Regular cleaning of the microscope and periodic review by a specialized technician are recommended. 4. The taking of a sample of cells from vaginal scrapings must be conducted by a duly trained health professional. The sample taking will be performed specifically on the area of the lesion, located with the aid of a colposcope. 5. The determination of viability in the sample is an important quality control, which indicates the percentage of live and dead cells present and that ensures that the DNA damage detected is due to an intraepithelial lesion or to CaCU and that it is not due to the majority of the cells are dead; the latter must be conducted with each sample to be studied. The Trypan Blue technique is a relatively easy methodology that may be conducted in any laboratory and is inexpensive and easy to interpret. 6. To verify that the DBD process was successful, it is suggested to stain one of the slide wells with 7 μL of propidium iodide 1:100 in antifade medium, incubate this in the dark for 15 min, and visualize this with a fluorescence microscope equipped with a Texas Red filter. A halo of small DNA fragments should be observed around the cell nucleus. 7. The procedure for denaturation and hybridization of the probes, that is, centromeric chromosome 1 CEP 1 (D1Z1) and centromeric chromosome 3 CEP 3 (D3Z1), is the same. If the probes are marked with different colors (e.g., green and red), both can be studied in a single hybridization experiment, by mixing 1 μL of each probe, 7 μL of hybridization buffer, and making up to 10 μL with ultra-pure water. DBD-FISH allows the use of two or more probes labeled with different fluorochromes at the same time (Fig. 4d). 8. The specific 1p36 probe (marked in red) includes an internal control that hybridizes to 1ptel (marked in green), for which two signals (red and green) will be observed, each using the specific filter (Texas Red and FITC) for each fluorochrome under the fluorescence microscope. 9. Commercial alpha-satellite probes do not include an internal control. That same slide can be stained with cytogenetic banding techniques to verify the nature of the chromosome in which hybridization has occurred.

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10. Regular washing is recommended, since fast washing sometimes leaves a fluorescence background noise. 11. Move the slide slowly for a few seconds within the solution to eliminate background noise. 12. The cell count must be conducted by two or more technicians at the same time. 13. It is recommended to determine whole-genome DNA hybridization as reference [23]. References 1. George N, Bhandari P, Shruptha P et al (2023) Multidimensional outlook on the pathophysiology of cervical cancer invasion and metastasis. Mol Cell Biochem 11:1–26 2. Sung H, Ferlay J, Siegel RL et al (2021) Global cancer statistics 2020: GLOBOCAN estimates of incidence and mortality worldwide for 36 cancers in 185 countries. CA Cancer J Clin 71(3):209–249 ˜ ez M, Arts T et al (1996) Use 3. Kurtycz D, Nun of fluorescent in situ hybridization to detect aneuploidy in cervical dysplasia. Diagn Cytopathol 15:46–51 4. Corte´s-Gutie´rrez EI, Da´vila-Rodrı´guez MI, Muraira-Rodrı´guez M et al (2005) Association between the stages of cervical cancer and chromosome 1 aneusomy. Cancer Genet Cytogenet 159(1):44–47 5. Friedberg EC (1996) Relationships between DNA repair and transcription. Annu Rev Biochem 65:15–42 6. Lee M, Nam ES, Jung SH et al (2014) 1p36.22 region containing PGD gene is frequently gained in human cervical cancer. J Obstet Gynaecol Res 40(2):545–553 7. Tyler-Smith C, Willard HF (1993) Mammalian chromosome structure. Curr Opin Genet Dev 3(3):390–397 8. Jeanpierre M (1994) Human satellite 2 and 3. Ann Ge´ne´t 37(4):163–171 9. Aze A, Sannino V, Soffientini P et al (2016) Centromeric DNA replication reconstitution reveals DNA loops and ATR checkpoint suppression. Nat Cell Biol 18(6):8684–8691 10. Bulten J, Melchers WJG, Kooy-Smits MM et al (2002) Numerical aberrations of chromosome 1 in cervical intraepithelial are strongly associated with infection hig-risk papillomavirus types. J Pathol 198:300–309 11. Atkin NB (1997) Cytogenetics of carcinoma of the cervix uteri: a review. Cancer Genet Cytogenet 95(1):33–39

12. Cottage ADS, Roberts I, Pett M et al (2001) Early genetic events in HPV immortalised keratinocytes. Genes Chrom Cancer 30:72–79 13. Wilting SM, Steenbergen RDM, Tijssen M, van Wieringen WN et al (2009) Chromosomal signatures of a subset of high-grade premalignant cervical lesions closely resemble invasive carcinomas. Cancer Res 15:647–655 14. Corte´s-Gutie´rrez EI, Garcı´a-Vielma C, Da´vilaRodrı´guez MI et al (2020) 1p36 is a chromosomal site of genomic instability in cervical intraepithelial neoplasia. Biotech Histochem 95(2):137–144 15. Verlaat W, Snijders PJF, Novianti PW et al (2017) Genome-wide DNA methylation profiling reveals methylation markers associated with 3q gain for detection of cervical precancer and cancer. Clin Cancer Res 23(14):3813–3822 16. Thomas LK, Bermejo JL, Vinokurova S et al (2014) Chromosomal gains and losses in human papillomavirus-associated neoplasia of the lower genital tract—a systematic review and meta-analysis. Eur J Cancer 50:85–98 17. Udumudi A, Jaiswal M, Rajeswari N et al (1998) Risk assessment in cervical dysplasia patients by single cell gel electrophoresis assay: a study of DNA damage and repair. Mutat Res 412:195–205 18. Corte´s-Gutie´rrez EI, Cerda-Flores RM, LealGarza CH (2000) Sister chromatid exchanges in peripheral lymphocytes from women with carcinoma of the uterine cervix. Cancer Genet Cytogenet 122(2):121–123 19. Leal-Garza CH, Cerda-Flores RM, LealElizondo E et al (2002) Micronuclei in cervical smears and peripheral blood lymphocytes from women with and without cervical uterine cancer. Mutat Res 515(1–2):57–62 20. Corte´s-Gutie´rrez EI, Da´vila-Rodrı´guez MI, Zamudio-Gonza´lez EA et al (2010) DNA damage in Mexican women with cervical

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Catalina Garcı´a-Vielma et al.

dysplasia evaluated by comet assay. Anal Quant Cytol Histol 32:207–213 21. Corte´s-Gutie´rrez EI, Da´vila-Rodrı´guez MI, Ferna´ndez JL et al (2011) DNA damage in women with cervical neoplasia evaluated by DNA breakage detection-fluorescence in situ hybridization. Anal Quant Cytol Histol 33: 175–118

22. Schneider CA, Rasband WS, Eliceiri KW (2012) NIH image to ImageJ: 25 years of image analysis. Nat Methods 9:671–675 23. Corte´s-Gutie´rrez EI, Ortı´z-Herna´ndez BL, Da´vila-Rodrı´guez MI et al (2013) 5-bp classical satellite DNA loci from chromosome-1 instability in cervical neoplasia detected by DNA breakage detection/fluorescence in situ hybridization (DBD-FISH). Int J Mol Sci 14: 4135–4147

Chapter 19 CRISPR-Based Split Luciferase as a Biosensor for Unique DNA Sequences In Situ Nicholas G. Heath and David J. Segal Abstract To date, CRISPR-based DNA targeting approaches have typically used fusion proteins between full fluorescent reporters and catalytically inactive Cas9 (dCas9) for imaging rather than detection of endogenous genomic DNA sequences. A promising alternative strategy for DNA targeting is the direct biosensing of user-defined sequences at single copy with single-cell resolution. Our recently described DNA biosensing approach using a dual fusion protein biosensor comprised of two independently optimized fragments of NanoLuc luciferase (NLuc) directionally fused to dCas9 paired with user-defined single-guide RNAs (sgRNAs) could allow users to sensitively detect unique copies of a target sequence in individual living cells using common laboratory equipment such as a microscope or a luminescence-equipped microplate reader. Here we describe a protocol for using such a DNA biosensor noninvasively in situ. Key words Microscopy, DNA biosensor, DNA detection, DNA imaging, Luminescence imaging, Fluorescence imaging, Live cell imaging, In situ imaging, Plate reader assay

1

Introduction The CRISPR/Cas gene editing system has recently been modified for imaging endogenous genomic loci, but the vast majority of current approaches employ fluorescent reporter-based probes, such as dCas9-GFP [1–8]. However, with such “always on” probes, signal is produced in both bound and unbound states, resulting in a high fluorescent background that negatively impacts the signal-tobackground ratio (SBR) [9]. Therefore, such probes require a high local concentration of binding events to distinguish signal from background, so they are mostly limited to imaging highly repetitive elements that can be targeted by one sgRNA or to unique sequences targeted by 20–30 or more sgRNAs [1, 3]. Imaging a short sequence present at a single copy has remained challenging. In addition to the major issue of accounting for higher background from unbound fluorescent probes leading to a naturally

Gal Haimovich (ed.), Fluorescence In Situ Hybridization (FISH): Methods and Protocols, Methods in Molecular Biology, vol. 2784, https://doi.org/10.1007/978-1-0716-3766-1_19, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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higher cellular auto-fluorescent background signal, fluorescencebased biosensing is plagued by other issues such as cellular phototoxicity and photobleaching of fluorophores and fluorescent proteins [10–12]. All of these issues contribute to a cumulative negative effect on SBR achievable with fluorescent probes. To increase sensitivity, luminescent reporters could offer an attractive alternative to fluorescent reporters in biosensor design as they have negligible auto-luminescent background signal. This is mainly because luminescence represents light produced from a catalytic reaction of an enzyme with its substrate as opposed to light produced from excitation by incident exogenous light [11, 12]. Despite luminescent reporters having the advantage of decreased background, the raw signal outputted from fluorescent reporters is brighter than available luminescent reporters [12]. However, a relatively new luciferase, NanoLuc, bridges this gap in signal intensity [13, 14]. NanoLuc luciferase offers several advantages over Firefly and Renilla luciferases including enhanced stability, significantly smaller size, and >150-fold enhancement in luminescence output [13, 14]. In addition, furimazine, the substrate for NanoLuc luciferase, is more stable and exhibits decreased levels of background activity than the substrate for Renilla luciferase, coelenterazine [13, 14]. Bipartite “turn on” DNA biosensors offer an exciting alternative strategy for targeting specific DNA sequences in live cells due to their higher sensitivity and specificity for a given target sequence compared to “always on” full fluorescent reporter-dCas9 fusion probes. Generally, on-target signal is higher upon binding to a single target sequence, while off-target or background signal is lower in the unbound state for such bipartite “turn on” probes, making them more advantageous than full reporter-based fusion probes for high sensitivity DNA biosensing applications. “Turn on” approaches to DNA biosensing using light-producing enzymes include activation of a chromophore by energy transfer from another activated chromophore or reassembly of a bright reporter molecule. However, recent “turn on” DNA biosensing approaches based on Fo¨rster resonance energy transfer (FRET) [15–19] have required more than three unique sgRNAs to detect individual endogenous genomic loci, while split reporter DNA or RNA biosensing has been described primarily by previous studies in vitro [20–24] with several studies describing using transcription activator-like effectors (TALEs) as DNA binding domains and split fluorescent proteins for DNA biosensing in live cells [25, 26]. To harness both the advantages of luminescent reporters and “turn on” biosensor design strategies, we recently developed a split luciferase DNA biosensor [9] based on the NanoLuc Binary Technology (NanoBiT) complementation reporter system created for NLuc [27] and catalytically inactive Cas9 (dCas9) from Streptococcus pyogenes. Applying this “turn on” DNA detection tool in live

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cells, we demonstrated a maximum of 11.5-fold signal-to-background ratio targeting various nonrepetitive endogenous genomic loci, including single nucleotide polymorphisms (SNPs). This split luciferase biosensor should be a broadly useful platform for many live cell DNA biosensing applications that require low copy number resolution and minimal destruction of highly valuable cell populations, including real-time genotyping of heterozygotes and homozygotes at a defined locus or even real-time detection of chromosomal rearrangements at specific junction points in situ. Here we describe a complete protocol to use our DNA biosensor to target unique copies of an endogenous sequence in individual living cells using transient transfection methods and common laboratory equipment such as a microscope or a luminescenceequipped microplate reader for signal readout (Fig. 1).

Fig. 1 Depiction of experimental process for split NLuc DNA biosensing assays with plate reader (left) and microscope (right) signal readouts using a LgBiTdSpCas9 + dSpCas9-SmBiT DNA biosensor

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Materials For all procedures prior to transfection, materials do not need to be prepared in a sterile working environment. Prepare all materials for transfection and signal measurement in a sterile biosafety cabinet.

2.1

Reagents

1. Q5 High-Fidelity 2X Master Mix (or equivalent high-fidelity polymerase master mix). 2. PCR purification equivalent).

kit

(QIAGEN,

Zymo

Research,

or

3. Gibson Assembly Master Mix. 4. Mammalian cell line(s) of choice (e.g., HEK293T or similar cell type amenable to transient transfection; see Note 1). 5. Complete growth media: DMEM, 10% (v/v) fetal bovine serum (FBS), 1% penicillin/streptomycin. Adjust culture media to cell line. 6. Opti-MEM® I Reduced Serum Medium. 7. 24-well and 96-well tissue culture plates. 8. 96-well opaque microplates.

white-side

translucent

bottom

assay

9. Lipofectamine 3000 or equivalent cationic liposome-based transfection system. 10. Nano-Glo Live Cell Assay. 11. Microscope slides and cover slips. 12. pUC19 plasmid (Addgene #50005). 13. LgBiT-dSpCas9, dSpCas9-SmBiT, and NLuc-dSpCas9 plasmid constructs (Addgene; see Fig. 1 and Note 2). 14. dSpCas9 sgRNA backbone plasmid (Addgene #41824). 15. Custom sgRNA oligos for cloning (see Ref. [28]). 16. AflII restriction enzyme. 2.2

Equipment

1. Thermocycler. 2. Electroporation/nucleofection apparatus. 3. Epifluorescence microscope: Leica DM6000B or equivalent. The microscope should be equipped with at least sCMOS camera but preferably equipped with EMCCD camera. 4. Microplate reader device. Must be equipped with luminescence detection capabilities.

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289

Methods Conduct all procedures at room temperature unless otherwise specified.

3.1

sgRNA Design

1. Linearize the dSpCas9 sgRNA expression vector backbone using the AflII restriction enzyme. 2. Each desired 19-bp sgRNA target sequence can be incorporated into two 60mer oligonucleotides that contain a 20-bp homologous region representing the desired sgRNA target sequence with 5′ position in the homologous region replaced by guanine [28]. These oligonucleotides also contain 40-bp homologous sequences to the sgRNA expression vector for subsequent molecular cloning via Gibson assembly or an analogous method (see Note 3). 3. Anneal and extend the oligonucleotides using three to four cycles in a thermocycler with Q5 high-fidelity polymerase (or equivalent) according to the following reaction setup and thermocycling parameters. Reaction setup. Q5 High-Fidelity 2X Master Mix

12.5 μL

10 μM forward sgRNA oligo

1.25 μL

10 μM reverse sgRNA oligo

1.25 μL

Nuclease-free H2O

10 μL

Thermocycling parameters. Initial denaturation

98 °C for 3 min

Cycle denaturation

98 °C for 10 s, 3–4 cycles

Cycle annealing

72 °C for 30 s, 3–4 cycles

Cycle extension

72 °C for 4 s, 3–4 cycles

Final extension

72 °C for 2 min

4. After oligonucleotide annealing and extension, use a PCR purification kit to purify the 100 bp dsDNA. Then, insert the purified dsDNA containing the desired 19-bp sgRNA target

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sequence into the AflII linearized sgRNA expression vector using a Gibson assembly reaction with the following setup (see Note 4).

3.2 Biosensor Transfection Setup for Microplate Reader Measurement

AflII-linearized sgRNA expression vector

100 ng (Variable—example amount 0.5 μL)

Annealed & extended sgRNA oligos

~20 ng (variable—Example amount 4 μL)

2X Gibson assembly master mix

10 μL

Nuclease-free H2O

Variable—Example amount 5.5 μL)

1. Seed 10,000 to 25,000 low-passage cells per well in 96-well opaque white-side translucent bottom microplates approximately 12–20 h before transfection (see Note 5). 2. Transiently transfect cells with 100 ng total DNA per well according to the instructions in the next step. Use either lipofectamine 3000 or equivalent cationic liposome-based transient transfection system or electroporation/nucleofection (see Note 6) (Fig. 2). 3. Set up two co-transfections for 0.1 fmol biosensor plasmid and 1 fmol biosensor plasmid (see Note 7). For 1 fmol biosensor plasmid, prepare 6.4 ng LgBiT-dSpCas9 plasmid, 6.12 ng dSpCas9-SmBiT plasmid, 2.44 ng of each sgRNA target plasmid, and 82.6 ng pMAX-GFP plasmid for each well. For 0.1 fmol biosensor plasmid, prepare 0.64 ng LgBiT-dSpCas9 plasmid, 0.62 ng dSpCas9-SmBiT plasmid, 0.25 ng of each sgRNA target plasmid, 82.6 ng pMAX-GFP plasmid, and 15.64 ng pUC19 plasmid for each well (see Notes 8 and 9). 4. Alongside on-target conditions where biosensor plasmids and sgRNA plasmids are transfected, we typically include several controls such as an equimolar transfection of NLuc-dSpCas9 plasmid and each of the split reporter fusion constructs individually (LgBiT-dSpCas9 alone and dSpCas9-SmBiT alone) to establish the dynamic range of the assay and an off-target condition to establish background signal which can be compared to on-target conditions to calculate signal-to-background ratios for the assay (see Note 10). 5. 24 hours post-transfection, check cells for GFP expression. If cells are not positive for green fluorescence or show lower transfection efficiency than expected, repeat steps 2–5. 6. Incubate cells for 24–72 h at 37 °C with 5% CO2 (see Note 11). 7. To measure fluorescent signals, use a microplate reader equipped to measure fluorescence. We typically use a high photomultiplier tube (PMT) sensitivity setting and at least 100 reads per well before taking any luminescent readings.

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Fig. 2 Image Postprocessing and Signal Analysis (a) Example image of LgBiT-dSpCas9 + dSpCas9-SmBiT + sgRNAs delivered to cells on a chamber slide and imaged at higher magnification (63X oil immersion) on a spinning disc confocal microscope equipped with an EMCCD camera. GFP fluorescence is shown in green and NanoLuc luminescence is shown in red. (b) Example image of LgBiT-dSpCas9 + dSpCas9-SmBiT + sgRNAs delivered to cells on a chamber slide and imaged at lower magnification (25X) on a STORM super-resolution microscope equipped with a sCMOS camera. GFP fluorescence is shown in green and NanoLuc luminescence is shown in magenta. (c) Imaging artifact (possibly an interference pattern) caused by external light sources present in the microscope room. (d) Resulting 8-bit segmented image from Trainable Weka Segmentation pipeline after several rounds of training. (e) 8-bit segmented image after thresholding and binarization. (f) Magnified image showing two examples of tight circled regions of interest (ROIs) in a noisy background. Numbers should be assigned by the Weka Segmentation algorithm to the ROIs corresponding to a specific group of segmented cells. This is the result that should be outputted from an appropriately trained Weka Segmentation algorithm

8. Equilibrate Nano-Glo® LCS Dilution Buffer to ambient temperature if using for the first time. Remove the Nano-Glo® Live Cell Substrate from storage, and mix. If the Nano-Glo® Live Cell Substrate has collected in the cap or on the sides of the tube, briefly spin tubes containing substrate in a microcentrifuge. 9. To measure luminescent signals produced from reassembly of NanoLuc luciferase bound to DNA, first prepare the luminescent substrate by reconstitution at ambient temperature. Prepare a 20-fold dilution of the Nano-Glo® Live Cell Assay Substrate in the Nano-Glo® LCS Dilution Buffer by mixing 1 volume of substrate with 19 volumes of dilution buffer (see Note 12).

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10. Aspirate medium from cells in 96-well plates and replace with 100 μL of buffered cell culture medium, such as Opti-MEM® I Reduced Serum Medium (see Note 13) containing 0–10% FBS. 11. Add 25 μL reconstituted furimazine substrate to each well of the 96-well plate (see Notes 14 and 15), and gently mix the plate by hand or with an orbital shaker (e.g., 15 s at 300–500 rpm). 12. Measure luminescence immediately after adding the substrate on a luminescence-equipped plate reader using a 0.25–2 s integration time and a high PMT sensitivity setting (if applicable) for a single time point. Alternatively, measure luminescence continuously over 2 h for multiple time points (see Note 16). 13. To normalize for variations in transfection efficiency, cell number, and/or cell viability between wells, we typically divide luminescent signals by fluorescent signals obtained for the same well at the same time point. Normalized signals between different sgRNA pairs in different wells can now be compared. 3.3 Biosensor Transfection Setup for Microscopy-Based Measurement

1. Seed 10,000 to 25,000 low-passage cells per well in 96-well tissue culture plates or seed 100,000 to 250,000 low-passage cells per well in 24-well tissue culture plates approximately 12–20 h before transfection. 2. Transiently transfect cells in 96-well plates with 100 ng total DNA per well or transiently transfect cells in 24-well plates with 500 ng total DNA per well using either the Lipofectamine 3000 or equivalent cationic liposome-based transient transfection system or electroporation/nucleofection, similar to Subheading 3.2, step 4 (see Note 17) (Fig. 2). 3. 24 h post-transfection, check cells for GFP expression. If cells are not positive for green fluorescence or show lower transfection efficiency than expected, repeat steps 2–3. 4. Incubate cells for 24–72 h at 37 °C with 5% CO2 (see Note 11). 5. Prior to preparing microscope slides, set the microscope settings for low-light luminescence imaging. On the microscope, select the brightfield channel and set lamp intensity to 0, exposure time to 30 s, and sCMOS or EMCCD gain to at least 2.0. 6. For 24-well plate transfections, split the cells to 1.5 × 105 cells/ mL in Opti-MEM® I Reduced Serum Medium containing 0–10% FBS, and for cells in 96-well plate transfections, split the cells to 0.25 × 105 cells/mL in Opti-MEM® I Reduced Serum Medium containing 0–10% FBS. 7. Quickly transfer approximately 25–50 μL of cell suspension to microscope slides (see Note 18).

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8. Combine cell suspension at a 1:1 volumetric ratio with reconstituted furimazine substrate and mix gently by pipette. 9. Carefully place cover glass over 1:1 mixture of cell suspension and reconstituted furimazine substrate and quickly transfer slide to microscope (see Note 19). 10. To measure luminescent signals from NLuc reassembly, cover cells with a dark box (can be any opaque covering that completely blocks exogenous light from hitting the cells) with all external light sources in the area around the microscope blocked. Leaving any external light sources uncovered can lead to artifacts (see Note 20) (Fig. 2c). Measure luminescence as soon as possible after transporting the microscope slide to the area where the microscope is housed. 11. Immediately after measuring luminescence, measure GFP fluorescence using an epifluorescence microscope (Leica DM6000B or equivalent—should be equipped with at least sCMOS camera but preferably equipped with EMCCD camera). We typically use a sCMOS gain setting of 1.0 or greater and a relatively low exposure time (