Mammalian Cell Engineering: Methods and Protocols 1071614401, 9781071614402

Citation preview

Methods in Molecular Biology 2312

Ryosuke Kojima Editor

Mammalian Cell Engineering Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Mammalian Cell Engineering Methods and Protocols

Edited by

Ryosuke Kojima Graduate School of Medicine, The University of Tokyo, Tokyo, Japan

Editor Ryosuke Kojima Graduate School of Medicine The University of Tokyo Tokyo, Japan

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1440-2 ISBN 978-1-0716-1441-9 (eBook) https://doi.org/10.1007/978-1-0716-1441-9 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Synthetic biology is an emerging interdisciplinary area between biology and engineering, and recent advances in the field have made it possible to program mammalian cells to perform desired functions, paving the way toward treating intractable diseases using smartly engineered designer cells. In light of Richard Feynman’s famous words “What I cannot create, I do not understand,” the engineering of new cellular functions will no doubt deepen our understanding of the function and mechanism of various biological processes, advancing our basic biological knowledge. The act of “engineering mammalian cells” will therefore prove useful in various aspects. Due to the inherently complex nature of mammalian cells however, it is often difficult to engineer mammalian cells in a controlled and predictable manner. Considering that there are still many remaining “black boxes” regarding the function and actuation mechanisms of intracellular components, we cannot utilize or even interpret the outcome of engineered cellular functions unless we engineer mammalian cells using reliable and defined methods. Conversely, it is also of importance to actively incorporate cutting-edge technologies, which for the first time enable control of biological phenomena that have previously been difficult to reach. As such, in this book, I aim to introduce detailed, step-by-step protocols of the state-ofthe-art engineering methods of mammalian cells that are useful for controlling the ability/ performance of engineered mammalian cells for future cell-based therapeutics, enabling better understanding of complex biological systems. The overall topics can be categorized roughly into five parts, as follows: Part I (Chapters 1–5) describes methods to engineer mammalian cells to sense biologically relevant inputs, such as specific cell contacts, soluble proteins, blood glucose levels, and intracellular miRNA levels, which could be useful for various cell-based next-generation therapeutics (e.g., CAR-T cell therapy, cell-encapsulation-based therapies, regenerative medicine). Part II (Chapters 6–10) offers methods to engineer mammalian cells to sense artificial inputs, such as light, ultrasound, and exogenous small molecules, which could be useful for rationally and remotely controlling transgene expression levels at will with desired spatiotemporal resolutions. Part III (Chapters 11–13) provides cutting-edge CRISPR-Cas-based methods to carry out highly multiplexed genome editing, spatiotemporally controlled genome editing, as well as allowing for the high-throughput analysis of genome editing results. Part IV (Chapters 14–18) describes methods to control and engineer biological events in mammalian cells in combination with chemical compounds/systems, which could be useful in studying various biological processes that are otherwise difficult to analyze, or to control cellular properties by unconventional manners. Part V (Chapters 19–20) provides methods to engineer specific mammalian cells in targeted manners, which may find use in the study of functions of specific cells in complex biological systems. I sincerely hope that readers will find this book a useful resource, allowing them to successfully carry out their own projects, ultimately contributing to the further advancement of the entire field of mammalian cell engineering.

v

vi

Preface

To conclude, I greatly appreciate the contributions from all the authors who took time to write their chapters while also being busy with their research. Finally, I extend my gratitude to John Walker and the Springer editorial staff for their kind guidance and assistance and to my family for their continuous and unconditional support. Tokyo, Japan

Ryosuke Kojima

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

v ix

PART I ENGINEERING MAMMALIAN CELLS TO SENSE BIOLOGICALLY RELEVANT INPUTS 1 Generation of CAR-T Cells by Lentiviral Transduction . . . . . . . . . . . . . . . . . . . . . . Atsushi Okuma 2 Synthetic Receptors for Sensing Soluble Molecules with Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Leo Scheller 3 Engineering Mammalian Cells to Control Glucose Homeostasis . . . . . . . . . . . . . . Jiawei Shao, Xinyuan Qiu, and Mingqi Xie 4 Using Engineered Mammalian Cells for an Epitope-Directed Antibody Affinity Maturation System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Akihiro Eguchi and Masahiro Kawahara 5 Purification of Specific Cell Populations Differentiated from Stem Cells Using MicroRNA-Responsive Synthetic Messenger RNAs . . . . . . . . . . . . . . Hideyuki Nakanishi and Hirohide Saito

PART II

3

15 35

59

73

ENGINEERING MAMMALIAN CELLS TO SENSE ARTIFICIAL INPUTS

6 Green Light-Controlled Gene Switch for Mammalian and Plant Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nils Schneider, Claire V. Chatelle, Rocio Ochoa-Fernandez, Matias D. Zurbriggen, and Wilfried Weber 7 Sonogenetic Modulation of Cellular Activities in Mammalian Cells. . . . . . . . . . . . Yao-Shen Huang, Ching-Hsiang Fan, Wei-Ting Yang, Chih-Kuang Yeh, and Yu-Chun Lin 8 Constructing Smartphone-Controlled Optogenetic Switches in Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuanhuan Yu, Guiling Yu, and Haifeng Ye 9 Constructing a Smartphone-Controlled Semiautomatic Theranostic System for Glucose Homeostasis in Diabetic Mice . . . . . . . . . . . . . . . . . . . . . . . . . . Guiling Yu, Yuanhuan Yu, and Haifeng Ye 10 Construction of Caffeine-Inducible Gene Switches in Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daniel Bojar

vii

89

109

125

141

159

viii

Contents

PART III PRECISE GENOME ENGINEERING TECHNIQUES USING CRISPR-CAS SYSTEMS 11

Multiplexed Genome Engineering with Cas12a. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 Niels R. Weisbach, Ab Meijs, and Randall J. Platt 12 Highly Multiplexed Analysis of CRISPR Genome Editing Outcomes in Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Soh Ishiguro and Nozomu Yachie 13 Optical Control of Genome Editing by Photoactivatable Cas9 . . . . . . . . . . . . . . . 225 Takahiro Otabe, Yuta Nihongaki, and Moritoshi Sato

PART IV ENGINEERING MAMMALIAN CELLS IN COMBINATION WITH CHEMICAL COMPOUNDS/SYSTEMS 14

Chemogenetic Control of Protein Localization and Mammalian Cell Signaling by SLIPT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sachio Suzuki, Yuka Hatano, Tatsuyuki Yoshii, and Shinya Tsukiji 15 Engineering Hydrogel Production in Mammalian Cells to Synthetically Mimic RNA Granules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hideki Nakamura 16 AgDD System: A Chemical Controllable Protein Aggregates in Cells . . . . . . . . . Yusuke Miyazaki 17 Intracellular Unnatural Catalysis Enabled by an Artificial Metalloenzyme . . . . . . Yasunori Okamoto and Ryosuke Kojima 18 Feeder-Free Human Induced Pluripotent Stem Cell Culture Using a DNA Aptamer-Based Mimic of Basic Fibroblast Growth Factor . . . . . . . . . . . . . Yuri Hayata, Ryosuke Ueki, and Shinsuke Sando

237

253 277 287

301

PART V NEW TECHNIQUES TO ENGINEER SPECIFIC MAMMALIAN CELLS IN A TARGETED MANNER 19

Protocol for De Novo Gene Targeting Via In Utero Electroporation. . . . . . . . . . 309 Yuji Tsunekawa, Raymond Kunikane Terhune, and Fumio Matsuzaki 20 Magnetically Single-Cell Virus Stamping. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321 Rajib Schubert Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

329

Contributors DANIEL BOJAR • Wallenberg Centre for Molecular and Translational Medicine, University of Gothenburg, Gothenburg, Sweden; Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden CLAIRE V. CHATELLE • Signalling Research Centres BIOSS and CIBSS and Faculty of Biology, University of Freiburg, Freiburg, Germany; DSM Nutritional Products, Kaiseraugst, Switzerland AKIHIRO EGUCHI • Department of Chemistry and Biotechnology, Graduate School of Engineering, The University of Tokyo, Bunkyo-ku, Tokyo, Japan CHING-HSIANG FAN • Department of Biomedical Engineering and Environmental Sciences, National Tsing Hua University, Hsinchu, Taiwan YUKA HATANO • Department of Life Science and Applied Chemistry, Nagoya Institute of Technology, Showa-ku, Nagoya, Japan YURI HAYATA • Department of Chemistry and Biotechnology, The University of Tokyo, Tokyo, Japan YAO-SHEN HUANG • Institute of Molecular Medicine, National Tsing Hua University, Hsinchu, Taiwan SOH ISHIGURO • School of Biomedical Engineering, University of British Columbia, Vancouver, BC, Canada; Research Center for Advanced Science and Technology, University of Tokyo, Tokyo, Japan MASAHIRO KAWAHARA • Department of Chemistry and Biotechnology, Graduate School of Engineering, The University of Tokyo, Bunkyo-ku, Tokyo, Japan; Laboratory of Cell Vaccine, Center for Vaccine and Adjuvant Research, National Institutes of Biomedical Innovation, Health and Nutrition (NIBIOHN), Ibaraki-shi, Osaka, Japan RYOSUKE KOJIMA • Graduate School of Medicine, The University of Tokyo, Bunkyo-ku, Tokyo, Japan; PRESTO, Japan Science and Technology Agency, Kawaguchi, Saitama, Japan YU-CHUN LIN • Institute of Molecular Medicine, National Tsing Hua University, Hsinchu, Taiwan; Department of Molecular Medicine, National Tsing Hua University, Hsinchu, Taiwan FUMIO MATSUZAKI • Laboratory for Cell Asymmetry, RIKEN Center for Biosystems Dynamics Research, Kobe, Japan; Laboratory of Molecular Cell Biology and Development, Department of Animal Development and Physiology, Graduate School of Biostudies, Kyoto University, Kyoto, Japan AB MEIJS • Department of Biosystems Science and Engineering, ETH Zurich, Basel, Switzerland YUSUKE MIYAZAKI • ANRI, Tokyo, Japan HIDEKI NAKAMURA • Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Nishikyo-ku, Kyoto, Japan; JSTERATO, Hamachi Innovative Molecular Technology for Neuroscience, Kyoto University, Nishikyo-ku, Kyoto, Japan HIDEYUKI NAKANISHI • Department of Biofunction Research, Institute of Biomaterials and Bioengineering, Tokyo Medical and Dental University (TMDU), Chiyoda-ku, Tokyo, Japan; Department of Life Science Frontiers, Center for iPS Cell Research and Application, Kyoto University, Sakyo-ku, Kyoto, Japan

ix

x

Contributors

YUTA NIHONGAKI • Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, MD, USA ROCIO OCHOA-FERNANDEZ • Institute of Synthetic Biology and iGRAD Plant Graduate School, University of Du¨sseldorf, Du¨sseldorf, Germany YASUNORI OKAMOTO • Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, Sendai, Japan ATSUSHI OKUMA • Center for Exploratory Research, Research & Development Group, Hitachi Ltd., Chiyoda City, Tokyo, Japan TAKAHIRO OTABE • Graduate School of Arts and Sciences, The University of Tokyo, Meguroku, Tokyo, Japan RANDALL J. PLATT • Department of Biosystems Science and Engineering, ETH Zurich, Basel, Switzerland; Department of Chemistry, University of Basel, Basel, Switzerland; Botnar Research Center for Child Health, Basel, Switzerland XINYUAN QIU • Key Laboratory of Growth Regulation and Translational Research of Zhejiang Province, School of Life Sciences, Westlake University, Hangzhou, Zhejiang, China; Department of Biology and Chemistry, College of Liberal Arts and Sciences, National University of Defense Technology, Changsha, Hunan, China HIROHIDE SAITO • Department of Life Science Frontiers, Center for iPS Cell Research and Application, Kyoto University, Sakyo-ku, Kyoto, Japan SHINSUKE SANDO • Department of Chemistry and Biotechnology, The University of Tokyo, Tokyo, Japan; Department of Bioengineering, The University of Tokyo, Tokyo, Japan MORITOSHI SATO • Graduate School of Arts and Sciences, The University of Tokyo, Meguro-ku, Tokyo, Japan LEO SCHELLER • Institute of Bioengineering, E´cole Polytechnique Fe´de´rale de Lausanne, Lausanne, Switzerland NILS SCHNEIDER • Signalling Research Centres BIOSS and CIBSS and Faculty of Biology, University of Freiburg, Freiburg, Germany; Celonic AG, Basel, Switzerland RAJIB SCHUBERT • Research and Early Development, Roche Molecular Systems, Pleasanton, CA, USA JIAWEI SHAO • Key Laboratory of Growth Regulation and Translational Research of Zhejiang Province, School of Life Sciences, Westlake University, Hangzhou, Zhejiang, China; Westlake Laboratory of Life Sciences and Biomedicine, Hangzhou, Zhejiang, China; Institute of Basic Medical Sciences, Westlake Institute for Advanced Study, Hangzhou, Zhejiang, China SACHIO SUZUKI • Department of Nanopharmaceutical Sciences, Nagoya Institute of Technology, Showa-ku, Nagoya, Japan RAYMOND KUNIKANE TERHUNE • Public Relations Division, Office of Global Communications, Kyoto University, Kyoto, Japan SHINYA TSUKIJI • Department of Nanopharmaceutical Sciences, Nagoya Institute of Technology, Showa-ku, Nagoya, Japan; Department of Life Science and Applied Chemistry, Nagoya Institute of Technology, Showa-ku, Nagoya, Japan YUJI TSUNEKAWA • Laboratory for Cell Asymmetry, RIKEN Center for Biosystems Dynamics Research, Kobe, Japan; Division of Molecular and Medical Genetics, Institute of Medical Science, University of Tokyo, Tokyo, Japan RYOSUKE UEKI • Department of Chemistry and Biotechnology, The University of Tokyo, Tokyo, Japan WILFRIED WEBER • Signalling Research Centres BIOSS and CIBSS and Faculty of Biology, University of Freiburg, Freiburg, Germany

Contributors

xi

NIELS R. WEISBACH • Department of Biosystems Science and Engineering, ETH Zurich, Basel, Switzerland MINGQI XIE • Key Laboratory of Growth Regulation and Translational Research of Zhejiang Province, School of Life Sciences, Westlake University, Hangzhou, Zhejiang, China; Westlake Laboratory of Life Sciences and Biomedicine, Hangzhou, Zhejiang, China; Institute of Basic Medical Sciences, Westlake Institute for Advanced Study, Hangzhou, Zhejiang, China NOZOMU YACHIE • School of Biomedical Engineering, University of British Columbia, Vancouver, BC, Canada; Research Center for Advanced Science and Technology, University of Tokyo, Tokyo, Japan; Institute for Advanced Biosciences, Keio University, Tsuruoka, Japan WEI-TING YANG • Institute of Molecular Medicine, National Tsing Hua University, Hsinchu, Taiwan HAIFENG YE • Synthetic Biology and Biomedical Engineering Laboratory, Biomedical Synthetic Biology Research Center, Shanghai Key Laboratory of Regulatory Biology, Institute of Biomedical Sciences and School of Life Sciences, East China Normal University, Shanghai, China CHIH-KUANG YEH • Department of Biomedical Engineering and Environmental Sciences, National Tsing Hua University, Hsinchu, Taiwan TATSUYUKI YOSHII • Department of Life Science and Applied Chemistry, Nagoya Institute of Technology, Showa-ku, Nagoya, Japan; PRESTO, Japan Science and Technology Agency (JST), Kawaguchi, Saitama, Japan GUILING YU • Synthetic Biology and Biomedical Engineering Laboratory, Biomedical Synthetic Biology Research Center, Shanghai Key Laboratory of Regulatory Biology, Institute of Biomedical Sciences and School of Life Sciences, East China Normal University, Shanghai, China YUANHUAN YU • Synthetic Biology and Biomedical Engineering Laboratory, Biomedical Synthetic Biology Research Center, Shanghai Key Laboratory of Regulatory Biology, Institute of Biomedical Sciences and School of Life Sciences, East China Normal University, Shanghai, China MATIAS D. ZURBRIGGEN • Institute of Synthetic Biology and iGRAD Plant Graduate School, University of Du¨sseldorf, Du¨sseldorf, Germany; CEPLAS—Cluster of Excellence on Plant Sciences, Du¨sseldorf, Germany

Part I Engineering Mammalian Cells to Sense Biologically Relevant Inputs

Chapter 1 Generation of CAR-T Cells by Lentiviral Transduction Atsushi Okuma Abstract CAR-T cell therapy is one of the most successful cell-based therapies. T cells are the most common cells to be genetically modified for cancer therapy, not only because T cells have cytotoxicity but also because they are easily cultured ex vivo and genetically modified with viral vectors. Hence, for nonexperts, T cell engineering is an ideal starting point for mammalian cell engineering or for development of therapeutics. Here, we have described a basic procedure for lentiviral transduction of human primary T cells to generate a CAR-T cell and assays to confirm CAR expression and function. Key words CAR-T cell, Lentivirus vector, Dynabeads, PEG precipitation, Retronectin, Flow cytometry, CAR-T cell activation assay

1

Introduction CAR-T cells have been successfully used as a clinical therapy for B cell malignancies [1–3]. More than 500 CAR-T cell-related clinical trials are currently in progress [4, 5]. The CAR is composed of a single-chain variable fragment (scFv) and signaling domains derived from TCR (CD3z) and costimulatory receptors such as CD28 or 4-1BB [1]. The scFv is derived from the antigen-binding part of an antibody and expressed on the cell surface. When the CARs bind to antigens on a target cell, T cells are stimulated by CAR signaling domains and kill target cells. CAR-T cell therapy has considerably expanded the promise of cell therapies using genetic engineering technology. Researchers in the fields of medicine and synthetic biology often develop new CAR-T cells as a model to prove the clinical usefulness of their synthetic biology products or techniques [6–10]. However, primary T cell handling and gene transfer techniques are considered as barriers. Lentiviral vectors have been widely used to engineer primary immune cells for adoptive cell therapies such as CAR-T cell therapy [11]. The advantages of lentiviral transduction include high transduction efficiency, high T cell viability after gene delivery, stable

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

3

4

Atsushi Okuma

expression of the transferred gene, and low risk of transformation [12]. Thus, we have described a detailed protocol for lentiviral transduction of human primary T cells which consists of lentivirus production, T cell isolation, and infection. This method can yield a 50–90% CAR-positive population in more than 1
108 T cells within a month from 1
106 T cells. We have also described rapid assays to validate CAR expression and function. Considering the target cell type and available equipment, the appropriate assay can be chosen to quantify CARs.

2

Materials

2.1 Vectors (See Note 1)

1. pHR-SFFV (Addgene #79121): Place CAR sequences of interest under SFFV promoter. 2. Packaging plasmid mix: Ratio of pCMVR8.74 (Addgene #22036): pMD2.G (Addgene #12259): pAdVAntage (Promega #E1711) ¼ 15: 5: 2.

2.2 Packaging Cell Transfection

1. Packaging cell: 293FT (Thermo Fisher Scientific #R70007). 2. Culture flask: T-175 tissue culture flask. 3. Culture media: Dulbecco’s Modified Eagle Medium (DMEM) containing 10% FBS, 10 U/mL penicillin (pen), 10 μg/m streptomycin (strep), 2 mmol/L L-glutamine, and 1 mmol/L sodium pyruvate. 4. Transfection Reagent: 0.323 g/L PEI solution, pH 8.0—Dissolve Polyethylenimine “Max” (Polyscience #24765-1) in culture grade water by mixing and heating to 50  C, then adjust the pH with NaOH and store at 80  C in 1 mL aliquots (see Note 2). 5. Phosphate-Buffered Saline (PBS(); see Note 3).

2.3 Virus Concentration

1. Virus production media: UltraCULTURE (Lonza #2-725F) containing 10 U/mL pen, 10 μg/m strep, 2 mmol/L L-glutamine, 1 mmol/L sodium pyruvate, and 5 mM sodium butylate. 2. PEG solution: Add 80 g of polyethylene glycol 8000 (final concentration of 40% w/v) and 14 g of NaCl (final concentration of 1.2 M) into 100 mL PBS. Mix by heating to 50  C until the solids are dissolved, adjust the final volume to 200 mL, and sterilize the solution using a 0.2-μm bottle top filter.

2.4

T Cell Isolation

1. Peripheral blood (see Note 4). 2. T cell isolation kit (STEMCELL Technologies #15063; see Note 5). 3. SepMATE (STEMCELL Technologies #86450).

Generation of CAR-T Cells by Lentiviral Transduction

5

4. Lymphoprep (STEMCELL Technologies #07801). 5. Frozen stock buffer: 90% FBS, 10% DMSO. 6. Cryotubes. 7. Freezing container. 2.5 T Cell Culture and Transduction

1. Primary T cell media: Add 50 mL of human serum (Valley Biomedical #HP1022; final concentration of 5%), 10 mL of 2-mercaptoethanol solution, 10 mL of 1 M N-acetyl-L-cysteine dissolved in PBS into 1 L of X-VIVO 15 (Lonza #04-418Q) and add recombinant IL-2 (final concentration of 100 U/mL) just before use. 2. T cell activator beads: Dynabeads human T-activator CD3/CD28 (Thermo Fisher Scientific #11131D). 3. Blocking buffer for Dynabeads: 0.1% bovine serum albumin and 2 mM EDTA in PBS sterilized using a 0.2-μm filter. 4. Magnet stand (STEMCELL #ST-18000). 5. RetroNectin (Takara bio #T100B): Dilute to 50 μg/mL with PBS. 6. Blocking buffer for RetroNectin-coated plate: 2% bovine serum albumin in PBS sterilized using a 0.2-μm filter. 7. 6-well nontreated plate.

2.6

Validation Assay

Antibodies: Anti-Myc-PE (Cell Signaling Technology #3739S; 1:100 dilution), Anti-CD69-APC (BioLegend #104514; 1:200 dilution). 1. FACS buffer: PBS containing 2% FBS and 0.01% w/v NaN3. 2. 96-well flat-bottom nontreated plate. 3. 96-well round-bottom plate.

3

Methods

3.1 Packaging Cell Transfection

1. Prepare nearly confluent (80–90%) packaging cells in a T175 flask (see Notes 6–8). 2. Mix 1.25 mL of PBS, 25 μg of packaging plasmid mix, and 25 μg of CAR vector of interest. 3. Mix 875 μL of PBS and 375 μL of PEI. 4. Combine with the solution in step 2 and incubate for 15 min at room temperature (RT). 5. Grab the flask upside-down, add the entire volume of the DNA-PEI solution to the culture medium, and mix by pipetting carefully to avoid detaching cells. 6. Culture at 37  C in 5% CO2 for 24 h.

6

Atsushi Okuma

3.2 Virus Concentration (See Note 9)

1. Discard the culture medium and add 25 mL of virus production medium. 2. Culture at 37  C in 5% CO2 for 24 h. 3. Collect conditioned medium and store at 4  C. 4. Add 25 mL of fresh virus production medium. 5. Culture at 37  C in 5% CO2 for 24 h. 6. Collect conditioned medium and store at 4  C. 7. Add 25 mL of fresh virus production medium. 8. Culture at 37  C in 5% CO2 for 24 h. 9. Collect conditioned medium and centrifuge it at 3000
g for 10 min at 4  C to remove cell debris. 10. Add 25 mL of PEG solution to the entire supernatant containing conditioned medium (75 mL), mix it gently but thoroughly, and store it at 4  C overnight. 11. Centrifuge the medium at 1600
g for 60 min at 4  C. 12. Carefully remove the supernatant without disturbing the pellet. 13. Thoroughly resuspend the viral pellet with 2 mL of PBS or desired medium (see Note 10). 14. Aliquot 500 μL each and store at 80  C (see Note 11).

3.3

T Cell Isolation

1. Add 750 μL of RossetteSep cocktail to 15 mL of peripheral blood from a healthy donor. 2. Mix the sample gently and incubate at RT for 10 min. 3. Dilute the sample with an equal volume (15 mL) of 2% FBS in PBS. 4. Add 15 mL of Lymphoprep to a SepMATE tube. 5. Add entire volume (30 mL) of diluted sample to the SepMATE tube containing Lymphoprep. 6. Centrifuge at 1200
g for 10 min. 7. Transfer the supernatant to a new 50-mL tube. 8. Top up the 50-mL tube with 2% FBS in PBS to wash isolated cells. 9. Centrifuge the sample at 300
g for 10 min. 10. Wash the cells with 2% FBS in PBS again. 11. Resuspend the cells with frozen stock buffer, aliquot 1
106 cells per cryotube, freeze them slowly in the freezing container, and store them in a liquid nitrogen tank.

Generation of CAR-T Cells by Lentiviral Transduction

3.4 T Cell Transduction

7

1. Adjust the concentration of isolated human primary T cells to 1
106 cells/mL and preculture in Primary T cell medium at 37  C in 5% CO2 overnight (see Note 12). 2. Resuspend the T cell activator beads by vortex and transfer the desired volume of beads to a 1.5-mL tube (50 μL per 1
106 T cells). 3. Add an equal volume of blocking buffer and incubate at RT for 5 min. 4. Place the tube on the magnet stand for 1 min and discard the supernatant. 5. Remove the tube from the magnet stand and resuspend the beads with culture medium. 6. Transfer the resuspended beads to T cell culture medium. 7. Culture at 37  C in 5% CO2 for 24 h. 8. Prepare Retronectin-coated plate by adding 2.25 mL of diluted Retronectin to a well of a 6-well nontreated plate and store it at 4  C overnight. 9. Aspirate the Retronectin solution and block with 3 mL of blocking buffer for 30 min at RT. 10. Aspirate the blocking buffer and wash with 3 mL of PBS. 11. Add 2 mL of the concentrated virus solution. 12. Centrifuge at 1200
g for 90 min at RT (see Note 13). 13. Discard the supernatant and wash with 3 mL of blocking buffer. 14. Dilute activated T cells to 2.5
105 cells/mL in fresh primary T cell medium and add 4 mL of diluted T cell culture to a viruscoated well. 15. Centrifuge at 1200
g for 10 min at RT. 16. Culture at 37  C in 5% CO2 for 48 h. 17. Remove the beads and add fresh primary T cell medium to adjust the cell concentration to 1
106 cells/mL. 18. Keep the cell concentration at 1
106 cells/mL by adding fresh primary T cell medium. 19. Around day 3–7, the transduced T cells are ready to be used for assay (see Notes 14 and 15).

3.5

Validation Assay

Surface expression confirmation: We usually place an Myc tag between scFv and the transmembrane domain—the Myc tag can be stained with an Myc antibody to confirm if CAR is expressed on the surface. 1. Transfer 1
105 transduced cells to a 1.5-mL tube and centrifuge at 300
g for 5 min.

8

Atsushi Okuma

-6 -5 -4

Split packaging cell Transfection Change media

-3

Collect media

-2

Collect media

-1

Collect media & Mix with PEG solution T cell activation Coat plate by Retronectin

Day 0

Virus concentration Infection

2

Remove activator beads Add fresh media

5

Check CAR expression

Fig. 1 Protocol timeline. A timeline of the transduction protocol in which infection is set as day 0. This protocol generates transduced T cells within 2 weeks

2. Suspend the pellet with anti-Myc-PE antibody diluted in FACS buffer. 3. Incubate at RT in the dark for 30 min. 4. Wash with FACS buffer twice. 5. Flow cytometry analysis (Fig. 1). CAR-T cell activation assay with plate-bound antigen: If a recombinant target antigen (e.g., Axl-Fc, R&D systems #154-AL) is available, it can be determined whether transduced T cells are activated by the plate-bound antigen. 1. Dilute the recombinant antigen to 10 μg/mL with PBS. 2. Add 100 μL of the diluted antigen to a well of a 96-well nontreated flat-bottom plate (1 μg/well; see Note 16). 3. Incubate at 4  C overnight. 4. Wash the plate with PBS three times to remove the unbound antigen. 5. Add 200 μL of 1
106 cells/mL transduced T cells. 6. Culture at 37  C in 5% CO2 for 24 h. 7. Centrifuge the cells at 300
g for 5 min. 8. Suspend the pellet with anti-Myc-PE and anti-CD69-APC antibodies diluted in FACS buffer. 9. Incubate at RT in the dark for 30 min. 10. Wash with FACS buffer twice. 11. Flow cytometry analysis (Fig. 2).

Generation of CAR-T Cells by Lentiviral Transduction

9

Fig. 2 CAR expression on the cell surface. A histogram of representative data of Myc staining on day 5. The gray histogram depicts nontransduced T cells as a negative control. The red histogram depicts anti-Axl-CAR-transduced T cells [8]

CAR-T cell activation assay with antigen-expressing cells: If a cell line expressing the target antigen is available (e.g., Axl+, SK-OV-3; ATCC #HTB-77; see Note 17), it can be used for this assay. If not, then a new target antigen-overexpressing cell line can be generated by transducing a cell line (e.g., K562, Nalm-6, Jurkat) lentivirally. 1. Mix transduced T cells (Effector) and target cells (Target) at the desired ratio (see Note 18; e.g., add 100 μL of 1
106 cells/mL Effector and 100 μL of 1
106 cells/mL Target to a well of a 96-well round-bottom plate. 2. Coculture at 37  C in 5% CO2 for 24 h. 3. Measure live target cells (see Note 19; Figs. 3 and 4).

4

Notes 1. Plasmid preparation is one of the most important steps for adequate transfection efficiency. If high toxicity of transfection is observed, plasmids purified by low endotoxin midiprep/ maxiprep kits (e.g., NucleoBond® Xtra Midi Plus EF, Takara bio #U0422A) may improve cell viability. 2. After the PEI solution is thawed, it can be kept at 4  C for at least 4 weeks.

10

Atsushi Okuma

Fig. 3 CAR-T cell activation assay using a plate-bound antigen. (a) A schematic illustration of the CAR-T cell activation assay using a plate-bound antigen. (b) Gating strategy to evaluate T cell activation. The expression of CD69, a T cell activation marker, is shown in the Myc (CAR)-positive population. (c) The dynamic range of the CAR response is depicted by CD69 expression measured after 24 h of culturing anti-Axl CAR-expressing T cells with varying amounts of plate-bound Axl protein (reproduced from [8] (licensed under CC BY 4.0))

3. The original protocol recommends 0.15 M NaCl, but there is no significant difference in the transfection efficiency when 0.15 M PBS is used. 4. If it is difficult to obtain human blood from healthy donors, cryopreserved peripheral blood mononuclear cells (PBMCs) are commercially available (e.g., STEMCELL Technologies #ST-70025) and can be used. 5. If CD8+ T cells are isolated from PBMCs (see Note 4), a different kit must be used (e.g., EasySep Neg Human CD8 T Cell Kit, STEMCELL Technologies #ST-19053). 6. Do not change the culture medium before transfection. 7. To prepare 80–90% confluent packaging cells, split 25 mL of 1.2
106 cells/mL 293FT cells in a T175 flask and culture for 18 h. Optimize the split cell number and culture time using your cells prior to this experiment. 8. Packaging cells in a T175 flask produces enough virus for transduction of 1
106 T cells. This can be scaled down as follows: T75 packaging cell/5
105 T cells/Retronectincoated 12-well plate, T25 packaging cell/1
105 T cells/ Retronectin-coated 48-well plate.

Generation of CAR-T Cells by Lentiviral Transduction

11

Fig. 4 Flow cytometry-based cell counting for the killing assay. (a) A schematic illustration of the killing assay. T cells express anti-Axl CAR and target cells express Axl. (b) Flow cytometry plots of the cell mixture after 24 h of coculture of 0.5
106 anti-Axl CAR T cells with 1
106 Axl+ Jurkat cells. Axl+ Jurkat cells express both GFP and mCherry. The numbers in the plots indicate the percentage of Jurkat cells in the whole mixture of cells, but the absolute number of Jurkat cells need to be counted for the killing assay. (c and d) Graphs that indicate the absolute number of Axl+ Nalm-6 (c) and Jurkat (d) cells after coculture at different E:T ratios. The number of Target cells at the start of the culture was normalized across all conditions. Black dots indicate the basal killing activity of primary T cells. 80%, E:T ¼ 4:1; 67%, E:T ¼ 2:1; 50%, E:T ¼ 1:1; 33%, E:T ¼ 1:2; 20%, E:T ¼ 1:4; 11%, E:T ¼ 1:8; 5.9%, E:T ¼ 1:16; 0.75%, E:T ¼ 1:32

9. After transfection, packaging cells produce lentivirus. Carefully handle and discard the medium, flask, and disposable pipettes according to your organization’s guidelines. 10. Optionally, you can check the titer of the concentrated virus (e.g., Lenti-X™ qRT-PCR Titration Kit; Clontech #Z1235). Titration is only usually performed when a huge batch of virus is made. 11. The virus can be kept at 80  C for at least 6 months. Do not repeatedly freeze and thaw them. 12. If the T cells were used immediately after isolation, this preculture step can be skipped. 13. Instead of this centrifuge step, incubation in a CO2 incubator for 4 h works as well. Usually, 50–90% Myc-positive cells are detected around day 5. However, in some cases, positive transduction rates were much lower (4 h) to validate the disease model; mice with blood glucose levels of higher than 10 mM are considered diabetic. 2. Implantation of encapsulated designer cells. (a) For implantation, resuspend the encapsulated cells (Subheading 3.3.1) in serum-free DMEM or MOPS at a density of ~5  106 cells/mL (2.5  104 capsules/mouse). Capsules can be counted by naked eye using a small representative sample of 10 mM of fasting blood glucose), which can be achieved through continuous feeding of high fat diet [38], injection of alloxan [5, 6] or streptozotocin [20], or genetic models such as Lepdb/db and Lepob/ob [39]. 9. Any closed-loop control system essentially aims to approach the biological performance and treatment efficacy of human islets [10]. A single dose of clinical-grade human islets typically shows a fivefold improvement of glucose-stimulated insulin secretion (GSIS) in vitro [40], restores normoglycemia within a few days following implantation, and should provide insulin independence to T1D patients for several years [41]. Indeed, these parameters are also the key observables when assessing the goodness of β-cells produced by stem cell differentiation [42]. One central drawback of stem cells–derived β-cells is that high-quality batches of β-cells cannot be freeze-stored for reuse, and that individual differentiation runs typically show very large variability in bioprocess efficiency [43]. Therefore, β-cell mimetic designer cells should ideally also enable costeffective and GMP-compliant production as well as compatibility with cryopreservation techniques to yield ready-to-use commercial products with clear specifications for qualityassurance, batch-to-batch identity, purity, and stability. 10. To validate and optimize the gene circuits to reach the intended functionality and performance with high throughput, transient transfections are preferred. Also, reporters that are easy to be detected (such as SEAP and Luciferase) are also preferred during this process to evaluate the dynamics of the circuits. 11. HEK-β. To engineer a synthetic promoter with optimal Ca2+ responsiveness, we transfected different calcium-specific reporter vectors into HEK-293 cells, such as pMX53 (PcFOSSEAP-pA), pHY30 (PNFAT1-SEAP-pA), pMX56 (PNFAT2SEAP-pA), and pKR32 (PNFkB-SEAP-pA). Each vector used a different calcium-specific promoter driving the expression of human placental secreted alkaline phosphatase SEAP as a reporter protein. Results showed that the synthetic promoter PNFAT2, which contained three tandem NFAT (nuclear factor of activated T cells) binding sites derived from the murine interleukin-4 (IL-4) promoter, was most responsive to membrane depolarization induced by potassium chloride. Building on this finding, we further optimized the PNFAT2 promoter by multimerizing the NFAT-repeats (50 -TACATTGGAAAATTT TAT-30 ) upstream of a minimal TATA-box promoter (50 -TAG AGGGTATATAATGGAAGCTCGACTTCCAG -30 ). Results

Engineering Mammalian Cells to Control Glucose Homeostasis

53

showed that PNFAT3 promoter, which had five NFATIL4 repeats, showed optimal induction ratio between resting and depolarized membrane potentials. Hence, we then used the plasmid pMX57 (PNFAT3-SEAP-pA) as the reporter vector to optimize the glucose-sensing gene componentry (GLUT2, GCK, KATP, Cav1.3). We found that ectopic overexpression of KATP or GCK did not improve glucose-induced calcium-dependent transcription, which was consistent to semiquantitative transcriptional profiling results showing that other isoforms of KATP and GCK were already endogenously expressed in HEK-293 cells. Although overexpression of GLUT2 increased overall SEAP expression levels, there was no significant changes in the fold-changes of glucose-induced SEAP expression. In fact, semiquantitative transcriptional profiling showed that the GLUT1 transporter was highly expressed in HEK-293 cells, which has highly similar functions as GLUT2. Instead, ectopic expression of the Cav1.3 channel was most critical for conferring glucose sensitivity to HEK-293 cells. Therefore, overexpression of the Cav1.3 channel and a synthetic promoter containing at least five NFATIL4 repeats driving the expression of a target gene of interest was sufficient to form a synthetic gene circuit that enables glucose-dependent gene expression in HEK-293 cells. 12. FRL. To achieve optimal far-red light-dependent SEAP expression, we first created different synthetic mammalian transactivators (FRTAs) by assembling BldD, p65 [65-kDa transactivator subunit of nuclear factor kB (NF-kB)], VP64 (tetrameric core of herpes simplex virus–derived transactivation domain), and HSF1 (heat shock factor 1) into various configurations and constructed multiple FRTA specific chimeric promoter (PFRL2.x) variants. We found that a combination of FRTA3 (p65-VP64-NLS-BldD) and PFRL2.13a (pA-(whiG)3PhCMVmin) showed the best FRL triggered BphS/YhjHmediated transcriptional response. The FRL system was functional in many mammalian cell lines, including HeLa, hMSCTERT, Hana3A and HEK-293, where it afforded over 60-fold inductions of far-red-light-dependent SEAP expression. 13. For stable transfection, several integration and selection methods can be applied. Transposase-directed and lentivirusmediated transgene integration are the most widely used approaches for effective integration of large-scale gene circuits into random loci of the host genome. Genome editing tools (CRISPR/Cas9, TALEN, etc.) can also be used to directly knock the gene circuit into specific loci. For multicomponent gene circuits, we prefer the use of the Sleeping Beauty transposase system that is based on random integration, as it usually

54

Jiawei Shao et al.

offers a large and highly variable pool of functional (and nonfunctional) individual cell clones to allow demand-driven selection according to a clear expectation profile. 14. For effective antibiotic selection, it is recommended not to cotransfect multiple transposons containing different resistance genes. 15. Antibiotic concentration should be adjusted based on the selection efficiency. 16. For integration of multiple transposons with different resistance genes, it is recommended to start with the “slowest” selection agent. Selection agent

Selection time

Zeocin (100 μg/mL)

7–10 days

Blasticidin (10 μg/mL)

2–3 days

Puromycin (0.5 μg/mL)

1–2 days

17. Cells should be diluted to obtain single colony generated from a single cell. Colonies should be rapidly removed and reseeded into the 96-well plate once trypsinized to avoid cell death. 18. The criteria of choosing a “high-functional” clone can vary from case to case. Parameters include fold-change, dynamic range and expression kinetics. 19. As type-1 diabetes mellitus is an autoimmune disorder characterized by repeated immune attack against pancreatic β-cells, it is recommended to encapsulate any kind of therapeutic cells (e.g., islets, β-like cells, designer cells) into immunoprotective semipermeable implants. This avoids the need for immunosuppression, resulting in optimal therapeutic activity and lifetime as well as patient safety. In diabetic mouse models, it is recommended to implant up to 5  106 cells per animal—a reference dosage reported in β-cell–based treatments [17]. For each treatment, cryopreserved (monoclonal) designer cells are thawed, passaged, and expanded to reach implantationcompatible cell numbers. To this end, it is especially valuable that designer cells can be cost-effectively produced, stored, and expanded according to standardized protocols, providing quality-assured and ready-to-use medicinal products with high batch-to-batch identity. As mentioned above, alginate microencapsulation is the state-of-the-art technology for implantation of glucose-sensitive insulin-producing cells to treat diabetes [21, 22, 31]. 20. In studies with stem cell-derived β-cells, alginate beads of 1.5 mm in diameter proved to be superior than smaller beads (500 μm) in terms of long-term efficacy in vivo [31].

Engineering Mammalian Cells to Control Glucose Homeostasis

55

21. This is the most critical step of the encapsulation process as determination of a proper flow rate indeed requires some experience. The magnetic stirrer should be turned on to prevent clumping of freshly formed polymeric beads. 22. STZ master mixes can be prepared for mice with similar body weights to simplify the injection and dilution process. 23. Weighting, dilution and injection of STZ should be conducted under low temperature while avoiding light exposure. 24. While homeostatic blood glucose levels (5–10 mM) of mice treated with HEK-β should be restored within 2–3 days and remain in this range over the entire experimental time window (e.g., 3 weeks), the glycemia of wild-type nondiabetic mice should not be affected by HEK-β implantation (Fig. 3a). This would indicate glucose-dependent insulin secretion and the absence of basal insulin expression potentially causing hypoglycemic side effects. 25. Blood insulin levels were measured with Mercodia insulin ELISA kits. Increase of blood insulin levels should correlate with the decrease in fasting glycemia of all treatment groups (Fig. 3b). Blood insulin levels of higher than 0.6 ng/mL are considered homeostatic. 26. Glucose tolerance tests (GTT) indicate the goodness of postprandial glucose metabolism. Whereas oral glucose tolerance tests (OGTT) more closely mimic an animal’s physiological response to meal intake, intraperitoneal glucose tolerance tests (IPGTT) directly reflects the pharmacokinetics of bioactive insulin. In fact, OGTT might be preferred to study the incretin effect in T2D models [44]. Here, in order to primarily assess the insulin-effect, IPGTT experiments are sufficient. Mice are considered GTT-positive when the glycemia at 120 min returns to similar levels as the glycemia at time zero (Fig. 3c). Area under the curve (AUC) analysis and two-way ANOVA analysis can also be used as criteria to judge GTT results (Fig. 3d). 27. Because the measurement of glycemia takes approximately 30 s per mouse, it is recommended not to do a same GTT experiment with more than 24 mice at once.

Acknowledgments We thank Minghui He for critical discussions on the manuscript. Work in the laboratory of M.X. (MingLab) is supported by Westlake Education Foundation, Tencent Foundation, the National Natural Science Foundation of China (32071429) and the Ministry of Science and Technology (2020YFA0909200).

56

Jiawei Shao et al.

References 1. NCD Risk Factor Collaboration (2016) Worldwide trends in diabetes since 1980: a pooled analysis of 751 population-based studies with 4.4 million participants. Lancet 387 (10027):1513–1530 2. Latres E, Finan DA, Greenstein JL et al (2019) Navigating two roads to glucose normalization in diabetes: automated insulin delivery devices and cell therapy. Cell Metab 29(3):545–563 3. Kieffer TJ (2016) Closing in on mass production of mature human Beta cells. Cell Stem Cell 18(6):699–702 4. Dolgin E (2016) Diabetes: encapsulating the problem. Nature 540(7632):S60–S62 5. Xie M, Ye H, Wang H et al (2016) Beta-cellmimetic designer cells provide closed-loop glycemic control. Science 354(6317):1296–1301 6. Auslander D, Auslander S, Charpin-El Hamri G et al (2014) A synthetic multifunctional mammalian pH sensor and CO2 transgenecontrol device. Mol Cell 55(3):397–408 7. Russell SJ, El-Khatib FH, Sinha M et al (2014) Outpatient glycemic control with a bionic pancreas in type 1 diabetes. N Engl J Med 371 (4):313–325 8. Chen Z, Wang J, Sun W et al (2018) Synthetic beta cells for fusion-mediated dynamic insulin secretion. Nat Chem Biol 14(1):86–93 9. Forlenza GP, Buckingham B, Maahs DM (2016) Progress in diabetes technology: developments in insulin pumps, continuous glucose monitors, and Progress towards the artificial pancreas. J Pediatr 169:13–20 10. Xie M, Aubel D, Fussenegger M (2017) Closed-loop control systems—the quest for precision therapies for diabetes. Curr Opin Syst Biol 5:32–40 11. Shapiro AM, Lakey JR, Ryan EA et al (2000) Islet transplantation in seven patients with type 1 diabetes mellitus using a glucocorticoid-free immunosuppressive regimen. N Engl J Med 343(4):230–238 12. Ricordi C, Goldstein JS, Balamurugan AN et al (2016) National Institutes of Healthsponsored clinical islet transplantation consortium phase 3 trial: manufacture of a complex cellular product at eight processing facilities. Diabetes 65(11):3418–3428 13. Wu J, Vilarino M, Suzuki K et al (2017) CRISPR-Cas9 mediated one-step disabling of pancreatogenesis in pigs. Sci Rep 7(1):10487 14. Yamaguchi T, Sato H, Kato-Itoh M et al (2017) Interspecies organogenesis generates autologous functional islets. Nature 542 (7640):191–196

15. Zarzeczny A, Scott C, Hyun I et al (2009) iPS cells: mapping the policy issues. Cell 139 (6):1032–1037 16. Pagliuca FW, Millman JR, Gurtler M et al (2014) Generation of functional human pancreatic beta cells in vitro. Cell 159(2):428–439 17. Rezania A, Bruin JE, Arora P et al (2014) Reversal of diabetes with insulin-producing cells derived in vitro from human pluripotent stem cells. Nat Biotechnol 32(11):1121–1133 18. Pepper AR, Pawlick R, Gala-Lopez B et al (2015) Diabetes is reversed in a murine model by marginal mass syngeneic islet transplantation using a subcutaneous cell pouch device. Transplantation 99(11):2294–2300 19. Ludwig B, Reichel A, Steffen A et al (2013) Transplantation of human islets without immunosuppression. Proc Natl Acad Sci U S A 110 (47):19054–19058 20. Krawczyk K, Xue S, Buchmann P et al (2020) Electrogenetic cellular insulin release for realtime glycemic control in type 1 diabetic mice. Science 368(6494):993–1001 21. Bochenek MA, Veiseh O, Vegas AJ et al (2018) Alginate encapsulation as long-term immune protection of allogeneic pancreatic islet cells transplanted into the omental bursa of macaques. Nat Biomed Eng 2(11):810–821 22. Jacobs-Tulleneers-Thevissen D, Chintinne M, Ling Z et al (2013) Sustained function of alginate-encapsulated human islet cell implants in the peritoneal cavity of mice leading to a pilot study in a type 1 diabetic patient. Diabetologia 56(7):1605–1614 23. de Vos P (2017) Historical perspectives and current challenges in cell microencapsulation. Methods Mol Biol 1479:3–21 24. Mount NM, Ward SJ, Kefalas P et al (2015) Cell-based therapy technology classifications and translational challenges. Philos Trans R Soc Lond Ser B Biol Sci 370(1680):20150017 25. Rostovskaya M, Bredenkamp N, Smith A (2015) Towards consistent generation of pancreatic lineage progenitors from human pluripotent stem cells. Philos Trans R Soc Lond Ser B Biol Sci 370(1680):20140365 26. Shao J, Xue S, Yu G et al (2017) Smartphonecontrolled optogenetically engineered cells enable semiautomatic glucose homeostasis in diabetic mice. Sci Transl Med 9(387):eaal2298 27. Yu Y, Wu X, Guan N et al (2020) Engineering a far-red light–activated split-Cas9 system for remote-controlled genome editing of internal organs and tumors. Sci Adv 6(28):eabb1777

Engineering Mammalian Cells to Control Glucose Homeostasis 28. Wroblewska L, Kitada T, Endo K et al (2015) Mammalian synthetic circuits with RNA binding proteins for RNA-only delivery. Nat Biotechnol 33(8):839–841 29. Lathuiliere A, Cosson S, Lutolf MP et al (2014) A high-capacity cell macroencapsulation system supporting the long-term survival of genetically engineered allogeneic cells. Biomaterials 35(2):779–791 30. Dolgin E (2014) Encapsulate this. Nat Med 20 (1):9–11 31. Vegas AJ, Veiseh O, Gurtler M et al (2016) Long-term glycemic control using polymerencapsulated human stem cell-derived beta cells in immune-competent mice. Nat Med 22 (3):306–311 32. de Vos P, Faas MM, Strand B et al (2006) Alginate-based microcapsules for immunoisolation of pancreatic islets. Biomaterials 27 (32):5603–5617 33. Hay CW, Docherty K (2003) Enhanced expression of a furin-cleavable proinsulin. J Mol Endocrinol 31(3):597–607 34. Kumagai-Braesch M, Jacobson S, Mori H et al (2013) The TheraCyte device protects against islet allograft rejection in immunized hosts. Cell Transplant 22(7):1137–1146 35. Ye H, Daoud-El Baba M, Peng RW et al (2011) A synthetic optogenetic transcription device enhances blood-glucose homeostasis in mice. Science 332(6037):1565–1568 36. Hernandez RM, Orive G, Murua A et al (2010) Microcapsules and microcarriers for in situ cell delivery. Adv Drug Deliv Rev 62 (7-8):711–730 37. Yin J, Yang L, Mou L et al (2019) A green tea-triggered genetic control system for treating diabetes in mice and monkeys. Sci Transl Med 11(515):eaav8826

57

38. Bojar D, Scheller L, Hamri GC et al (2018) Caffeine-inducible gene switches controlling experimental diabetes. Nat Commun 9 (1):2318 39. Ye H, Xie M, Xue S et al (2017) Self-adjusting synthetic gene circuit for correcting insulin resistance. Nat Biomed Eng 1(1):0005 40. Ariyachet C, Tovaglieri A, Xiang G et al (2016) Reprogrammed stomach tissue as a renewable source of functional beta cells for blood glucose regulation. Cell Stem Cell 18(3):410–421 41. Shapiro AM, Pokrywczynska M, Ricordi C (2017) Clinical pancreatic islet transplantation. Nat Rev Endocrinol 13(5):268–277 42. Zhu S, Russ HA, Wang X et al (2016) Human pancreatic beta-like cells converted from fibroblasts. Nat Commun 7:10080 43. Lipsitz YY, Bedford P, Davies AH et al (2017) Achieving efficient manufacturing and quality assurance through synthetic cell therapy design. Cell Stem Cell 20(1):13–17 44. Chambers AP, Sorrell JE, Haller A et al (2017) The role of pancreatic Preproglucagon in glucose homeostasis in mice. Cell Metab 25 (4):927–934. e923 45. Stanley SA, Gagner JE, Damanpour S et al (2012) Radio-wave heating of iron oxide nanoparticles can regulate plasma glucose in mice. Science 336(6081):604–608 46. Xue S, Yin J, Shao J et al (2017) A syntheticbiology-inspired therapeutic strategy for targeting and treating Hepatogenous diabetes. Mol Ther 25(2):443–455 47. Bai P, Liu Y, Xue S et al (2019) A fully human transgene switch to regulate therapeutic protein production by cooling sensation. Nat Med 25(8):1266–1273

Chapter 4 Using Engineered Mammalian Cells for an Epitope-Directed Antibody Affinity Maturation System Akihiro Eguchi and Masahiro Kawahara Abstract Antibodies have been attracting attention as therapeutic tools owing to their high affinity and specificity. To develop potent antibodies, affinity maturation, epitope regulation, and using target antigens in native form are pivotal requirements. Here we describe a method to conduct epitope-directed affinity maturation of antibodies using engineered mammalian cells. This method utilizes protein chimeras that transduce cell death signaling in response to antibody binding. As the competition of antibody binding inhibits the cell death signaling, only affinity-matured antibodies retaining the same epitope as an original one can be selected using cell survival as readout. Key words Antibody, Single-chain Fv, Epitope, Library screening, Receptor engineering, Membrane protein, Cell death signaling

1

Introduction Antibodies bind to specific targets with high affinity, so they have been widely used as therapeutic and analytical tools [1–3]. Upon developing antibodies against membrane proteins as therapeutics, their epitope is pivotal because the epitope determines the function of antibodies [4]. Furthermore, as the antibodies recognize their specific antigens expressed on the cell surface, the antigens used in selection should be displayed on mammalian cells in their native forms. As an antibody selection and affinity maturation system, a phage display method is the most common and sophisticated [5, 6]. However, this method does not meet the abovementioned demands for the epitope regulation and antigen display fashion. Here we develop engineered mammalian cells for epitopedirected antibody affinity maturation [7]. To realize epitopedirected affinity maturation, we focus on the binding competition of antibodies and design two chimeric proteins. The first one is a dimerizing antibody consisting of an original prematured antigen-specific single-chain Fv (scFv) clone and a mutant of

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

59

60

Akihiro Eguchi and Masahiro Kawahara

Dimerizing antibody

…

FKBPF36V:

…

AP20187 Original scFv: Antigen:

Antigen/Fas chimera

FKBPF36V:

Receptor Fas:

Death signaling

Fig. 1 Design of the antigen/Fas chimera and dimerizing antibody. The antigen/Fas chimera is a fusion protein consisting of ECD of a target antigen, TM of an erythropoietin receptor mutant, FKBPF36V, and ICD of a receptor Fas. The dimerizing antibody consists of an scFv against the antigen and FKBPF36V. AP20187 induces dimerization of the extracellular and intracellular FKBPF36V moieties independently and then oligomerization of the antigen/Fas chimera. ICD of Fas induces cell death signaling upon their oligomerization

FK506-binding protein 12 (FKBPF36V) whose dimerization is tunable by adding a chemical dimerizer AP20187 [8] (see Fig. 1). The other is named as an antigen/Fas chimera that is a fusion protein consisting of the extracellular domain (ECD) of a target antigen, the transmembrane domain (TM) of an erythropoietin receptor mutant, FKBPF36V, and the intracellular domain (ICD) of a death signaling receptor Fas. These two proteins are genetically encoded in retroviral vectors, with which cells are transduced to establish engineered cells. When the dimerizer AP20187 is added to the engineered cells expressing these two proteins, two FKBPF36V moieties cooperatively work to oligomerize the antigen/Fas chimera (see Fig. 1). As the receptor Fas induces cell death signaling by clustering [9], the receptor chimera induces death of the cells. However, when the cells additionally secrete another free scFv that binds to the same epitope as the original one with higher affinity, the free scFv blocks the binding of the dimerizing antibody and thus inhibits the cell death induction (see Fig. 2). Using an scFv library as a source of the free scFv, library screening for antibody affinity maturation can be conducted. After incubation of the engineered cells with the dimerizer AP20187, we can select affinity-matured scFvs only by harvesting the survived cells and recovering the integrated scFv sequences from their genome.

Epitope-Directed Antibody Affinity Maturation System …

61

…

Cell death Compete! secretion

+ Free scFv

Cell survival Fig. 2 Schematic illustration of the affinity maturation using cell survival as a readout. When AP20187 is added, the cells expressing two chimeric proteins, the antigen/Fas chimera and dimerizing antibody, die due to the activation of Fas (upward arrow). The binding of the dimerizing antibody and subsequent cell death signaling can be blocked by a free scFv secreted by the cells, if the free scFv has higher affinity and the same epitope as the original scFv (down arrow)

2

Materials All solutions are prepared using ultrapure water (18.2 MΩ/cm). Store enzymes, antibiotics, retronectin, AP20187, and DNA products at 20  C. Store competent cells at 80  C. Store IL-3, protamine sulfate, ethidium bromide (EtBr), and proprium iodide at 4  C. Store other reagents at room temperature.

2.1 Vector Construction

1. Host vectors (see Fig. 3): Three plasmids for retroviral transduction are necessary and these plasmids should code different antibiotic-resistance genes. In this chapter, we use a neomycinresistance gene for expressing the antigen-FKBPF36V-Fas chimera (see Fig. 3a, containing the illustration of the strategy of vector construction described in Subheading 3.1), a blasticidin-resistance gene for expressing scFv-FKBPF36V (see Fig. 3b), and a puromycin-resistance gene for expressing free scFv (see Fig. 3c). In addition, all plasmids code an ampicillinresistance gene for cloning in E. coli.

62

Akihiro Eguchi and Masahiro Kawahara pMK vector (plasmid DNA)

a:

Ψ

IRES

Stuffer

5’ LTR

FKBPF36V

+

NeomycinR

Fas ICD

3’ LTR

: PCR Primers for host.

Target antigen

: PCR Primers for insert. Black box is overlap sequence for DNA assembly

DNA assembly

Ψ

b:

IRES

Target antigen

5’ LTR

FKBPF36V

Fas ICD

Ψ 5’ LTR

c:

3’ LTR

BlasticidinR

3’ LTR

IRES

Leader

Original scFv

Leader

Free scFv

Ψ 5’ LTR

NeomycinR

FKBPF36V

IRES

T2A

PuromycinR

EGFP

3’ LTR

Fig. 3 Illustration of vectors for expressing proteins and the way to construct the vectors. (a) Plasmid DNA coding the antigen/Fas chimera. Stuffer (dotted rectangle) should be replaced by the gene coding the target antigen. Normal arrows represent primers for PCR of the host plasmid, whereas arrows with a black rectangle represent those for PCR of the target antigen. The black rectangles represent 15-bp overlap sequences for DNA assembly reactions. (b) Plasmid DNA coding the dimerizing antibody. (c) Plasmid DNA coding the free scFv

2. Target gene: Genes coding a target membrane protein and an original scFv. 3. Reagents for PCR: DNA polymerase, primers, and dNTPs. 4. Thermal cycler. 5. Restriction enzymes (PstI and ClaI are used for library vector construction). 6. DpnI enzyme. 7. Enzyme for DNA assembly. 8. 50 TAE buffer: 2 M Tris, 1 M acetic acid, 50 mM EDTA. Weigh 242 g of Tris base and 18.6 g of EDTA-2Na. Dissolve them in 800 mL of water, add 60.5 mL of acetic acid, and adjust volume to 1 L with additional water.

Epitope-Directed Antibody Affinity Maturation System

63

9. Agarose gel: Weigh 1.0 g of agarose and add it in 100 mL of 1 TAE buffer. Dissolve it using a microwave, add EtBr solution (final concentration: 0.5 μg/mL), and pour the mixed solution into a gel plate (see Note 1). 10. Electrophoresis chamber and gel plate for agarose gel preparation. 11. Loading dye for agarose gel electrophoresis. 12. UV irradiator (see Note 2). 13. Gel extraction kit. 14. Spectrophotometer. 15. Chemical competent cell. 16. Autoclave. 17. Test tubes for small-scale culture (5 mL). 18. 37  C incubator. 19. 37  C shaker (horizontal shaking at 200 rpm for culture in test tubes). 20. LB agar plate: Weigh 1.0 g of tryptone, 0.5 g of yeast extract, 1.0 g of NaCl, and 1.5 g of agar. Add them to 100 mL of water in a flask and autoclave the solution. Add antibiotics (100 μL of 50 mg/mL ampicillin) after autoclaving and pour the solution to 5–6 dishes of 10 cm dishes (see Note 3). 21. LB medium: Weigh 1.0 g of tryptone, 0.5 g of yeast extract, and 1.0 g of NaCl. Add them to 100 mL of water, dispense 5 mL of the solution into each test tube, and autoclave it. 22. High-speed centrifuge. 23. Kit for plasmid DNA mini-preparation. 2.2 Establishment of the Engineered Cells

1. Clean bench and safety cabinet. 2. CO2 incubator. 3. Centrifuge. 4. Autoclave. 5. Cell culture dishes and plates: 6 cm dishes, 10 cm dishes, 96-well plates, 24-well plates, and 6-well plates. Medium volume is adjusted to 5 mL for 6 cm dishes, 10 mL for 10 cm dishes, 200 μL for 96-well plates, 1 mL for 24-well plates, and 3 mL for 6-well plates. 6. Cell culture: A murine pro-B Ba/F3 cell line is cultured in RPMI1640 medium containing 10% fetal bovine serum (FBS), 30 μg/mL kanamycin, penicillin–streptomycin, and 1 ng/mL murine IL-3, with 37  C/5% CO2 atmosphere in an incubator. A viral packaging cell line Plat-E is cultured in Dulbecco’s modified Eagle’s medium (DMEM) containing

64

Akihiro Eguchi and Masahiro Kawahara

10% FBS, 30 μg/mL kanamycin, penicillin–streptomycin, 1 μg/mL puromycin, and 10 μg/mL blasticidin with 37  C/ 10% CO2 atmosphere in an incubator. 7. 0.05% Trypsin–EDTA. 8. Reagents for lipofection. 9. Dulbecco’s phosphate-buffered saline (DPBS): Weigh 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4·2H2O, and 0.2 g of KH2PO4. Dissolve them in 800 mL of water, adjust volume to 1 L with additional water, and autoclave the solution. 10. 2% BSA/DPBS: Weigh 0.2 g of bovine serum albumin (BSA) and dissolve it in 10 mL of DPBS. Filter the solution using a 0.22 μm PVDF filter. 11. Retronectin. 12. 0.22 μm PVDF filter. 13. 10 mL Syringe. 14. Antibiotics: puromycin, blasticidin, and G418 (neomycin). 15. Propidium Iodide. 16. Flow Cytometer. 2.3 Library Preparation and Screening

1. 3 M Sodium acetate (pH 5.2). 2. Ethanol (100% and 70% in water). 3. Gel extraction kit. 4. DNA ligase (e.g., Ligation high). 5. Electrocompetent cells. 6. Recovery medium (the product recommended by the manufacture of electrocompetent cells). 7. Electroporator. 8. Cuvette. 9. Autoclave. 10. Test tubes for small scale culture. 11. 500 mL baffled flask 12. 37  C incubator. 13. 37  C shaker which enables reciprocal shaking at 200 rpm for culture in test tubes and orbital shaking at 110 rpm for culture in large plates and flasks. 14. LB agar large plate: The same preparation as 2.1.20 except the final solution is poured into a large plate (e.g., a 245 mm  245 mm plate). 15. LB medium: The same preparation as Subheading 2.1, item 21, but without dispensing into 5 mL test tubes.

Epitope-Directed Antibody Affinity Maturation System

65

16. Kit for plasmid DNA midi-preparation. 17. High-speed centrifuge. 18. Protamine sulfate. 19. Chemical dimerizer AP20187.

3

Methods

3.1 Vector Construction

1. Design primers to amplify target genes following the guideline of final plasmid products (see Fig. 3). Basically, a 15-bp overlap sequence should be included in primers for successful DNA assembly reactions in the following step 6. 2. Conduct PCR. Mix DNA polymerase, dNTPs, primers, and template DNA following the guideline of polymerase you use. 3. Add 1 μL of DpnI enzyme (20,000 U/mL) to digest the template DNA, and incubate the mixture for 1 h at 37  C. 4. Mix the product with loading dye, apply the mixture to 1% agarose gel and run electrophoresis at 150 V for 20 min, and check whether target DNA is properly amplified based on the molecular weight of the amplicon using a UV irradiator. 5. Cut the amplified target DNA out of the gel under UV irradiation, and purify the DNA using gel extraction kit. 6. Measure the concentration of the purified DNA by A260 absorbance using a spectrophotometer. Mix 50 ng of the host fragment and threefold amount of the insert fragment, and add water up to 4 μL. Add 1 μL of a 5  In-fusion HD enzyme and incubate the reaction mixture for 15 min at 50  C. 7. Transform 50 μL of chemical competent cells with 5 μL of the reaction mixture by incubation for 5 min on ice (see Note 4). 8. Heat-shock the cells for 30 s at 42  C, and return them on ice. 9. Add 200 μL of LB medium, and seed it all onto an LB plate (see Note 5). 10. Incubate it overnight at 37  C (see Note 6). 11. Pick up several colonies to test tubes with 5 mL of LB medium, add 50 μg/mL ampicillin, and incubate them for 12–16 h at 37  C, shaking at 200 rpm. 12. Centrifuge the suspension at 13,000  g for 1 min, and harvest the cells. 13. Extract plasmids from the cell pellets using a miniprep kit. 14. Confirm the sequence of the plasmids and use a correct one in the following experiments.

66

Akihiro Eguchi and Masahiro Kawahara

3.2 Transduction of Vectors Coding the Antigen/Fas Chimera and Dimerizing Antibody to Ba/F3 Cells (See Note 7)

Day

1: Seeding

1. Maintain Plat-E cells in DMEM supplemented with puromycin and blasticidin in 10 cm cell culture dishes at 37  C, 10% CO2. 2. Remove the medium and wash the cells by DPBS. Add 1 mL of 0.05% trypsin–EDTA and incubate the cells at 37  C until the cells are almost detached. 3. Resuspend the cells by 9 mL of DMEM and seed 1.0  106 cells/well in a 6-well plate and incubate the cells at 37  C, 10% CO2. Day 0: Transfection 4. Mix 3 μg of plasmids and 6 μL of Plus reagent in 300 μL OptiMEM. 5. Mix 6 μL of Lipofectamine LTX or Lipofectamine 3000 in 300 μL Opti-MEM. 6. Mix the solution of steps 4 and 5, and incubate it for 15 min at room temperature. 7. Add the solution to the cells cultured in step 3, and incubate the cells at 37  C, 10% CO2. Two plasmids coding the antigen/ Fas chimera or dimerizing antibody should be separately transfected. Day 1: Medium replacement and preparation of a retronectincoated plate 8. Twenty-four hours after transfection, remove the culture medium and add 2 mL of fresh medium. 9. Add 100 μL of 20 μg/mL retronectin solution to each well of a nontreated 24-well plate, and leave the plate for 2 h at room temperature. 10. Remove the solution, add 500 μL of 2% BSA/DPBS to each well, and incubate for 30 min at room temperature. 11. Remove 2% BSA/DPBS and wash the wells by DPBS. 12. Store the plate at 4  C. Day 2: Transduction of Ba/F3 cells 13. Forty-eight hours after transfection, harvest the cell culture supernatant, filter it through a 0.22 μm PVDF filter, mix the medium from two wells in which plasmids coding two chimeric proteins were separately transfected, transfer the filtrate to the retronectin-coated plate, and incubate the plate for 4–6 h at 37  C in 5% CO2. 14. Remove the suspension and wash the wells by DPBS.

Epitope-Directed Antibody Affinity Maturation System

67

15. Add 5.0  104 Ba/F3 cells suspended in RPMI1640 with 1 ng/mL IL-3 to each well, and incubate the plate at 37  C in 5% CO2. Day 4: Harvest Ba/F3 cells 16. Remove the medium and wash the wells by DPBS. 17. Add 200 μL of 0.05% trypsin–EDTA to each well and incubate at 37  C until the cells are almost detached. 18. Resuspend the cells in each well by RPMI1640 with 1 ng/mL IL-3, and transfer the cell suspension to a 6 cm dish. Day 5~: Antibiotic selection 19. Seed transductants and nontransduced Ba/F3 cells separately in a 24-well plate at 1.0  105 cells/well in RPMI1640 with 1 ng/mL IL-3 and corresponding antibiotics. We recommend the following antibiotics concentration: 10 μg/mL for blasticidin and 800 μg/mL for G418 (Geneticin), the latter of which is the antibiotics targeted by the neomycinresistance gene. 20. After nontransduced Ba/F3 cells completely die, collect transductants and maintain them. 3.3 Cell Cloning (See Note 8)

1. Serially dilute transductants to 800 cells/mL by RPMI1640. 2. Dilute parental Ba/F3 cells to 1.6  105 cells/mL by RPMI1640. 3. Add 1 mL each of the cell suspensions prepared in steps 1 and 2 to 18 mL RPMI1640. The final cell densities of transductants and parental Ba/F3 cells become 4 cells/mL and 8.0  104 cells/mL, respectively. 4. Add 1 ng/mL IL-3 and corresponding antibiotics to the cell suspension. 5. Seed the cell suspension to two 96-well plates at 200 μL/well. 6. After 1-week incubation, choose single colonies and scale up them (see Note 8).

3.4 Evaluation of Cloned Cells Expressing the Antigen/Fas Chimera and Dimerizing Antibody (See Note 7)

1. Prepare 500 μL of RMPI1640 medium containing 200 nM AP20187 and 2 ng/mL IL-3 in a 24-well plate. 2. Centrifuge the cloned cells at 380  g for 3 min, discard the supernatant, and wash the cells by DPBS once. 3. Resuspend the cells by RPMI1640 medium and measure the cell density. 4. Adjust the cell density at 1.0  105 cells/mL by RPMI1640 medium, and add 500 μL of the cell suspension to the 24-well plate.

68

Akihiro Eguchi and Masahiro Kawahara

5. Incubate the cells for 2 days. 6. Prepare a mixture containing 45.85 μL of DPBS, 4 μL of Flow Count Fluorosphere, and 0.15 μL of 1 mg/mL propidium Iodide. 7. Mix 100 μL of the cell suspension and 50 μL of the mixture prepared in step 6. 8. Measure the number of survived and dead cells using flow cytometry (see Note 9). 9. Compare the efficiency of cell death induction and use the clone showing the highest cell death rate for the following experiments. 3.5 Preparation of a Plasmid Coding an scFv Library

1. Prepare primers that amplify a library sequence of interest (see Note 10). 2. Conduct PCR. In order to prepare a DNA fragment containing two restriction enzyme recognition sites, PCR can be conducted for several times using extended primers (see Fig. 4). In that case, the PCR product should be purified by agarose gel extraction as shown in Subheading 3.1, steps 4–6. 3. Digest 3 μg of the host vector and 3 μg of the purified PCR product by a pair of restriction enzymes (PstI and ClaI) for 1 h at 37  C (see Note 11). 4. Purify the products by agarose gel extraction as shown in Subheading 3.1, steps 4–6. 5. Mix 1 μg each of the host and insert fragments. 6. Conduct ethanol precipitation. Add 1/10 volume of sodium acetate, add 2.5 times volume of ethanol, incubate the mixture for 20 min at room temperature, centrifuge the mixture for 20 min at 21,500  g, aspirate the supernatant, add the same volume of 70% ethanol, centrifuge it for 20 min at 21,500  g, aspirate the supernatant, and dry the pellet until residual ethanol is evaporated completely. 7. Dissolve the pellet by 5 μL of water, add 5 μL of Ligation high, and incubate the mixture for 4 h at room temperature. 8. Conduct ethanol precipitation again, dissolve the pellet by water, and adjust the DNA concentration to 100 ng/μL. 9. Take 2.5 μL of the vector (100 ng/μL) into an ice-cold microtube. 10. Add 50 μL of electrocompetent cells gently to the microtube, and stir slowly by pipetting (see Note 12). 11. Transfer the mixture to a cuvette and pulse at the condition of 2000 V, 25 μF, 200 Ω, 1 ms (see Note 13).

Epitope-Directed Antibody Affinity Maturation System

Ψ 5’ LTR

IRES

Leader

Free scFv

69

T2A

PuromycinR

EGFP

3’ LTR

Library

Free scFv PstI site

ClaI site

PstI and ClaI Digestion

PCR and Digestion Library

Ligation scFv Library

Ψ 5’ LTR

IRES

Leader

T2A

PuromycinR

EGFP

3’ LTR

Fig. 4 Illustration of the way to construct the vector coding the scFv library. As the scFv sequence is flanked by PstI and ClaI recognition sites unique in the host vector, these two restriction enzymes are used for excision. A primer containing degenerate NNS codons is used for constructing a library (arrow with a white rectangle) and another round of PCR is conducted so that the amplicon is flanked by the recognition sites of the two restriction enzymes (normal arrow). The library vector can be obtained by the ligation of the digested amplicon and host vector

12. Immediately add 200 μL of recovery medium and incubate for 1 h at 37  C. 13. Seed the cells to an LB large plate (see Note 14). 14. After overnight incubation, add 40 mL of LB medium with ampicillin to the LB large plate, and shake the plate at 110 rpm for 1 h on the 37  C shaker. 15. Collect the medium to a culture flask, and add new LB medium up to 125 mL. 16. Shake the flask at 220 rpm at 37  C until OD600 becomes higher than 0.7. 17. Collect the cells by centrifugation at 5000  g for 15 min. 18. Extract the plasmid using a midiprep kit.

70

Akihiro Eguchi and Masahiro Kawahara

3.6 Library Screening (See Note 7)

Day

1 to Day 1

1. Produce retroviral vectors as described in steps 1–8 of Subheading 3.2 except the following. Scale up the amount of DNA from 3 μg to 10 μg. Mix DNA, 3.75 μL of Lipofectamine, and 5 μL of Plus reagent in 300 μL of Opti-MEM. Day 2 2. Collect the retroviral supernatant and dilute it in half by RPMI1640. 3. Prepare 6.0  106 cloned Ba/F3 cells expressing the antigen/ Fas chimera and dimerizing antibody in a 15 mL tube. 4. Centrifuge it at 380  g for 3 min, discard the supernatant, and wash the cells by DPBS. 5. Resuspend the cells by the solution of step 2, add 1 ng/mL IL-3 and 10 μg/mL protamine sulfate, and seed the mixture in a 10 cm dish. 6. Incubate the cells for 6 h in 37  C, 5% CO2 condition. 7. Dispense the cell suspension into three 10 cm dishes. 8. After 2 days incubation, check the efficiency of transduction by measuring a GFP-positive rate using flow cytometry (see Note 15). Using protamine sulfate, transduction efficiency would be generally smaller than 10%. Day 3~ 9. Seed the cells to 96-well plates at the density of 5.0  104 cells/ mL with 100 nM AP20187 and 1 μg/mL puromycin. 10. After 7 days incubation, scale up the grown cells to 24-well plates, and finally to 6-well plates. 11. Collect the cells and extract the genomes using a genome extraction kit. 12. Amplify the scFv region of the genome by PCR and read the sequence to obtain the information of selected scFvs.

4

Notes 1. EtBr is a carcinogen, so should not handle it with bare hands. Gel and buffer containing EtBr should be discarded properly. Buffer should be mixed with activated carbon to remove EtBr from the solution. Alternative noncarcinogenic reagents can be used. 2. Do not directly see the UV light. Wear UV-protecting glasses.

Epitope-Directed Antibody Affinity Maturation System

71

3. To avoid inactivation by heat, antibiotics should be added after cooling down the flask until we can keep touching. Pour the solution to dishes as soon as possible after adding antibiotics to avoid gelling. 4. Chemical competent cells are vulnerable to heat. Be careful to keep cells always on ice. 5. The LB gel plate is vulnerable, so spread the solution gently using a bacterial spreader. 6. The LB gel plate should be stored upside down in order not to accumulate water droplets on the lid and fall them on the plate. 7. All steps should be conducted in a clean bench or in a safety cabinet in a biosafety level-2 room for retroviral transduction. Wipe instruments and reagents with 70% ethanol before bringing them into the bench or cabinet. 8. As there may be several colonies in one well, check the colony formation status everyday by microscopy. Additionally, check whether the cells are derived from single colonies by measuring the surface expression level of the antigen/Fas chimera by immunostaining. 9. Cell death rate is calculated by a ratio of propidium iodidestained cells to all cells. 10. The following types of scFv libraries can be applied; naı¨ve or animal-immunized libraries; artificially randomized libraries created by error-prone PCR or site-directed mutagenesis using degenerate primers (e.g., those containing degenerate NNS codons). Here, the strategy using degenerate primers is introduced. 11. Because electroporation necessitates relatively concentrated DNA, and because multiple DNA purification steps result in DNA loss, the PCR and ligation reactions should be scaled up. The product should be purified by ethanol precipitation. Other purification methods such as those using AMPure can be used alternatively. DNA assembly may be used for vector construction instead of ligation. 12. Electrocompetent cells are also vulnerable to heat. Be careful to keep cells always on ice. Do not pipette cells. Tubes, tips, and cuvettes should be cooled on ice in advance. 13. Before electroporation, confirm that the cell suspension covers the bottom of the cuvette, and then wipe out the ice-derived water droplets outside of the cuvette. Take care not to touch the instrument upon electroporation. 14. To correctly estimate the colony number appeared on the large plate, make dilution series using several normal-sized plates. Before seeding the cells to the large plate, prepare 103 to 105-

72

Akihiro Eguchi and Masahiro Kawahara

fold dilution series and seed them to the normal-sized plates. Thus, the number of colonies in the large plate can be roughly calculated from the number of colonies in the normal-sized plates. 15. The GFP positive rate should be smaller than 10% to decrease the cells undergoing integration of multiple scFv genes. References 1. Scott AM, Wolchok JD, Old LJ (2012) Antibody therapy of cancer. Nat Rev Cancer 12:278–287 2. Ibrahim AD (2006) Immunoassay methods and their applications in pharmaceutical analysis: basic methodology and recent advances. Int J Biomed Sci 2:217–235 3. June CH, Sadelain M (2018) Chimeric antigen receptor therapy. N Engl J Med 379:64–73 4. Tvorogov D, Anisimov A, Zheng W, Lepp€anen VM, Tammela T, Laurinavicius S, Holnthoner W, Heloter€a H, Holopainen T, Jeltsch M, Kalkkinen N, Lankinen H, Ojala PM, Alitalo K (2010) Effective suppression of vascular network formation by combination of antibodies blocking VEGFR ligand binding and receptor dimerization. Cancer Cell 18:630–640 5. Smith GP (1985) Filamentous fusion phage: novel expression vectors that display cloned antigens on the virion surface. Science 228:1315–1317

6. McCafferty J, Griffiths AD, Winter G, Chiswell DJ (1990) Phage antibodies: filamentous phage displaying antibody variable domains. Nature 348:552–554 7. Eguchi A, Nakakido M, Nagatoishi S, Kuroda D, Tsumoto K, Nagamune T, Kawahara M (2019) An epitope-directed antibody affinity maturation system utilizing mammalian cell survival as readout. Biotechnol Bioeng 116:1742–1751 8. Clackson T, Yang W, Rozamus LW, Hatada M, Amara JF, Rollins CT, Stevenson LF, Magari SR, Wood SA, Courage NL, Lu X, Cerasoli F Jr, Gilman M, Holt DA (2006) Redesigning an FKBP-ligand interface to generate chemical dimerizers with novel specificity. Proc Natl Acad Sci U S A 95:10437–10442 9. Kaufmann T, Strasser A, Jost PJ (2011) Fas death receptor signalling: roles of Bid and XIAP. Cell Death Differ 19:42–50

Chapter 5 Purification of Specific Cell Populations Differentiated from Stem Cells Using MicroRNA-Responsive Synthetic Messenger RNAs Hideyuki Nakanishi and Hirohide Saito Abstract Pluripotent stem cells have the potential to differentiate into various cell types that can be used for basic biological studies, drug discovery, and regenerative medicine. To obtain reliable results using the differentiated cells, the contamination of nontarget cells should be avoided. microRNAs (miRNAs) can serve as indicators to distinguish target and nontarget cells, because the activities of miRNAs are different among cell types. In this chapter, we introduce a method to purify target cells using synthetic messenger RNAs (mRNAs) that respond to cell-specific miRNAs. The method is composed of five steps: mRNA sequence design, template DNA preparation by PCR, in vitro mRNA transcription, mRNA transfection into cells, and fluorescence-activated cell sorting. This synthetic mRNA-based cell purification method will advance various applications of pluripotent stem cells. Key words Cell sorting, microRNA, Stem cell, Differentiation, Synthetic mRNA, Synthetic biology, Regenerative medicine

1

Introduction Pluripotent stem cells have the potential to differentiate into various cell types that can be used for both biological studies and biomedical applications including regenerative medicine and drug discovery. However, after the differentiation, stem cell-derived populations sometimes contain undifferentiated cells or other nontarget cell types. Such unwanted cells can confound drug discovery or cause adverse effects such as tumorigeneses in regenerative medicine. Thus, the purification of target cell types is critical for reliable studies and safe clinical applications. Accordingly, in this chapter, we describe a method to purify target cell types based on differences in microRNA (miRNA) activity.

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

73

74

Hideyuki Nakanishi and Hirohide Saito

miRNAs are small (approximately 22 nt) noncoding RNAs that repress the translation of their target messenger RNAs (mRNAs). There are various miRNAs (e.g., more than 2600 mature miRNAs are discovered in human cells [1]) and each has a unique sequences. To repress translation, a miRNA binds directly to an mRNA via a complementary sequence. Importantly, miRNA activity depends on the cell type. For example, the activity of hsa-miR-302a-5p, one of the human miRNAs, is high in human pluripotent stem cells (embryonic stem (ES) and induced pluripotent stem (iPS) cells), but low in HeLa cells and midbrain dopaminergic neuronal cells [2] [3]. Thus, miRNA activity can be an indicator to distinguish cell types. To purify living specific cell types based on miRNA activity, we have developed a miRNA-responsive synthetic reporter mRNA system [4]. Because these reporter mRNAs contain complementary miRNA sequences, the reporter gene expression is repressed if the target miRNA activity is high in cells transfected with the system. Thus, we can purify cells by fluorescence-activated cell sorting (FACS) (Fig. 1). In this chapter, we describe the procedure for cell purification using these miRNA-responsive reporter mRNAs from the mRNA sequence design to cell sorting.

2

Materials

2.1 Template DNA Preparation by PCR

1. pDNAs coding fluorescent proteins (e.g., tagRFP, EGFP). 2. Nuclease-free water. 3. High fidelity PCR enzymes (e.g., KOD -Plus- Neo (Toyobo)). 4. The PCR primers listed below. Sequences described as “NNN. . .. . .NNN” depend on the fluorescent protein-coding genes and miRNAs to be detected. Details are shown in Subheading 3.2. 1. Primer set for miRNA responsive reporter ORF PCR Forward primer-1

50 -AGAAAAGAAGAGTAAGAAGAAATATAAGACACCG GTCNNN. . .. . .NNNGCCACCATGNNN. . .. . .NNN-30

Reverse primer-1

50 -GCCCCGCAGAAGGTCTAGATTCANNN. . .. . .NNN-30

2. Primer set for reference reporter ORF PCR Forward primer- 50 -CACCGGTCGCCACCATGNNN. . .. . .NNN-30 2 Reverse primer- 50 -GCCCCGCAGAAGGTCTAGATTCANNN 2 . . .. . .NNN-30

Cell Purification by MicroRNA-Responsive Synthetic mRNA

75

Fig. 1 An overview of the cell purification using miRNA-responsive reporter mRNAs. miRNA-responsive and reference reporter mRNAs are synthesized by in vitro transcription. Both reporter mRNAs are transfected into unpurified cells, followed by fluorescence-activated cell sorting (FACS). Target and nontarget cells can be distinguished and sorted based on the fluorescence ratio of the miRNA-responsive and reference reporters (i.e., the ratio is low in cells with high miRNA activity)

76

Hideyuki Nakanishi and Hirohide Saito 3. Primer set for 30 UTR PCR Forward primer-3

50 -TCTAGACCTTCTGCGGGGC-30

Reverse primer-3

50 -TTTTTTTTTTTTTTTTTTTTCCTACTCAGGCTTTATTC AAAGACCAAG-30

30 UTR template DNA

50 -TCTAGACCTTCTGCGGGGCTTGCCTTCTGGCCATG CCCTTCTTCTCTCCCTTGCACCTGTACCTCTTGGTC TTTGAATAAAGCCTGAGTAGG-30

4. Primer set for miRNA responsive reporter Full PCR Forward primer-4

50 -CAGTGAATTGTAATACGACTCACTATAGGGCGAATTA AGAGAGAAAAGAAGAGTAAGAAGAAATATAAGACACC-30

Reverse primer-4

50 -TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTCCTACTCAGGCTTTATTCA-30

5. Primer set for reference reporter Full PCR Forward primer-5

50 -CAGTGAATTGTAATACGACTCACTATAGGGCGA-30

Reverse primer-4

50 -TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTTTTTTTTTTTTTTTTTTTTTTTCCTACTCAGGCTTTATT CA-30

50 UTR template DNA

50 -CAGTGAATTGTAATACGACTCACTATAGGGCGAATTAAGAGAG AAAAGAAGAGTAAGAAGAAATATAAGACACCGGTCGCCACCA TG-30

5. Thermal cycler. 6. DpnI. 7. Agarose. 8. Tris-Acetate-EDTA (TAE) Buffer. 9. Microwave oven. 10. Electrophoresis apparatus for agarose gels. 11. DNA ladder. 12. Loading dye for agarose gel electrophoresis. 13. Nucleic acids staining reagent. 14. Gel imager. 15. PCR product purification kit. 16. Centrifuge. 17. Spectrophotometer.

Cell Purification by MicroRNA-Responsive Synthetic mRNA

2.2 mRNA Preparation by In Vitro Transcription

77

1. T7 transcription kit (e.g., MEGAscript T7 Transcription Kit (ThermoFisher Scientific)). 2. Cap analog (e.g., Anti-Reverse Cap Analog (TriLink)). 3. Modified NTP (e.g., 0 -triphosphate (TriLink)).

N1-methylpseudouridine-5-

4. RNA purification kit. 5. Alkaline phosphatase. 6. Electrophoresis apparatus to check RNA size and quality. 2.3

Cell Culture

1. Cells of interest. 2. Medium and culture plate suitable for the cells. 3. Suitable reagent to detach the cells.

2.4 mRNA Transfection

1. Transfection reagent.

2.5

1. Cell strainer.

Flow Cytometry

2. Serum free medium.

2. Flow cytometer. 3. Tubes to collect sorted cells.

3

Methods

3.1 Choice of Appropriate Fluorescent Proteins

3.2 Design of Primers for ORF PCR

1. Check the laser-filter sets of the flow cytometer to be used. At least two different laser-filter sets are necessary. 2. Search for fluorescent proteins with excitation and emission wavelength peaks close to the laser and filter wavelengths, respectively. For example, if the wavelength of the laser is 561 nm and that of the filter is 582/15 nm, examples of suitable fluorescent proteins are mKO2 (excitation: 551 nm, emission: 565 nm), DsRed (excitation: 558 nm, emission: 583 nm), and tagRFP (excitation: 555 nm, emission: 584 nm). Although dimeric fluorescent proteins can also be used, for a clear separation of miRNA-positive and -negative cells, we recommend using monomeric fluorescent proteins. At least two fluorescent proteins that can be detected by different laser-filter sets (e.g., Azami Green and tagRFP) are needed. A fluorescent protein database such as FPbase (https://www. fpbase.org/) [5] may be helpful for finding suitable proteins. Template DNAs for the in vitro transcription are prepared by two rounds of PCR. In the first round, the fluorescent protein gene and 30 UTR are amplified (ORF PCR and 30 UTR PCR, respectively). In the second round, these two DNA fragments are fused, and T7 promoter and poly-A sequences are added by fusion PCR (Fig. 2).

78

Hideyuki Nakanishi and Hirohide Saito

Fig. 2 Scheme for preparing template DNAs for in vitro transcription. Template DNAs for in vitro transcription are obtained by two rounds of PCRs. In the first round (ORF PCR), the partial 50 and 30 UTRs are added to the coding region. Template DNAs containing all necessary sequences for the in vitro transcription are obtained in the second round (Full PCR)

In this section, we describe the procedure to design the primers used in ORF PCR. The sequences of other primers are described in Subheading 2.1. 1. Design the forward and reverse primers to amplify the fluorescent protein genes to be used as a miRNA-responsive and a reference reporter, respectively. If the fluorescent protein gene in the template pDNA is adjacent to a Kozak sequence and a stop codon, then these sequences should also be included. Primer design tools such as Primer3Plus (http://primer3plus. com/cgi-bin/dev/primer3plus.cgi) may be helpful for designing the primers. 2. Add the partial 50 UTR sequence (50 -AGAAAAGAAGAGTAA GAAGAAATATAAGACACCGGTC-30 for miRNA-responsive reporters and 50 -CACCGGTC-30 for reference reporters) to the 50 end of the forward primers. If the fluorescent protein gene in the template pDNA is not adjacent to a Kozak sequence, then add the Kozak sequence (Fig. 3). In the case of a miRNA-responsive reporter, complementary sequences of the miRNA to be detected should also be added. Sequences of miRNAs to be detected can be obtained from miRbase (http://www.mirbase.org) [1] (see Note 1). For example, when hsa-miR-21-5p (50 -UAGCUUAUCA GACUGAUGUUGA -30 ) is the target and human codonoptimized monomeric Azami Green is the reporter, the primer sequence shown below can be used (the complementary sequence of hsa-miR-21-5p is underlined).

Cell Purification by MicroRNA-Responsive Synthetic mRNA

79

Fig. 3 Primer design for ORF PCR. The partial 50 and 30 UTR sequences are added to the forward and reverse primers, respectively. In the case of a miRNA-responsive reporter, a miRNA target site (complementary sequence of miRNA) is also added

50 - AGAAAAGAAGAGTAAGAAGAAATATAAGACACC GGTCTCAACATCAGTCTGATAAGCTAGCCACCATGGT GAGCGTGATCAAGCCCGAGA-30 3. Add the partial 30 UTR sequence (50 -GCCCCGCAGAAGGTC TAGAT-30 ) to the 50 end of the reverse primers. If the fluorescent protein gene in the template pDNA is not adjacent to a stop codon, then add a stop codon (Fig. 3). For example, when hmAG1 is used as the reporter, the primer sequence shown below can be used. 50 - GCCCCGCAGAAGGTCTAGATTCACTTGGCCTG GCTGGGC-30 3.3 Template DNA Preparation by PCR

1. Mix the components for the ORF PCR and 30 UTR PCR, respectively. Examples of mixtures using KOD -Plus- Neo are shown below. The sequences of each primer are described in Subheading 2.1. ORF PCR for miRNA-responsive reporter Component

Volume

Final concentration

10 PCR buffer for KOD -Plus- Neo

5μl

1

dNTPs (2 mM)

5μl

0.2 mM each

MgSO4 (25 mM)

3μl

1.5 mM

Forward primer-1 (10μM)

1.5μl

0.3μM

Reverse primer-1 (10μM)

1.5μl

0.3μM

Template pDNA (50 ng/μl)

0.1–1μl

0.1–1 ng/μl

KOD -Plus- Neo (1 U/μl)

1μl

0.02 U/μl

dH2O

Up to 50μl

80

Hideyuki Nakanishi and Hirohide Saito

ORF PCR for reference reporter Component

Volume

Final concentration

10 PCR buffer for KOD -Plus- Neo

5μl

1

dNTPs (2 mM)

5μl

0.2 mM each

MgSO4 (25 mM)

3μl

1.5 mM

Forward primer-2 (10μM)

1.5μl

0.3μM

Reverse primer-2 (10μM)

1.5μl

0.3μM

Template pDNA (50 ng/μl)

0.1–1μl

0.1–1 ng/μl

KOD -Plus- Neo (1 U/μl)

1μl

0.02 U/μl

dH2O

Up to 50μl

30 UTR PCR Component

Volume

Final concentration

10 PCR buffer for KOD -Plus- Neo

5μl

1

dNTPs (2 mM)

5μl

0.2 mM each

MgSO4 (25 mM)

3μl

1.5 mM

Forward primer-3 (10μM)

1.5μl

0.3μM

Reverse primer-3 (10μM)

1.5μl

0.3μM

30 UTR template DNA (10 nM)

1.5μl

0.3 nM

KOD -Plus- Neo (1 U/μl)

1μl

0.02 U/μl

dH2O

31.5μl

2. According to the manufacturer’s instructions, set a thermal cycler to perform the ORF and 30 UTR PCR. An example using KOD -Plus- Neo is described below. Predenature: 94  C, 2 min. Denature: 98  C, 10 s. Extension: 68  C, 30 s/kb. (Repeat the denature and extension steps 19 times (total 20 cycles).) 3. Add 1μl of DpnI (3–20 U/μl) to the PCR solution and incubate at 37  C for 0.5–1 h to remove the template pDNA. If the template pDNA lacks a T7 promoter sequence, this step can be ignored.

Cell Purification by MicroRNA-Responsive Synthetic mRNA

81

4. Perform the electrophoresis of a small portion of the PCR solution (e.g., 5μl) using 1.2% agarose gel to check the size of the amplified DNA (100 V, 25 min). Note that the sufficient amount of PCR solution for detection depends on the sensitivity of the nucleic acid staining reagent and the gel imager. 5. If the size of the amplified DNA is appropriate, purify it from the residual PCR solution by using a PCR product purification kit. 6. Quantify the concentration of purified DNA by Nanodrop or a comparable spectrophotometer. 7. Mix the components for the Full PCR, respectively. Examples of mixtures using KOD -Plus- Neo is shown below. Full PCR for miRNA-responsive reporter Component

Volume

Final concentration

10 PCR buffer for KOD -Plus- Neo

5μl

1

dNTPs (2 mM)

5μl

0.2 mM each

MgSO4 (25 mM)

3μl

1.5 mM

Forward primer-4 (10μM)

1.5μl

0.3μM

Reverse primer-4 (10μM)

1.5μl

0.3μM

3 UTR PCR product (500 nM)

1μl

10 nM

ORF PCR product (50 ng/μl)

0.1–1μl

0.1–1 ng/μl

KOD -Plus- Neo (1 U/μl)

1μl

0.02 U/μl

dH2O

Up to 50μl

0

Full PCR for reference reporter Component

Volume

Final concentration

10 PCR buffer for KOD -Plus- Neo

5μl

1

dNTPs (2 mM)

5μl

0.2 mM each

MgSO4 (25 mM)

3μl

1.5 mM

Forward primer-5 (10μM)

1.5μl

0.3μM

Reverse primer-4 (10μM)

1.5μl

0.3μM

5 UTR template DNA (500 nM)

1μl

10 nM

30 UTR PCR product (500 nM)

1μl

10 nM

ORF PCR product (50 ng/μl)

0.1–1μl

0.1–1 ng/μl

KOD -Plus- Neo (1 U/μl)

1μl

0.02 U/μl

dH2O

Up to 50μl

0

82

Hideyuki Nakanishi and Hirohide Saito

8. According to the manufacturer’s instructions, set a thermal cycler to perform the Full PCR. An example using KOD -Plus- Neo is described below. Predenature: 94  C, 2 min. Denature: 98  C, 10 s. Extension: 68  C, 30 s/kb. (Repeat the denature and extension steps 29 times (total 30 cycles).) 9. Perform agarose gel electrophoresis, purification, and quantification of the Full PCR products following the same procedure as for the ORF PCR products (Subheading 3.3, steps 4–6) (see Note 2). 3.4 mRNA Synthesis by In Vitro Transcription

1. Mix the components of the in vitro transcription reaction as described below and incubate the mixture at 37  C for 4–6 h. Due to their high translational efficiency, low immunoresponse [6, 7], and high miRNA-responsiveness [8], we recommend N1-methyl-pseudoUTP-containing mRNAs as a first choice. An example of a mixture using MEGAscript T7 Transcription Kit is described below. Component

Volume

Final concentration

10 T7 reaction buffer

1μl

1

100 mM ARCA

0.6μl

6 mM

75 mM GTP

0.2μl

1.5 mM

75 mM ATP

1μl

7.5 mM

75 mM CTP

1μl

7.5 mM

100 mM N1-methyl-pseudoUTP

0.75μl

7.5 mM

T7 enzyme mix

1μl

Template DNA (Full PCR product)

400 ng

dH2O

Up to 10μl

40 ng/μl

An alternative combination of modified nucleotides using pseudoUTP and 5-methyl-CTP [9] is shown below. Component

Volume

Final concentration

10 T7 reaction buffer

1μl

100 mM ARCA

0.6μl

6 mM

75 mM GTP

0.2μl

1.5 mM

75 mM ATP

1μl

7.5 mM (continued)

Cell Purification by MicroRNA-Responsive Synthetic mRNA

83

Component

Volume

Final concentration

100 mM 5-methyl-CTP

0.75μl

7.5 mM

100 mM pseudoUTP

0.75μl

7.5 mM

T7 enzyme mix

1μl

Template DNA (Full PCR product)

400 ng

dH2O

Up to 10μl

40 ng/μl

2. Add 1μl of TURBO DNase and incubate at 37  C for 30 min. 3. According to the kit-appended protocol, purify the RNA by using an RNA purification kit. 4. Mix the components of the dephosphorylation reaction and incubate at 37  C for 30 min. An example of a mixture using rApid alkaline phosphatase is described below. Component

Volume

Purified RNA 10 rApid alkaline phosphatase buffer

5μl

rApid alkaline phosphatase

1μl

dH2O

Up to 50μl

5. Purify the RNA by an RNA purification kit. 6. Measure the concentration of the purified RNAs by NanoDrop or a comparable spectrophotometer. 7. According to the kit-appended protocol, check the size and quality of the RNA by Bioanalyzer or a comparable electrophoresis apparatus (see Note 3). 3.5 mRNA Transfection and Flow Cytometry

1. According to the transfection reagent-appended protocol, cotransfect the miRNA-responsive and reference reporter mRNAs into the cells (see Note 4). The reporter mRNAs should be transfected as a single transfection complex (i.e., the single transfection complex should contain both mRNAs. Do not transfect two mRNAs as two separate transfection complexes containing only one of them). An appropriate molar ratio of miRNA-responsive and reference reporter mRNAs depends on the selected fluorescent proteins and the laser and filter sets of the cell sorter. If the cells aggregate, we recommend detaching them into single cells before the transfection to improve the transfection efficiency. 2. One day after the transfection, detach the cells from the plates and suspend them in medium or FACS buffer.

84

Hideyuki Nakanishi and Hirohide Saito

3. Filtrate the cells by using cell strainers (see Note 5). 4. According to the manufacturer’s instructions, sort the cells based on the ratio of miRNA responsive/reference reporter fluorescence using FACS Aria or a comparable cell sorter (see Note 6).

4

Notes 1. Candidate miRNAs can be selected based on a microarray analysis or small RNA sequencing. miRNAs whose expression is highly different between target and nontarget cells are good candidates. Alternatively, miRNAs whose target cell-specific expression is confirmed in multiple studies are also promising candidates. However, highly expressed miRNAs are not always highly active [4]. Thus, we recommend preparing multiple miRNA-responsive reporter mRNAs and comparing their responsiveness. 2. If extra bands are observed by an agarose gel electrophoresis, modulate the PCR conditions or change the PCR enzymes. Ramp rates of the thermal cyclers can also have an effect. Alternatively, the main PCR products can be purified using a gel purification kit. 3. In some apparatus, the electrophoretic speed of the RNAs containing modified nucleotides differs from RNAs containing only natural nucleotides. In addition, highly structured RNAs tend to show multiple bands. 4. A suitable transfection reagent depends on the cell type. We recommend Lipofectamine MessengerMAX (ThermoFisher Scientific) as the first choice, but other transfection reagents such as Stemfect (ReproCELL) or Lipofectamine Stem (ThermoFisher Scientific) may show higher transfection efficiency in some cell types. If transfection by transfection reagents remains difficult, electroporation can be an alternative method. 5. To avoid cell aggregation, we recommend filtrating the cells immediately before sorting. 6. If the separation of target and nontarget cells is not sufficient, increasing the copy number of the miRNA target sites may improve the miRNA-sensitivity [10–12]. Alternatively, find miRNAs whose activity is higher in nontarget cells than in target cells and insert their complementary sequences into the reference reporter mRNA. The selective decrease of the reference reporter expression in nontarget cells increases the miRNA-responsive/reference reporter fluorescence ratio to eventually improve the separation of the target and nontarget cells (Fig. 4) [13].

Cell Purification by MicroRNA-Responsive Synthetic mRNA

85

Fig. 4 Schemes for clearly separating target and nontarget cells. When miRNA-responsive reporter expression is sufficiently repressed in the target cells, the target and nontarget cells can be clearly separated (a). In contrast, when the repression is not sufficient, it is difficult to clearly separate the target cells from the nontarget cells (b). The separation can be improved by inserting additional miRNA target sites into 50 or 30 UTRs of the miRNA-responsive reporter mRNAs to enhance the miRNA sensitivity (c). Alternatively, target sites for miRNAs that are highly active in nontarget cells but not in target cells can be inserted into UTRs of the reference reporter mRNAs to increase the miRNA-responsive/reference fluorescence ratio in nontarget cells (d)

86

Hideyuki Nakanishi and Hirohide Saito

Acknowledgments We would like to thank Dr. Peter Karagiannis and Ms. Miho Nishimura (Kyoto University) for English proofreading and administrative support, respectively. References 1. Kozomara A, Griffiths-Jones S (2011) MiRBase: integrating microRNA annotation and deep-sequencing data. Nucleic Acids Res 39: D152–D157 2. Parr CJC, Katayama S, Miki K, Kuang Y, Yoshida Y, Morizane A, Takahashi J, Yamanaka S, Saito H (2016) MicroRNA-302 switch to identify and eliminate undifferentiated human pluripotent stem cells. Sci Rep 6:32532 3. Hirosawa M, Fujita Y, Parr CJC, Hayashi K, Kashida S, Hotta A, Woltjen K, Saito H (2017) Cell-type-specific genome editing with a microRNA-responsive CRISPR-Cas9 switch. Nucleic Acids Res 45:e118 4. Miki K, Endo K, Takahashi S et al (2015) Efficient detection and purification of cell populations using synthetic MicroRNA switches. Cell Stem Cell 16:699–711 5. Lambert TJ (2019) FPbase: a communityeditable fluorescent protein database. Nat Methods 16:277–278 6. Andries O, Mc Cafferty S, De Smedt SC, Weiss R, Sanders NN, Kitada T (2015) N1-methylpseudouridine-incorporated mRNA outperforms pseudouridineincorporated mRNA by providing enhanced protein expression and reduced immunogenicity in mammalian cell lines and mice. J Control Release 217:337–344 7. Svitkin YV, Cheng YM, Chakraborty T, Presnyak V, John M, Sonenberg N (2017) N1-methyl-pseudouridine in mRNA enhances

translation through eIF2α-dependent and independent mechanisms by increasing ribosome density. Nucleic Acids Res 45:6023–6036 8. Parr CJC, Wada S, Kotake K, Kameda S, Matsuura S, Sakashita S, Park S, Sugiyama H, Kuang Y, Saito H (2020) N 1-Methylpseudouridine substitution enhances the performance of synthetic mRNA switches in cells. Nucleic Acids Res 48:1–7 9. Kariko´ K, Muramatsu H, Welsh FA, Ludwig J, Kato H, Akira S, Weissman D (2008) Incorporation of pseudouridine into mRNA yields superior nonimmunogenic vector with increased translational capacity and biological stability. Mol Ther 16:1833–1840 10. Nakanishi H, Miki K, Komatsu KR, Umeda M, Mochizuki M, Inagaki A, Yoshida Y, Saito H (2017) Monitoring and visualizing microRNA dynamics during live cell differentiation using microRNA-responsive non-viral reporter vectors. Biomaterials 128:121–135 11. Matsuura S, Ono H, Kawasaki S, Kuang Y, Fujita Y, Saito H (2018) Synthetic RNA-based logic computation in mammalian cells. Nat Commun 9:4847 12. Endo K, Hayashi K, Saito H (2019) Numerical operations in living cells by programmable RNA devices. Sci Adv 5:eaax0835 13. Endo K, Hayashi K, Saito H (2016) Highresolution identification and separation of living cell types by multiple microRNAresponsive synthetic mRNAs. Sci Rep 6:21991

Part II Engineering Mammalian Cells to Sense Artificial Inputs

Chapter 6 Green Light-Controlled Gene Switch for Mammalian and Plant Cells Nils Schneider, Claire V. Chatelle, Rocio Ochoa-Fernandez, Matias D. Zurbriggen, and Wilfried Weber Abstract The quest to engineer increasingly complex synthetic gene networks in mammalian and plant cells requires an ever-growing portfolio of orthogonal gene expression systems. To control gene expression, light is of particular interest due to high spatial and temporal resolution, ease of dosage and simplicity of administration, enabling increasingly sophisticated man–machine interfaces. However, the majority of applied optogenetic switches are crowded in the UVB, blue and red/far-red light parts of the optical spectrum, limiting the number of simultaneously applicable stimuli. This problem is even more pertinent in plant cells, in which UV-A/B, blue, and red light-responsive photoreceptors are already expressed endogenously. To alleviate these challenges, we developed a green light responsive gene switch, based on the light-sensitive bacterial transcription factor CarH from Thermus thermophilus and its cognate DNA operator sequence CarO. The switch is characterized by high reversibility, high transgene expression levels, and low leakiness, leading to up to 350-fold induction ratios in mammalian cells. In this chapter, we describe the essential steps to build functional components of the green light-regulated gene switch, followed by detailed protocols to quantify transgene expression over time in mammalian cells. In addition, we expand this protocol with a description of how the optogenetic switch can be implemented in protoplasts of A. thaliana. Key words AdoB12, CarH, Green Light, Light-switchable gene expression, Mammalian cells, Optogenetics, Plant cells

1

Introduction Inducible gene switches are key components of synthetic biological systems and have enabled numerous applications in vitro, in cellulo, and in vivo [1, 2]. Among others, gene switches have been used to generate genetic oscillators [3], Boolean logic gates [4], platforms for drug discovery [5], closed loop–controlled gene networks [6], cell therapeutic networks [7], and even biomaterials [8]. Initially, gene inducers were based on naturally occurring gene switches, responding to chemical cues [9]. However, these systems have

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

89

90

Nils Schneider et al.

inherent drawbacks, notably low spatial resolution due to lateral diffusion and low temporal resolution due to the need for inducer removal from the medium. These drawbacks prompted the development of a second generation of genetic switches that can be controlled with light [10–12]. Such optogenetic gene expression systems are generally based on plant and bacterial photoreceptors functionally modified by fusing them to DNA binding domains and transcriptional activators/repressors. Advantages of these systems are spatial and temporal resolution that cover all biologically relevant scales but also ease of dosage, orthogonality, minimized invasiveness, and controllable reversibility. Optogenetic switches have therefore become outstanding tools to interface the control of biological functions in vivo with programmable machines. First optogenetic systems mostly used phytochromes, cryptochromes and LOV-domains as optogenetic components, covering the absorption spectrums of Red/Far-red, blue, and UVB light, see www.optobase.org [13]. The crowding of numerous optogenetic system in these regions of the optical spectrum limits the use of multiple wavelengths for orthogonal systems and has left an unused gap in the middle of the optical spectrum. Even more important, plant-based phytochrome and cryptochrome optogenetic switches cannot easily be used for synthetic biology applications in plants without the risk of cross talk with endogenous photoreceptors and therefore unwanted downstream effects [14]. Thus, the lack of dedicated green light receptors in plants opens a window of opportunity to control gene expression in plants with green light. Hence, to enlarge the toolbox of orthogonal optogenetic gene switches in mammalian cells and to expand optogenetic systems to plant cells, we developed a first green light responsive gene switch [15]. The gene expression system is based on the bacterial transcriptional repressor CarH found in gram-negative bacteria such as Myxococcus xanthus and Thermus thermophilus. In these bacteria, CarH controls the expression of the carotenogenic gene cluster to protect cells against phototoxic stress [16–18]. Photosensitivity is conferred to CarH by the cofactor and chromophore 50 deoxyadenosylcobalamin AdoB12, which has an absorption peak at 525 nm. In the dark AdoB12 binds to CarH and induces homo-tetramerization, in which state CarH has a high affinity for the cognate DNA operator CarO. Illumination of the cells leads to photolysis of AdoB12 via disruption of a Co-C bond, followed by the disassembly of the CarH-AdoB12 complex, which in turn leads to monomerization and detachment of CarH from the DNA [19]. The first uses of CarH for optically controlled biological systems included optogenetic control of transgene expression in bacteria [20], activation of fibroblast growth factor receptor signaling [21] but also to generate a light-switchable protein hydrogel based on CarH and SpyTag-SpyCatcher chemistry [22].

Green Light-Controlled Gene Switch

91

The natural gene switch in bacteria is active in darkness and repressed by green light. In adapting the system to the use in mammalian cells, we also aimed to modify the system for constitutive expression, which can be conditionally suppressed by green light treatment. To that end, we generated two basic constructs: (1) A reporter gene, preceded by varying numbers of CarO binding sequences along with the minimal promoter PhCMVmin, derived from the human cytomegalovirus immediate early promoter; (2) A fusion protein of CarH and VP16, a strong transactivation domain derived from the Herpes simplex virus. Transfection and expression in cells supplemented with AdoB12 lead to reporter gene expression in the dark, while green light-illumination suppresses gene expression [15]. The functional components of the system, as well as the fundamental mechanism of the switch, are illustrated in Fig. 1. The CarH-based gene switch significantly expands the toolbox to build specific, orthogonal and precisely controlled gene networks in both mammalian cells and plant cells. To enable the reader to implement the system, we here provide a detailed description of how to build reporter vectors by cloning varying numbers of CarO upstream of a transgene of choice. Next, we detail how to

Fig. 1 Illustration of the CarH based optogenetic gene switch. Top: In the dark, cofactor Ado-B12 binds to the CarH-VP16 fusion protein, resulting in tetramerization and binding to CarO repeat sequences. The transactivation domain VP16 recruits the transcription preinitiation complex on the minimal promoter PhCMVmin, resulting in the expression of the gene of interest. Bottom: Illumination with green light (525 nm) leads to photolysis of the bond between 50 deoxyadenosyl (Ado) and cobalamin. Binding of a histidine residue (His) of CarH triggers a conformational change, leading to monomerization and detachment of CarO. The expression of the gene of interest is stopped

92

Nils Schneider et al.

determine temporal expression dynamics to quantitatively measure the output of the system in mammalian cells. Last, we describe the implementation of the system in plant protoplasts.

2

Materials

2.1 Cloning of Multiple CarO Repeats

1. As donor plasmid, a vector is required that contains the gene of interest under the control of a minimal human CMV promoter (PhCMVmin sequence: CCTATATAAGCAGAGCTCGTTTAGT GAACCGTCAGATCGCCTGGAGACGCCATCCACGCTG TTTTGACCTCCATAGAAGACACCGGGACCGATCCAG CCT) and followed by a polyadenylation signal such as SV40 poly(A). In this exemplary case, we use the plasmid pHB145, which is available upon request. As backbone plasmid, we recommend the use of the pRSET-A vector, which contains an upstream XbaI and a downstream ClaI site. 2. Q5 polymerase (New England Biolabs), dNTPs (New England Biolabs), thermocycler. 3. Agarose (Roth), TAE buffer, DNA loading buffer (New England Biolabs), agarose gel electrophoresis chamber, ethidium bromide (Roth), gel imaging system for long-wave UV transillumination, gel extraction kit (Qiagen). 4. XbaI (New England Biolabs), NheI (New England Biolabs), ClaI (New England Biolabs). 5. T4 DNA ligase (New England Biolabs). 6. E. coli Top10 chemically competent cells, thermo-shaker, LB Medium, agar plates containing the antibiotic for the employed vector backbone, appropriate selection antibiotic. 7. Miniprep Kit (Qiagen), Midiprep Kit (Macherey-Nagel).

2.2 Mammalian Cell Culture, Transfection, and Light Treatment

1. Vectors to constitutively express CarH-VP16 fusion proteins are available upon request for mammalian expression hosts (pHB144) as well as for plants (pROF254). The reporter plasmids can be cloned as described in Subheading 2.1 or are available upon request with CarO2-PhCMVmin-SEAP-PA (pCVC034), CarO4-PhCMVmin-SEAP-PA (pCVC035) and CarO8-PhCMVmin-SEAP-PA (pCVC036). 2. HEK-293 cells (DSMZ), cell culture hood and incubator, 10 cm dishes, 12-well plates, 50 ml conical centrifuge tubes, DMEM (PAN) supplemented with 10% (v:v) fetal calf serum (FCS) (PAN) and 1% (v:v) penicillin–streptomycin (PAN), Dulbecco’s phosphate-buffered saline, trypsin–EDTA (PAN), bright-field microscope, CASY cell counter including CASY cups and CASY ton buffer.

Green Light-Controlled Gene Switch

93

3. High-quality plasmid DNA (midi-prep), Opti-MEM (ThermoFisher Scientific), sterile filtered PEI solution containing 1 mg/ml of linear polyethylenimine (MW 25000), 15 ml conical centrifuge tubes, vortex mixer. 4. Coenzyme B12 solution: prepare 2.5 mM stock by dissolving 25 mg of coenzymeB12 (Sigma-Aldrich) in 6.33 ml of ultrapure water, wrap aliquots in aluminum foil to protect from light and store at 20  C until further use. Illumination boxes with 525 nm light-emitting diodes (LED) of controllable intensity. The illumination boxes used for this protocol are custom made, consist of PVC, protect the samples from light contamination and have an active gas exchange [23]. As safe light, red light (660 nm, 10μmol m2 s1) can be used. Photometer Sanawa LP1 or similar. 5. Eppendorf tubes, peqGOLD TriFast RNA extraction kit (Peqlab/VWR), V-bottom 96-well plate. 2.3 Quantification of mRNA Expression in Mammalian Cells

1. Chloroform, isopropanol (RNase free), RNase free water, fume hood with air exchange, cooling centrifuge for Eppendorf tubes. 2. RNA Clean & Concentrator-5 with included DNase (Zymo Research). 3. NanoDrop 1000 spectrophotometer, agarose (Roth), TAE buffer, DNA loading buffer (New England Biolabs), agarose gel electrophoresis chamber, ethidium bromide (Roth), gel imaging system for long-wave UV transillumination. 4. Primers SEAP_Fw (50 - CATGGACATTGACGTGATCCT ), SEAP_RV (50 -CACCTTGGCTGTAGTCATCTG), Actin_Fw (50 -CCCTGGAGAAGAGCTACGAG), Actin_Rv (50 -TCCAT GCCCAGGAAGGAAG), Absolute QPCR Mix SYBR Green ROX (ThermoFisher Scientific), 96-well PCR plate (ThermoFisher Scientific), adhesive qPCR Plate Seals (ThermoFisher Scientific), qPCR device qTOWER 2.0/2.2 (Analytik Jena) or similar, qPCR analysis software.

2.4 Quantification of SEAP Expression in Mammalian Cells

1. Transparent flat-bottom 96-well plate, microplate reader for absorbance measurement, heat block for 96-well plates, multichannel pipette or multistep pipette for volumes of 20μl. 2. 2 SEAP buffer: 21% (v:v) diethanolamine, 20 mM L-homoarginine, 1 mM MgCl2, pH 9.8, store at 4  C in the dark. pNPP solution: 120 mM para-nitrophenyl phosphate dissolved in water, store at 20  C.

94

Nils Schneider et al.

2.5 Reporter Gene Expression in Protoplasts 2.5.1 Protoplast Preparation

1. A. thaliana seeds: wild type, Columbia-0. Autoclaved water. Autoclaved filter paper strips. 12 cm2 petri dish. 2. SCA medium: 0.32% Gamborg B5 basal salt powder with vitamins, 4 mM MgSO4, 43.8 mM sucrose, 0.8% (w:v) Phytoagar, pH 5.8. 0.1% (v:v) Gamborg B5 Vit Mix, stored at 4  C. 3. Parafilm, sterile hood, 1.5 ml tubes, 80% ethanol, absolute ethanol (99.5%). 4. A. thaliana sterilization solution: 5% calcium hypochlorite (w: v), 0.02% Triton X-100 (v:v), 80% EtOH, 4  C. 5. Centrifuge, 15 ml round bottom tubes, wide-orifice tips, scalpel. 6. MMC solution: 10 mM MES, 40 mM, CaCl2, 85 g l1 mannitol, pH 5.8, stored at 4  C. 7. MMM solution: 15 mM MgCl2, 5 mM MES, 85 g l1 mannitol, pH 5.8, stored at 4  C. 8. 10 enzyme stock solution: 5% (w:v) cellulase Onozuka R10 and 5% (w:v) macerozyme R10 in MMC, stored at 20  C. 9. MSC solution: 10 mM MES, 0.4 M sucrose, 20 mM MgCl2, 85 g l1 mannitol, pH 5.8, stored at 4  C. 10. W5 solution: 2 mM MES, 154 mM NaCl, 125 mM CaCl2, 5 mM KCl, 5 mM glucose, pH 5.8, stored at 4  C. 11. Disposable 70μm pore size sieves, Rosenthal cell counting chamber.

2.5.2 Protoplast Transformation, Light Treatment, and Luciferase Assay

1. CarH-VP16 (pROF254), luciferase reporter plasmids CarO2 (pROF250), CarO4 (pROF251), CarO8 (pROF252). 2. MMM solution: see above. 3. PEG solution: 2.5 ml of 0.8 M mannitol, 1 ml of 1 M CaCl2, 4 g PEG4000, and 3 ml H2O, prepare fresh, mix thoroughly and incubate at 37  C until dissolution. 4. PCA solution: 0.32% (w:v) Gamborg B5 basal salt powder with vitamins, 2 mM MgSO4, 3.4 mM CaCl2, 5 mM MES, 0.342 mM L-glutamine, 58.4 mM sucrose, 80 g l1 glucose, 8.4μM Ca-pantothenate, 4μg ml1 biotin, pH 5.8, 0.1% (v:v) Gamborg B5 Vitamin, 50μg ml1 ampicillin, stored at 4  C. 5. AdoB12 (see above). 6. 525 nm light-emitting diode (LED) illumination boxes, nontreated 6-well and 24-well plates, white 96-well flat bottom plates, plate reader with luminescence read-out. 7. Firefly luciferase substrate: 20 mM tricine, 2.67 mM MgSO4, 0.1 mM EDTA, 33.3 mM DTT, 0.52 mM ATP, 0.27 mM acetyl-CoA, 0.47 mM D-luciferin, 5 mM NaOH, 264μM MgCO3, pH 8, stored at 80  C.

Green Light-Controlled Gene Switch

3

95

Methods

3.1 Cloning of Multiple CarO Repeats

Cloning of multiple identical sequence repeats is not straightforward and generally cannot be circumvented by ordering synthetic gene fragments. The following protocol describes a generic strategy to clone any desired number of short sequence repeats in front of a gene of interest (GOI), see Fig. 2. 1. Design and order a forward primer containing the restriction site XbaI, followed by the sequence of CarO, the restriction site NheI and an annealing region of ~20 bp towards the donor plasmid upstream of the PhCMVmin promoter (see Note 1). The resulting oligonucleotide should resemble the following sequence: NNNNNNTCTAGA ACACTCCGCAGAGATGTACAAAAGCTTGACAAAAACCTA GCTAGC NNNNNNNNNNNNNNNNNNNN with bold: restriction sites, italic: CarO sequence, underlined: binding region to donor plasmid (see Note 2). 2. Design and order a reverse primer that binds with ~20 bp to the donor plasmid downstream of the gene of interest and the polyadenylation signal. Incorporate a ClaI restriction site at the 50 end of the primer. The resulting oligonucleotide should resemble the following sequence: NNNNNNATCGAT NNNNNNNNNNNNNNNNNNNN, with bold: restriction sites, underlined: binding region to donor plasmid. When designing sequences, please be aware that ClaI and XbaI restriction enzymes are sensitive to methylation (see Note 3). 3. Use forward and reverse primers of steps 1 and 2 to PCR-amplify the cassette of XbaI-CarO-NheI-PhCMVminGOI-poly(A)-ClaI and the donor plasmid (here: pHB145) with the Q5 polymerase, following the manufacturer’s instructions. 4. Gel purify the resulting PCR fragment. 5. Digest both the backbone plasmid and the purified PCR fragment of step 4 with XbaI and ClaI and gel purify both fragments. 6. Ligate the digested PCR fragment into the digested plasmid backbone to obtain the CarO1-reporter-plasmid. 7. Digest CarO1-reporter-plasmid with NheI and ClaI and gel purify the backbone. 8. Digest CarO1-reporter-plasmid with XbaI and ClaI and gel purify the excised fragment. 9. Ligate the purified backbone of step 7 and the purified fragment of step 8 to obtain the CarO2-reporter-plasmid.

96

Nils Schneider et al.

Fig. 2 Cloning of multiple CarO repeats. The iterative addition of CarO repeats is based on the use of restriction enzymes with compatible cohesive ends (XbaI and NheI). A detailed description of the required steps is given in Subheading 3.1

Green Light-Controlled Gene Switch

97

10. Transform the ligation product in Top10 chemically competent cells via heat shock. After the addition of LB medium, incubate the transformed cells for 1 h at 30  C, streak out on antibiotic-containing agar plates and incubate overnight at 30  C (see Note 4). 11. Pick multiple single colonies and inoculate each in 3 ml of LB containing the appropriate selection antibiotic and incubate overnight at 30  C. Isolate the plasmid DNA and analyze via Sanger sequencing to assure the correct insertion of the correct amount of CarO repeats. 12. Iteratively increase the number of CarO repeats by repeating the steps 7–11 (see Note 5). 13. For final use of CarO plasmids in mammalian cell culture, prepare Midi-Preps to assure a sufficient quantity and quality. 3.2 Mammalian Cell Culture, Transfection, Light Incubation, and Harvest

1. Culture HEK-293 cells in two 10 cm dishes for 2–3 days until they reach 80% confluency. Alternative cell lines can be employed as well (see Note 6). Pre-heat trypsin–EDTA, DPBS, and DMEM complete medium to 37  C. 2. For each 10 cm dish, remove the medium, wash cells gently with 10 ml DPBS and add 1 ml of trypsin–EDTA. Incubate the cells approximately for 3 min at 37  C until cells detach from the surface upon gentle agitation. Add 9 ml of DMEM complete medium, resuspend the cells and pool the cells of both 10 cm dishes into one 50 ml centrifugation tube. Spin the cells down at 300  g for 3 min, decant the supernatant and resuspend the cells in 20 ml of DMEM complete medium. To determine the cell count with the CASY-counter, dilute 100μl of the cell suspension in 10 ml CASYton buffer in a CASY cup. Determine the cell concentration automatically using the CASY counter. 3. Dilute HEK-293 cells to a concentration of 250,000 cells ml1 and apply 1 ml of cell suspension into each well of four entire 12-well plates. After seeding, transfer the plates into the cell culture incubator. Ensure a homogeneous distribution of the cells on the surface by moving the plates back and forth several times in a linear motion. 4. Observe the attachment and growth of the cells with the bright-field microscope. For transfection, the cells should ideally arrive at a confluency of ~30%. 5. For transfection, mix for each well 1200 ng of pHB144 (CarHVP16) and 300 ng of the reporter construct pCVC036 (CarO8-PhCMVmin-SEAP) in 50μl of Opti-MEM and mix thoroughly. To assure homogeneity, prepare one master mix for all wells to be transfected, containing a total of 60μg of pHB144, 15μg of pCVC036 and fill up to 5 ml with Opti-MEM (see Note 7).

98

Nils Schneider et al.

6. For transfection, mix for each well 45μl of Opti-MEM and 5μl of PEI solution. To assure homogeneity, prepare one master mix for all wells containing 250μl of PEI solution and 4750μl of Opti-MEM to be transfected and mix thoroughly. 7. Mix the reagents prepared in steps 5 and 6 and immediately mix by vigorous vortexing for at least 10 s. 8. Incubate the transfection mixture of step 7 for 15 min at room temperature, vortex shortly and add 100μl dropwise onto the surface of the medium within each well of the four 12 well plates. Transfer the plates into the incubator and gently move the plates back and forth multiple times. After ~1 h, the first cells are expected to start to express the CarH-VP16 protein. However, since CarH binding to CarO is dependent on AdoB12 administration, the system is not light sensitive at this stage. Therefore, no further action is required (see Note 8). 9. After incubation for 24 h, the transcription factor is robustly expressed and can be loaded with the chromophore. Before starting, switch the laboratory to a red safe-light condition, at which the chromophore has an absorption minimum. Keep working at safe-light until the handling of living cells is finished in step 12. Next, add for each transfected well 8μl of the 2.5 mM AdoB12 stock solution to obtain a final concentration of 20μM (see Note 9). 10. For time point 1, immediately harvest 200μl supernatant medium of three wells (for triplicate measurements) into Eppendorf tubes and store the samples at 20  C until further use. To harvest cells for RNA samples, wash HEK-293 cells gently off of the surface with the remaining medium and transfer into an Eppendorf tube (see Note 10). Spin the cells down by centrifugation for 3 min at 300  g. Decant the supernatant and transfer the tube under a hood with air exchange. Immediately add 600μl of peqGOLD TriFast solution to the cell pellet (see Note 11). Pipette the mixture several times up and down to obtain thorough cell lysis. Keep the sample for 5 min at RT. Store the cell lysate in Trifast at 80  C until further use in Subheading 3.4. 11. Transfer two 12-well plates for green light exposure into the light box with LEDs with an emission peak at 525 nm and adjusted to 5μmol m2 s1. Transfer the 12-well plate for no light exposure into the light box with the LEDs switched off (see Note 12). 12. For time points 2 to 8, harvest supernatant and cells after 1 h, 2 h, 4 h, 6 h, 10 h, 24 h and 48 h as described in step 10 (see Note 13).

Green Light-Controlled Gene Switch

3.3 Quantification of SEAP Expression in Mammalian Cells

99

1. Thaw frozen samples of supernatants and transfer them into a single V-bottom 96-well plate. Use adhesive tape to seal the plate. Incubate the plate for 30 min at 65  C on a heat block to inactivate heat-labile phosphatases other than SEAP. 2. Centrifuge the plate at 1000  g for 3 min to pellet cell debris and condensed water. 3. Transfer 100μl 2 SEAP buffer into 48 wells of a transparent flat-bottom 96-well plate. Add 20μl of each supernatant sample as well as 60μl of DMEM (see Note 14). 4. Thaw the pNPP solution and add 20μl to each well using a multichannel or multi-step pipette (see Note 15). Immediately mix the samples by agitating the plate for 30 s. Measure the absorbance at 405 nm every minute for at least 30 min. 5. Calculate the slope S ¼ ΔA 405/Δt (min1) for the linear range of absorbance increase. With the slope, use Lambert– Beer’s law to calculate the enzymatic activity EA with EA ¼ S  ν/εpNP  d, where εpNP ¼ 18,600 M1 cm1 is the extinction coefficient of the product, d is the length of the light path in cm, and ν is the dilution factor VMeasurement/VSample (200μl/20μl in this example). SEAP activity is generally given in U l1 ¼μmol min1 l1. Representative results are shown in Fig. 3.

Fig. 3 Representative results of light-dependent reporter gene expression over time. Left y-axis: SEAP reporter protein expression of cells grown in the dark (black square) and cells exposed to green light (white square, 525 nm, 5μmol m2 s1) for the indicated periods of time. Right y-axis: fold-change in SEAP-mRNA expression towards t ¼ 0 of cells grown in dark (black triangle) and cells exposed to green light (white triangle). Data are means  Stdev (n ¼ 3). (Reprinted (adapted) with permission from [15]. Copyright (2018) American Chemical Society)

100

Nils Schneider et al.

3.4 Quantification of mRNA Expression in Mammalian Cells

1. Thaw the cell lysates in Trifast prepared in Subheading 3.2. 2. Under a fume hood with air exchange, add 120μl of chloroform to each sample and shake vigorously for 15 s. Incubate the samples for 5 min at room temperature. 3. Centrifuge the samples for 10 min at 12,000  g. After centrifugation, genomic DNA and proteins are separated into the lower phenol–chloroform interphase, while RNA is accumulated in the upper aqueous phase. 4. Transfer the upper aqueous phase into new tubes and discard the phenol-chloroform phase (see Note 16). Add 300μl of RNase free isopropanol and incubate on ice for 15 min. 5. Centrifuge the samples for 10 min at 12,000  g at 4  C. The resulting RNA pellet should be clearly visible. Carefully remove the supernatant and air-dry the pellet. Resuspend the pellet in 20μl of RNase free water. RNA isolation based on phenol and guanidineisothiocyanate does not exclude plasmid DNA from the RNA fraction. Because plasmid DNA encoding the SEAP reporter gene will interfere with the quantification of SEAP mRNA via qRT-PCR, it is pivotal to treat the RNA samples with DNase first. 6. Use DNase I to treat 10μg of RNA for each sample and in a total volume of 50μl. Incubate the sample for 30 min at room temperature. 7. Purify the RNA samples following the instructions of the RNA Clean & Concentrator-5 Kit. Elute the RNA in 15μl RNase free water. 8. Measure the RNA concentration with the NanoDrop spectrophotometer and verify that the absorbance ratio 260/280 is ~2.0. Run at least 200 ng of each RNA sample on an agarose gel to verify that the RNA is still intact. If intact, two sharp bands corresponding to rRNA 28S and 18S can be observed. 9. Use 500 ng of RNA for each condition to synthesize cDNA with the First Strand cDNA Synthesis Kit. Use supplied oligo (dT)18 and/or random primers and follow the manufacturer’s instructions step-by-step to obtain cDNA. It is important to include negative controls in which the reverse transcriptase is omitted to verify that no DNA contaminations remain. A negative control without RNA is recommended to verify that reagents are not contaminated. 10. Use the Absolute qPCR SYBR Green ROX Mix to prepare the reaction mixtures for qPCR. For SEAP mRNA quantification prepare technical triplicates of each of the three biological triplicates. Mix 10μl of 2 reaction mixture, 1.4μl of forward primer SEAP_Fw, 1.4μl of reverse primer SEAP_Rv, 0.5μl of

Green Light-Controlled Gene Switch

101

cDNA and 6.7μl ultra-pure water (see Note 17). Prepare one master mix (without cDNA) for all reactions to ensure homogeneity. For quantitative normalization use the same cDNA samples, however, use primers targeting ACTB Actin_Fw and Actin_Rv. In addition to the cDNA samples of the time series, include reactions for negative controls that were not treated with reverse transcriptase as well as the RNA-negative control. Mix all reagents in the real-time qPCR plate and seal the plate to prevent evaporation. 11. Perform quantitative real-time PCR and measure the fluorescence of SYBR Green and ROX after every cycle for 40 cycles in the real-time thermal cycler (see Note 18). Normalize the fluorescence of the reporter fluorophore SYBR Green by using the ROX fluorescence data. Determine Cq values with the ΔΔCq method to infer the relative gene expression levels. Representative results of mRNA expression over time are shown in Fig. 3. 3.5 Protoplast Preparation

1. Rinse seeds multiple times with 80% (v:v) ethanol to remove large particles (dirt, plant material) and sterilize the seeds with 1 ml of the A. thaliana sterilization solution for 10 min under agitation. 2. Replace the sterilization solution and wash with 1 ml of 80% (v: v) ethanol, incubate for 5 min under agitation and repeat the washing step. 3. Replace the solution with 1 ml absolute ethanol (99.5%) and incubate for 1 min under agitation. Remove the ethanol and let the seeds dry completely under the sterile hood. 4. Place several autoclaved strips of filter paper in a 12 cm2 plate containing SCA medium. Add sterile water to the seeds and place 200–300 seeds in line on each paper strip. Seal the plate with parafilm and incubate the seeds at 23  C in a growth chamber, with a 16 h light–8 h dark regime for 1 to 2 weeks. 5. Harvest A. thaliana leaves while avoiding to include roots and seeds. With a scalpel, slice the leaves in a volume of 2 ml of MMC and transfer the sliced plant material into a new petri dish containing 7 ml of MMC (see Note 19). 6. Add 1 ml of 10 enzyme stock solution (final enzyme concentration: 0.5%). Seal the dish with parafilm and cover with aluminum foil. Allow the enzymatic digestion to proceed overnight (12–16 h) in the dark at 22  C. 7. Slowly pipet up and down the digested leaf material to release the protoplasts from the plant material (see Note 20). Pass the solution through a sieve with a pore size of 70μm. Transfer the filtered protoplast solution into a 15 ml round bottom tube. Use a dedicated tube for each plate of digested leaf material.

102

Nils Schneider et al.

8. Pellet protoplasts by centrifugation for 20 min at 100  g (see Note 21). Remove supernatant and resuspend in 10 ml MSC. Very carefully overlay the 10 ml of protoplast solution with 2 ml of MMM (see Note 22). 9. Centrifuge for 10 min at 80  g to accumulate the protoplasts at the interphase of MSC and MMM. Collect the interphase and transfer into a new 15 ml conical tube with 7 ml of W5 solution. This step can be performed up to three times. 10. Pellet the collected protoplasts for 10 min at 100  g and resuspend them in a defined volume of W5 for counting. Determine the cell density using a Rosenthal cell counting chamber. Pellet the protoplasts at 80  g for 5 min, discard the supernatant and add an appropriate volume of MMM solution to obtain a density of 5  106 cells ml1. 3.6 Protoplast Transformation, Light Treatment, and Luciferase Assay

To demonstrate the implementation of the green light-dependent gene switch in plant protoplast, the influence of number CarO repeats on reporter gene expression is described. To that end, protoplasts are transformed with CarO2, CarO4, and CarO8, and either exposed to green light or darkness. Representative results are shown in Fig. 4.

Fig. 4 Representative results of light-dependent reporter gene expression in plant protoplasts. Protoplasts incubated with 20μM of AdoB12 in the dark (gray bars) or in green light (white bars, 525 nm, 5μmol m2 s1) express luciferase reporter dependent on light condition and the number of CarO repeats. The displayed values are means  SEM (n ¼ 6). RLU ¼ relative luminescence units. (Reprinted (adapted) with permission from [15]. Copyright (2018) American Chemical Society)

Green Light-Controlled Gene Switch

103

1. Transform the protoplast with the high-quality plasmid DNA encoding CarH-VP16 (pROF254) and either the luciferase reporter plasmid CarO2 (pROF250), CarO4 (pROF251) or CarO8 (pROF252). For each transformation, prepare a total amount of 30μg of DNA in H2O with a molar ratio of 1:3. Adjust the total volume to 20μl with MMM. To assure homogeneity, it is recommended to prepare one master mix for all identical transformation conditions. 2. Per well of a 6-well culture plate, transfer 20μl DNA solution to the edge of the surface (slightly tilt the plate for easier pipetting in the following steps). Dispense 100μl of the protoplast solution to each well (500,000 protoplasts per well) and mix by gentle pipetting. Incubate for 5 min. 3. Gently shake the 6-well plate from side to side to distribute the protoplasts and DNA along the edge before directly adding slowly 120μl of PEG solution drop-by-drop. Do not mix after the addition of PEG. Incubate for 8 min and quickly add 120μl MMM and 1.2 ml of PCA. Gently mix after the addition of PCA, the final volume obtained should be at least 1.6 ml. 4. After transformation, split the protoplasts in different 24-well plates for either dark or green light treatment, where each well should contain at least 150,000 protoplasts (ca. 480μl of the protoplast suspension) needed for the measurement of 6 replicates. Add AdoB12 to a final concentration of 20μM and seal the plates with parafilm. 5. Illuminate the plates with either green light (525 nm, 5μmol m2 s1) or keep in the dark as described in Subheading 3.2. 6. After 24 h, perform the luciferase assay to quantify reporter gene expression levels. It is recommended to perform the experiment in a dark room with red safe light. First, gently mix the protoplast suspension with a pipette and transfer 80μl into the wells of a white 96-well flat bottom plate. Include 6 replicates for each condition. Add 20μl of firefly luciferase substrate, shake the plate for 10 s to obtain homogeneous substrate repartition and measure the luminescence for 20 min (integration time 0.1 s). 7. For each well, plot the luminescence values over time. For each set of experimental replicates of each experimental condition, select a time frame in which the luminescence remains stable. This plateau generally excludes an initial phase, in which protoplasts are still taking up the luciferase substrate, and occasionally also a decay of the luminescence signal (see Note 23). For the selected time frame and for each well, calculate the average luminescence value (see Note 24). Representative results of luciferase expression in protoplasts are shown in Fig. 4.

104

4

Nils Schneider et al.

Notes 1. The restriction enzymes XbaI and NheI contain compatible cohesive ends, a prerequisite of the described cloning strategy. After ligation, the restriction site is destroyed. 2. The first six random nucleotides are generally recommended for restriction cleavage close to the end of DNA fragments. 3. ClaI is blocked by DAM methylation for sequences ATCGAT C and GATCGAT and XbaI for sequences TCTAGATC and GATCTAGA. Accordingly, prevent the use of random nucleotides resulting in these MTase recognition sites. 4. Cultivating transformed bacteria at 30  C reduces the risk of losing repetitive sequence repeats of CarO due to homologous recombination relative to incubation at 37  C. If time is not a constraint, performing all bacterial growth steps at room temperature is recommended. 5. By combining different backbones and fragments, any desired count of CarO can be achieved. 6. Apart from human HEK-293 cells (ATCC®: CRL-1573), the CarH-based optogenetic switch was also successfully tested in HeLa (human, ATCC®: CCL-2), NIH/3T3 (mouse, ATCC®: CCL-2) and COS-7 (African green monkey, ATCC®: CRL-1651) cells. However, to obtain the same fold-change expression levels as in HEK-293 cells, optimization of transfection and cell culture media are advised. 7. The quantities are calculated to contain a safety margin for pipetting errors. 8. The required addition of the chromophore is an advantage of this particular system, which allows handling cells safely without restricting the light conditions, in contrast to other widely used optogenetic systems that are active without the addition of an exogenous chromophore. This might be especially advantageous when creating stable cell lines containing the CarHbased gene switch. 9. AdoB12 is not stable in DMEM and will slowly degrade with a half-life time of approx. 24 h. In consequence, the use of higher initial concentrations of AdoB12 will lead to longer sustained expression. 10. HEK-293 cells can be detached easily by repeatedly aspirating the medium. For other cell types, use trypsin–EDTA for detachment. 11. peqGOLD TriFast contains phenol, which has corrosive effects on the respiratory tract and skin. Use only with appropriate precautions/safety measures.

Green Light-Controlled Gene Switch

105

12. Measure light intensities with the photometer. Importantly, in the case of the employed Sanwa LP1 device, the use of wavelengths different than the reference wavelength requires a conversion with a wavelength-dependent correction coefficient. In addition, assure that the fan in the illumination box is running for both green light and dark conditions to enable gas exchange. 13. Ideally, check cell viability and density under a bright-field microscope equipped with a red-light filter. 14. The dilution of the supernatant is necessary since otherwise the absorbance measurement rapidly reaches saturation. 15. In case air-bubbles have formed, they can and should be destroyed with a wooden toothpick prior to measurement. 16. TriFast has to be disposed of as hazardous waste. 17. The efficiency of the PCR for the indicated primers amplifying cDNA of SEAP and ACTB were determined as 100%, a repeat of the analysis is therefore not necessary. 18. The specificity of the PCR product can be measured in the end by performing a melt curve analysis and by running the PCR products on an agarose gel. 19. It is recommended to use sterile tweezers/forceps to transfer the leaves from the square/growth plate into the petri dish. 20. Pipetting is done with wide orifice tips or Stripettes to avoid damaging the protoplasts. This note applies to all the steps involving pipetting of protoplasts. 21. Use medium acceleration and lowest deceleration settings for all centrifugation steps. 22. Gentle inversion of the tube before adding the MMM solution leads to a clean phase separation. 23. The first phase of increasing luminescence typically takes 1–2 min. The signal then reaches a plateau phase, which can vary in duration depending on the configuration of transfected plasmids or other experimental conditions. In the case of the system and experimental conditions shown in here, the decay occurs approximately 3–7 min after the start of the plateau phase. 24. Experimental outliers can be excluded by statistical analysis following the guidelines for bicistronic reporter assay data provided by Jacobs and Dinman [24].

106

Nils Schneider et al.

Acknowledgments We would like to thank Susanne Knall for her excellent technical assistance during cloning of CarO repeats. This work was supported by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) under Germany’s Excellence Strategy— EXC-2189—Project ID: 390939984 to WW and CEPLAS— EXC—2048/1—Projekt ID 390686111 to MDZ, the iGRAD Plant IRTG 1525 to ROF and MDZ, and a grant to WW (WE 4733/7-1). References 1. Weber W, Fussenegger M (2012) Emerging biomedical applications of synthetic biology. Nat Rev Genet 13(1):21–35 2. Ausl€ander S, Fussenegger M (2016) Engineering gene circuits for mammalian cell–based applications. Cold Spring Harb Perspect Biol 8(7):a023895 3. Tigges M, Marquez-Lago TT, Stelling J et al (2009) A tunable synthetic mammalian oscillator. Nature 457(7227):309–312 4. Kramer BP, Weber W, Fussenegger M (2003) Artificial regulatory networks and cascades for discrete multilevel transgene control in mammalian cells. Biotechnol Bioeng 83 (7):810–820 5. Weber W, Schoenmakers R, Keller B et al (2008) A synthetic mammalian gene circuit reveals antituberculosis compounds. Proc Natl Acad Sci U S A 105(29):9994–9998 6. Xie M, Ye H, Wang H et al (2016) β-cell–mimetic designer cells provide closedloop glycemic control. Science 354 (6317):1296–1301 7. Kojima R, Aubel D, Fussenegger M (2016) Toward a world of theranostic medication: programming biological sentinel systems for therapeutic intervention. Adv Drug Deliv Rev 105:66–76 8. Wagner HJ, Sprenger A, Rebmann B et al (2016) Upgrading biomaterials with synthetic biological modules for advanced medical applications. Adv Drug Deliv Rev 105:77–95 9. Weber W, Fussenegger M (2010) Synthetic gene networks in mammalian cells. Curr Opin Biotechnol 21(5):690–696 10. Mu¨ller K, Naumann S, Weber W et al (2015) Optogenetics for gene expression in mammalian cells. Biol Chem 396(2):145–152 11. Kolar K, Weber W (2017) Synthetic biological approaches to optogenetically control cell signaling. Curr Opin Biotechnol 47:112–119

12. Olson EJ, Tabor JJ (2014) Optogenetic characterization methods overcome key challenges in synthetic and systems biology. Nat Chem Biol 10(7):502–511 13. Kolar K, Knobloch C, Stork H et al (2018) OptoBase: a web platform for molecular optogenetics. ACS Synth Biol 7(7):1825–1828 14. Mu¨ller K, Siegel D, Rodriguez Jahnke F et al (2014) A red light-controlled synthetic gene expression switch for plant systems. Mol BioSyst 10(7):1679–1688 15. Chatelle C, Ochoa-Fernandez R, Engesser R et al (2018) A green-light-responsive system for the control of transgene expression in mammalian and plant cells. ACS Synth Biol 7 (5):1349–1358 16. Ortiz-Guerrero JM, Polanco MC, Murillo FJ et al (2011) Light-dependent gene regulation by a coenzyme B12-based photoreceptor. Proc Natl Acad Sci U S A 108(18):7565–7570 17. Jones AR (2017) The photochemistry and photobiology of vitamin B 12. Photochem Photobiol Sci 16(6):820–834 18. Padmanabhan S, Jost M, Drennan CL et al (2017) A new facet of vitamin B12: gene regulation by cobalamin-based photoreceptors. Annu Rev Biochem 86:485–514 ˜o R, Elı´as19. Padmanabhan S, Pe´rez-Castan Arnanz M (2019) B12-based photoreceptors: from structure and function to applications in optogenetics and synthetic biology. Curr Opin Struct Biol 57:47–55 20. Garcia-Moreno D, Polanco MC, NavarroAviles G et al (2009) A vitamin B12-based system for conditional expression reveals dksA to be an essential gene in Myxococcus xanthus. J Bacteriol 191(9):3108–3119 21. Kainrath S, Stadler M, Reichhart E et al (2017) Green-light-induced inactivation of receptor signaling using cobalamin-binding domains. Angew Chem Int Ed 56(16):4608–4611

Green Light-Controlled Gene Switch 22. Wang R, Yang Z, Luo J et al (2017) B12-dependent photoresponsive protein hydrogels for controlled stem cell/protein release. Proc Natl Acad Sci U S A 114 (23):5912–5917 23. Mu¨ller K, Zurbriggen MD, Weber W (2014) Control of gene expression using a red-and

107

far-red light–responsive bi-stable toggle switch. Nat Protoc 9(3):622–632 24. Jacobs JL, Dinman JD (2004) Systematic analysis of bicistronic reporter assay data. Nucleic Acids Res 32(20):e160–e160

Chapter 7 Sonogenetic Modulation of Cellular Activities in Mammalian Cells Yao-Shen Huang, Ching-Hsiang Fan, Wei-Ting Yang, Chih-Kuang Yeh, and Yu-Chun Lin Abstract Ultrasound is acoustic waves that can penetrate deeply into tissue in a focused manner with limited adverse effects on cells. As such, ultrasound has been widely used for clinical diagnosis for several decades. Ultrasound induces bioeffects in tissues, providing potential value in therapeutic applications. However, the intrinsic millimeter scale of the ultrasound focal zone represents a challenge with respect to minimizing the illuminated regions to perturb target cells in a precise manner. To control a specific cell population or even single cells, sonogenetic tools that combine ultrasound and genetic methods have been recently developed. With these approaches, several ultrasound-responsive proteins are heterologously introduced into target cells, which enhances the cells’ ability to respond to ultrasound stimulation. With optimization of the ultrasound parameters, these tools can specifically manipulate activities in genetically defined cells but not in unmodified cells present in the ultrasound-illuminated regions. These approaches provide new strategies for noninvasive modulation of target cells in various therapeutic applications. Key words Ultrasound, Genetics, Sonogenetics, Prestin, Noninvasive therapy, Microbubbles

1

Introduction Ultrasound (US) acoustic waves, which have frequencies >20 kHz, are inaudible to humans. Owing to its high penetrability, focusing, and safety, US has been widely used in clinical diagnosis and therapy for over 50 years [1–4]. The frequency of US can be adjusted to fulfill different demands. US at higher frequencies results in greater resolution but suffers from lower penetrability. Conversely, US at lower frequencies can penetrate to deeper tissues with lower spatial resolution [5–7]. In addition to defining the physical properties of US, researchers have also studied US-induced bioeffects. Depend-

Yao-Shen Huang, Ching-Hsiang Fan and Wei-Ting Yang contributed equally to this work. Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

109

110

Yao-Shen Huang et al.

ing on the parameters of US used on tissues, US can trigger thermal effects and mechanical effects [8, 9]. The acoustic energy in US-excited tissues is converted to heat and in turn raises the local temperature [10–12]. Besides its thermal effects, the rarefactions and compressions of US longitudinal waves induce mechanical bioeffects due to the propagation of waves through different media [8, 9]. These US-mediated bioeffects have been used in numerous clinical applications including eliminating tumor cells, increasing blood flow, improving dermal contraction, and accelerating the healing rates of injured tissues [13–16]. The bioeffects of US can be further enhanced by nano/microparticles that are responsive to acoustic waves. Among these agents, one of the most commonly used is an US contrast agent, microbubbles (MBs), which are composed of a thin lipid layer filled with air or a high–molecular weight gas [17–20]. As the US waves propagate at different rates in different media, the acoustic waves change the tension of the gas-liquid interface of MBs, resulting in rapid oscillation of the MBs in the incident wave. Under high acoustic pressure, US can trigger MB cavitation accompanied by vibration forces for transiently increasing the permeability of cellular membrane or opening blood–brain barriers, which permits gene or drug delivery as well as manipulating cellular activities under the control of US excitation [21–23]. Moreover, destruction of cells induced by excessive US and MBs can be used to eliminate cancer cells or abnormal tissues [24–26]. Although these approaches are powerful, MBs are not stable in circulation (
5-min turnover rate in the bloodstream) and cannot be used to target extravascular tissues [27]. Even though low-frequency US can noninvasively access deep tissue, its millimeter-scale spatial resolution prevents it from being used to perturb activities of target cells (on the scale of micrometers) [28]. One way to overcome this limitation is by combining US with genetic tools that make the target cells more responsive to US. These emerging new approaches, referred to as sonogenetics, provide a potential solution for using US to manipulate the activities of target cells. Several mechanosensitive ion channels including TRPA1, TREK-1, MscL, Piezo1, and TRP4 have been heterologously expressed in cells, resulting in the ability to use US to perturb genetically defined cells via mechanical stimulation [29– 33]. Moreover, cells transfected with TRPV1, a thermosensitive ion channel, are also activated by US stimulation via a thermal effect [34]. Apart from the ion channels mentioned above, an engineered auditory-sensing protein, Prestin(N7T, N308S) can endow human cells with US sensitivities. US at 0.5 MHz is sufficient to trigger a calcium influx in human cells expressing Prestin(N7T, N308S) [35]. Moreover, the expression of Prestin(N7T, N308S) in deep regions of the mouse brain can respond to transcranial US, resulting in increased c-FOS expression [35, 36]. Here we describe the

Sonogenetic Modulation of Cellular Activities

111

detailed procedures for setting up an US apparatus, engineering Prestin protein, genetically modifying target cells in culture and cells within the mouse brain via adeno-associated virus or MBs, and illuminating the target cells with US to modulate cell activities.

2 2.1

Materials US Apparatus

1. Single-element focused US transducer (V389; Olympus, Tokyo, Japan). 2. Waveform the USA).

generator

(AFG3251;

Tektronix,

Oregon,

3. Radio-frequency power amplifier (325LA; Electronics & Innovation, New York, the USA). 4. Calibrated polyvinylidene difluoride type hydrophone (model HGL-0085, ONDA, Sunnyvale, the USA; calibration range ¼ 0.25–20 MHz; spatial resolution: 85 μm). 5. Oscilloscope (LT354; LeCory Co., Chestnut Ridge, the USA). 2.2 Live-Cell Imaging System

1. Inverted fluorescence microscope. 2. 20 objective (NA ¼ 0.75). 3. 60 oil objective (NA ¼ 1.4). 4. Environmental chamber. 5. CMOS camera. 6. NIS-Elements AR software (Nikon, Tokyo, Japan).

2.3 Cell Culture and Transfection

1. HEK29T cells (Human embryonic kidney cells). 2. Culture medium: Dulbecco’s modified Eagle’s medium (DMEM), 10% (v/v) fetal bovine serum (FBS), penicillin (5 U/mL), streptomycin (50 μg/mL) (see Note 1). 3. Phosphate-buffered saline (PBS): pH 7.4. 4. Trypsin-EDTA solution. 5. 37  C CO2 incubator. 6. Hemocytometer. 7. Poly-D-lysine. 8. 25-mm glass coverslip. 9. Six-well culture plates. 10. LT-1 transfection reagent. 11. Opti-MEM. 12. Five-chamber cell-stack (CF5). 13. Polyethylenimine (PEI 25000).

112

Yao-Shen Huang et al.

Table 1 The primers used to generate mutated prestin plasmid DNA Primer name

Sequence (50 –30 )

N7T- Forward

AGACCATGGATCATGCTGAAGAAACCGAAATCCC TGCAGAGACCCAGAGG

N7T- Reverse

CCTCTGGGTCTCTGCAGGGATTTCGGTTTCTTCAGCATGATCCATGGTC T

N308S- Forward GGGACTGGCATTTCTGCAGGATTTTCCCTACATGAGTCCTACAGTGTGGA N308S- Reverse

TCCACACTGTAGGACTCATGTAGGGAAAATCCTGCAGAAATGCCAGTCCC

14. 0.45-μm polyethersulfone (PES) filter. 15. PEG solution (Polyethylene glycol 8000). 16. PBS/pluronic F68/NaCl solution: PBS, 0.001% (v/v) pluronic F68, 200 mM NaCl. 17. Benzonase. 18. ATP (10 μM). 2.4 Construction of Plasmid DNAs

1. CFP-R-GECO plasmid DNA is obtained from Dr. Takanari Inoue at Johns Hopkins University School of Medicine. 2. Venus-Prestin (NM 001289787) was obtained from Dr. Jian Zuo at St. Jude Children’s Research Hospital. 3. Restriction enzymes: HindIII, XbaI, XhoI, BamHI, DpnI. 4. Primers (see Table 1). 5. dNTPs (10 mM). 6. 5 Phusion HF Buffer. 7. Phusion DNA Polymerase. 8. Thermocycler.

2.5 Preparation of MBs

1. 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC). 2. 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[carboxy(polyethylene glycol)-2000] (DSPE-PEG 2000). 3. 1,2-dipalmitoyl-3-trimethylammonium-propane (DPTAP). 4. Perfluoropropane (C3F8) gas. 5. Multisizer 3 Coulter counter.

2.6 Animal Experiments

1. Mouse: C57BL/6jNarl mice (age: 7–10 weeks; weight: 30–35 g). 2. Anesthetization: mixture of Zoletil 50 and Rompun 2%, mixture of oxygen (0.8 L/min) and 2% vaporized isoflurane. 3. Stereotaxic instrument.

Sonogenetic Modulation of Cellular Activities

113

4. Surgical instruments. 5. Needles: small needle (27 gauge), steel needles (33 gauge). 6. Injector-attached syringe. 7. US coupling gel. 2.7 Immunohistochemical Staining

1. Paraformaldehyde (4%). 2. Optimum cutting temperature compound. 3. Freezing microtome. 4. 12-well plate. 5. Blocking solution: PBS containing 5% normal goat serum. 6. Antibodies: anti-c-Fos (1:1000), anti-tyrosine hydroxylase (1:500), DyLight 594-conjugated secondary antibody (1:500). 7. 40 ,6-diamidino-2-phenylindole (DAPI). 8. Fluorescence mounting medium.

3

Methods

3.1 Introducing Two Mutants into Mouse Prestin DNA

Generate Venus-tagged mouse Prestin mutant genes via sitedirected mutagenesis. The sequences of all the primers used in this protocol are listed in Table 1. 1. Set up a PCR reaction mix with each PCR tube containing the following: 0.5 μL (1 μg/μL) template Venus-mPrestin (wildtype) plasmid DNA, 0.5 μL (10 μM) forward primer, 0.5 μL (10 μM) reverse primer (Table 1), 0.5 μL dNTPs (10 mM), 5 μL Phusion 5 HF Buffer, 0.25 μL Phusion DNA Polymerase, and sterilized ddH2O to 25 μL. Mix gently by pipetting and collect all liquid in the bottom of the tube with a quick spin (see Note 2). 2. Transfer the tubes to a thermocycler with the following cycling conditions: 30 s at 98  C for the initial denaturation; 16 cycles of 30 s at 98  C, 30 s at 57  C, and 10 min at 72  C for amplification; 10 min at 72  C for the final extension; 4  C for cooling down. Store the sample at 4  C if necessary. 3. To remove the original template, add 2.5 μL DpnI into each PCR tube and incubate at 37  C for 1 h. These DNA fragments could be amplified by bacterial transformation. Verify the plasmid via DNA sequencing before being used in subsequent steps.

3.2

DNA Transfection

1. Add 50 μL poly-D-lysine (0.1 mg/mL) onto the central area of a 25-mm glass coverslip in a six-well culture plate and incubate at 37  C in 5% CO2 for 20 min. Remove the poly-D-lysine from

114

Yao-Shen Huang et al.

the coverslip and add 100 μL sterilized ddH2O to wash the central area of the coverslip. Use an aspirator to remove this wash solution. 2. Mix together CFP-R-GECO, Venus-Prestin plasmid DNAs (1.5 μg for each DNA) and LT-1 transfection reagent at a ratio of 1:3 DNA (μg) and transfection reagent (μL) in 40 μL Opti-MEM. Mix this transfection mixture thoroughly by pipetting and incubate at room temperature for 20 min (see Note 3). 3. Remove the culture medium with an aspirator and add 5 mL sterile PBS to gently wash HEK293T cells grown in a 10-cm culture dish to 60–80% confluence (see Note 4). Remove the PBS and harvest the cells by adding 1 mL trypsin-EDTA solution and then incubate the cells for 3–5 min at 37  C. Add 4 mL fresh DMEM to stop trypsinization and resuspend cells. Collect the cell suspension in a 15-mL tube. Use 10–15 μL of the cell suspension to count the cells on a hemocytometer, counting cell number for transfection. Collect the required number of cells (2.5  105 cells) into another tube and centrifuge at 100 centrifugal force (g) for 5 min. Replace the supernatant with fresh DMEM containing 10% FBS (500 μL medium/2.5  105 cells) and resuspend by pipetting. 4. Add the transfection mixture (step 2) to the prepared HEK293T cells (2.5  105 cells in 500 μL medium) and pipette gently. Seed 80 μL of cells onto each poly-D-lysine– coated 25-mm glass coverslip in the six-well plate (step 1) and then incubate the cells at 37  C in 5% CO2 for 1 h (see Note 5). After the incubation, add 2 mL DMEM containing 10% FBS to each well and incubate the cells at 37  C in 5% CO2 for 18–24 h prior to US stimulation. 3.3 Preparation of DNA-Loaded MBs

1. The MBs are prepared via the thin-film hydration method (Fig. 1) [38]. Three phospholipids, 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-distearoyl-sn-glycero-3phosphoethanolamine-N-[carboxy(polyethylene glycol)2000] (DSPE-PEG 2000), and 1,2-dipalmitoyl-3-trimethylammonium-propane (DPTAP) are dissolved in chloroform at a weight ratio of 9:1:1 in an airtight vial (Fig. 1). 2. The chloroform is then removed by evaporation such that a thin film of lipids remained on the wall of the vial (Fig. 1). 3. To dissolve the thin film, 1 mL of degassed PBS containing 1% (w/v) glycerol is added to the vial (Fig. 1). 4. The vial is placed in a sonicator for 5 min to disperse the lipid solution at room temperature (Fig. 1).

Sonogenetic Modulation of Cellular Activities

115

Fig. 1 Flow chart of DNA-loaded MBs preparation. Red arrows represent adding required materials into the vial

5. The vial is degassed and refilled with perfluoropropane (C3F8) gas (Fig. 1). 6. The MB suspension is then produced by intense shaking using a home-built agitator (Fig. 1). 7. The MB suspension is centrifuged for 2 min at 500  g, and the top layer is transferred to 1 mL of ddH2O in a clean vial (Fig. 1). 8. The MB suspension is gently dispersed and counted as well as sized using a Multisizer 3 Coulter counter to characterize their size distribution and particle concentration. 9. The DNA (7500 ng) of interest is gently mixed into the MB solution (concentration: 108 MBs/mL; volume: 0.1 mL) for 30 min. The DNA will naturally attach to the shell of MBs by electrostatic interaction (Fig. 1) [39].

116

Yao-Shen Huang et al.

10. The MB suspension is then centrifuged for 2 min at 500  g, and the top layer is transferred to 1 mL of ddH2O in a clean vial to remove unreacted DNA (Fig. 1). 3.4 AdenoAssociated Virus Production

1. HEK293T cells are seeded into a five-chamber cell-stack (CF5) at a concentration of 2.5  108 cells/CF5 and are incubated at 37  C in 5% CO2 for 24–36 h. 2. Prepare each plasmid DNA (RepCap, pHelper, and transfer plasmid) at a concentration of 1 μg/μL and calculate the amount of each plasmid needed to have a 1:1:1 molar ratio with 2.5 mg total DNA for each CF5. 3. Add the plasmid DNAs to a sterile 250-mL bottle containing 176 mL Opti-MEM and mix well. Add 7.5 mL of Polyethylenimine (1:3 DNA (μg)/Polyethylenimine (μL) ratio) to the bottle. Shake the bottle up-and-down for 30 s and incubate the bottle at room temperature for 15 min. During the incubation, prepare a sterile 500-mL bottle containing 350 mL DMEM and 2% FBS. At the end of the incubation period, add the transfection mixture to the bottle containing the DMEM and mix well. 4. Remove the medium from the CF5, gently add the transfection mixture to the CF5, and then incubate the cells at 37  C in 5% CO2 for 96 h. 5. Harvest the cells and medium by tapping the sides of the CF5 on a soft surface, then transfer cells and medium into multiple 50-mL tubes. Add 20 mL PBS to the CF5 to collect additional cells and transfer these cells into a new 50-mL tube. Centrifuge the 50-mL tubes at 1000  g at 4  C for 10 min. Transfer the supernatant to a sterile 500-mL bottle and keep the pellet on ice (this is the pellet referred to below in step 8). 6. Filter supernatant through a 0.45-μm polyethersulfone (PES) filter and add 25 mL PEG solution (40% [w/v] polyethylene glycol 8000 solution in ddH2O) for each 100 mL supernatant. Use a stir bar to stir slowly at 4  C for 1 h, and then hold at 4  C for 3 h without stirring to allow full precipitation. 7. Transfer the entire sample to 50-mL tubes and centrifuge at 2818  g at 4  C for 15 min. Remove the supernatant and completely resuspend the pellet in 10 mL PBS/pluronic F68/NaCl solution. Keep the resuspended pellet on ice. 8. Add 10 mL of PBS/pluronic F68/NaCl solution to resuspend the cell pellet (from step 5), lyse the pellet with four 1-s sonication pulses (50% amplitude) with at least 15 min on ice between each pulse. Centrifuge the cell pellet at 3220  g at 4  C for 15 min. Transfer the lysate to the tube containing the resuspended virus from step 9 (below).

Sonogenetic Modulation of Cellular Activities

117

9. To degrade any residual DNA, add benzonase at 50 U/mL of viral suspension in PBS (step 7) and incubate at 37  C for 45 min. Transfer the viral suspension to centrifuge tubes and centrifuge at 2415  g at 4  C for 10 min. Then transfer the clarified supernatant to a new tube. 10. Store the viral solution at 80  C for long-term storage (see Note 6). 3.5 Acoustic Peak Negative Pressure and US Focal Zone Measurement

1. For acoustic peak negative pressure measurement, the US transducer is mounted on a three-axis stage and the focus point of the US transducer is collimated at the tip of a calibrated polyvinylidene difluoride type hydrophone. The hydrophone is connected with an oscilloscope for collecting the voltage traces produced by US pressure waves (Fig. 2). 2. The acoustic peak negative pressure then could be obtained via multiplying the recorded voltage value and hydrophone sensitivity value (14.5 kPa/mV). 3. For US focal zone measurement, the hydrophone measurement is conducted with spot scanning (width: 25 mm, length: 25 mm). The scanning interval is set at 1 mm. The measurements are conducted in an acrylic water tank that is filled with distilled and degassed water at 25  C.

Fig. 2 Illustrations of acoustic peak negative pressure measurement and US focal zone measurement. A hydrophone was placed in front of an US transducer to measure the acoustic pressure of the US generated from the transducer. To estimate the focal zone of US, the US transducer was mounted on a three-axis stage and performed spot scanning (width: 1 mm of step interval for 25 mm; length:1 mm of step interval for 25 mm)

118

Yao-Shen Huang et al.

3.6 In Vitro US Stimulation

1. US stimulation (Frequency: 0.5 MHz; acoustic negative pressure, 0.5 MPa; 2000 cycles; pulse repetition frequency, 10 Hz; 3-s duration) is applied using a single-element FUS transducer. The US transducer is driven by a waveform generator and a radio-frequency power amplifier to transmit the US pulses (Fig. 3). 2. A water cone filled with degassed water is attached to the US transducer assembly, after which the surface of the cone is submerged into the culture dish medium (Fig. 3). 3. Before starting each experiment, we use a glass-bottom dish which has been embedded two acoustically transparent vesselmimicking cellulose tubes (200 μm in diameter) in a cross shape. The point of intersection of the two tubes is positioned at the center of the dish (0.5 cm above the bottom). The point of intersection is also marked onto the bottom of the glass dish by a colored pen. 4. To align the focus of the US transducer and optical objective, the location of the holder is first appropriately adjusted such that the mark made in step 3 appeared at the center of the objective.

Fig. 3 Schematic representation of the live-cell imaging and US-exposure equipment. The cells are seeded on a cover glass held by an Attofluor Cell Chamber (4). The environmental chamber (3) provides an enclosed space to ensure cell imaging under physiological conditions (temperature, 37  C; humidity, >90% relative humidity; 5% CO2). A water cone (2) filled with degassed water is attached to the US transducer assembly (1), after which the bottom surface of the cone is lowered into the culture dish medium. The US waves are generated by a transducer (1) that is driven by a waveform generator and amplifier. The effects of US stimulation on cells can be simultaneously observed by an inverted microscope

Sonogenetic Modulation of Cellular Activities

119

5. Use the US transducer in pulse echo mode and positioning the location of the US transducer more precisely, we ensure that the focal area of the US pulse is sonicated at the point of intersection of the two tubes. 6. We then move the location of the US transducer down 0.5 cm. 7. We verify the successful positioning optical and acoustic foci by injecting MBs into the cellulose tube and observing the occurrence of MBs destruction by US (see Note 7). 8. To record the calcium influx in cells in real-time, we replace the glass dish with a culture dish containing the cells of interest. 3.7 Live-Cell Imaging and Data Analysis

1. Transfer HEK293T cells cultured on poly-D-lysine–coated 25-mm glass coverslips in six-well plates to Attofluor Cell Chambers immersed in serum-free DMEM without phenol red, and then hold these cells under a 5% CO2 atmosphere at 37  C in an environmental chamber (Fig. 3). 2. Conduct time-lapse imaging throughout the procedure (Fig. 4). Expose cells to US at 30 s for 3 s and add ATP at 210 s to induce a calcium response in the cells as a positive control (see Notes 8 and 9). 3. Carry out image analysis (i.e., determine the probability of US-excitable cells) with NIS-Elements AR software. Count the number of CFP and YFP double-positive signals, which represents the total cell number. 4. Time-dependent normalized fluorescence intensity (F/F0) is used for the probability analysis. Measure the average intensity of the R-GECO signal at 0–30 s (gray region in Fig. 4) as the initial fluorescence intensity (F0). After US stimulation, use the maximum intensity of R-GECO from the period highlighted in yellow (Fig. 4) as (F). A value of F/F0 that is 1.2 is defined as US-excitable cells. 5. Calculate the percentage of US-excitable cells relative to the total number of cells as determined by CFP and YFP fluorescence (step 3) (see Note 10).

Fig. 4 Time-lapse imaging of live cells is performed according to this procedure. Focused US (FUS) is applied to stimulate cells at 30 s for 3 s. ATP treatment as a calcium response inducer is used as a positive control to confirm R-GECO activity in HEK293T cells. For data analysis, the gray shading represents the baseline of the R-GECO intensity. Fluorescence intensity is then recorded continuously from 30 to 150 s (yellow shading)

120

Yao-Shen Huang et al.

3.8 In Vivo Gene Delivery 3.8.1 Adeno-Associated Virus Infection

1. To anesthetize a mouse before surgery, inject the mouse intraperitoneally with a 1:1 mixture (1.7 mL/kg) of Zoletil 50 and Rompun 2%. 2. Place the mouse in a stereotaxic apparatus. 3. Shave the fur over the skull region and cut the skin to expose the skull (width: 15 mm, length: 15 mm). 4. Keep the wound open with small hooks and thin the skull with a hand-held drill (bregma, 3 mm; left, 0.5 mm). 5. Stop the drill when the skull is perforated. Take a small needle and carefully perforate the dura matter. Edges of the craniotomy. 6. Draw up 1 μL of the adeno-associated virus solution (refer Subheading 3.4) into an injector with a steel needle. The injector is attached to a syringe (KD Scientific Inc., Holliston, the USA). The syringe is assembled on a stereotaxic arm. 7. Bring the tip of the microneedle to the correct x and y position and lower it until it touches the exposed dura. 8. After the microneedle penetrates the dura, slowly lower it to the desired z coordinate of the injection site (ventral tegmental area, VTA; bregma, 3 mm; left, 0.5 mm; depth, 4.2 mm), and wait 5 min before injection. 9. Inject a total of 1 μL virus solution at a rate of 0.5 μL/min. 10. Wait 10 min and then withdraw the microneedle slowly to avoid backflow of the virus solution to the surface. 11. Suture the skin and apply triple antibiotic ointment to the wound. Two weeks later, mice are exposed to transcranial US stimulation.

3.8.2 Sonotransfection

1. Anesthetize a mouse with a 1:1 mixture (1.7 mL/kg) of Zoletil 50 and Rompun 2%. 2. Secure the animal on a stereotaxic apparatus with ear bars and a bite bar. 3. Shave the fur over the skull region and cut the skin to expose the skull (width: 15 mm, length: 15 mm). 4. Attach a water cone filled with degassed water to an US transducer assembly. 5. Move the US transducer to the target site (
3.0 mm posterior to the bregma and 0.5 mm lateral to the sagittal suture, 4.2 mm below the skull surface; left hemisphere of the brain). 6. Apply US coupling gel between the cone surface and the head of the mouse. 7. Inject 100 μL of MB solution (1  108 DNA-loaded MBs/mL; DNA, 7500 ng; refer to Subheading 3.3) retro-orbitally.

Sonogenetic Modulation of Cellular Activities

121

8. Wait 20 s for MBs circulating into brain and deliver US (frequency: 1 MHz; acoustic peak negative pressure, 0.5 MPa; 5000 cycles; pulse repetition frequency, 5 Hz; 240-s duration) to the VTA. Two days later, mice are exposed to transcranial US stimulation. 3.9 In Vivo US Stimulation

1. Anesthetize a mouse with a mixture of oxygen (0.8 L/min) and 2% vaporized isoflurane. 2. Secure the gene delivered mouse which has undergone one of the gene delivery methods described in Subheading 3.8 on a stereotaxic apparatus with ear bars and a bite bar. 3. Attach a water cone filled with degassed water to an US transducer assembly. 4. Move the US transducer to the sonication site (
3.0 mm posterior to the bregma and 0.5 mm lateral to the sagittal suture, 4.2 mm below the skull surface; left hemisphere of the brain). 5. Apply US coupling gel between the cone surface and the head of the mouse. 6. Deliver US into VTA region. (Frequency: 0.5 MHz; acoustic peak negative pressure, 0.5 MPa; 2000 cycles; pulse repetition frequency, 10 Hz; 3-s duration). Nighty min later, sacrifice mice immediately and conduct immunohistochemical staining (Subheading 3.10).

3.10 Immunohistochemical Staining

1. Sacrifice mice which have undergone US stimulation and perfuse with 4% paraformaldehyde via the left ventricle until the right atrium appear colorless perfusion fluid. 2. Remove brain tissues, embed samples rapidly in optimum cutting temperature compound, and store at 50  C. 3. Slice tissues into coronal sections (thickness, 15 μm) on a freezing microtome. Collect the sections into a 12-well plate containing 2 mL 0.1 M PBS. 4. Incubate these sections in 5% goat serum in PBS for 1 h to minimize the background signal due to non-specific antibody binding. 5. Incubate the sections in antibody against c-Fos (1:1000) or antibody against tyrosine hydroxylase (1:500) in blocking solution overnight at 4  C. 6. Incubate the sections with DyLight 594–conjugated secondary antibody (1:500) in PBS for 1 h at 37  C. 7. Is the sections with PBS for three times and label the cellular nuclei by DAPI (1:10000) for 10 min at room temperature. 8. Use fluorescence mounting medium to fix the slides with and coverslips and stored flat in the dark at 20  C.

122

4

Yao-Shen Huang et al.

Notes 1. All the FBS mentioned here is inactivated by heating to 56  C for 30 min. Heat-Inactivated FBS can be purchased If you want to avoid the heating process in water bath. 2. If you have multiple reactions (1 tube for one reaction) in one PCR process, prepare a premix tube containing reagents you need, gently vortex and spin down, then aliquot the premixed PCR mixture to PCR tubes to minimize the pipetting error. 3. Vortex DNA solution and pipetting thoroughly before drawing in. 4. Before transfection, prewarm solution at 37  C in the water bath for 10–15 min, including DMEM, PBS, trypsin, poly-Dlysine, and Opti-MEM. 5. If you need to seed transfected-cell solution in a volume more than 80 μL onto coverslip, increase the volume of poly-D-lysine for covering more area of coverslip. 6. Viral solution can be store at 4  C for few days. Please avoid to repeatedly freeze and thaw the virus, it will reduce the infection efficiency. 7. After using MBs to verify FUS excitation, clean the cone by washing with ddH2O, make sure that there are no MBs remained on the cone. 8. ATP addition should be gentle to avoid detaching cells from surface and disturbing imaging analysis. 9. The Attofluor Cell Chamber should be thoroughly cleaned by washing with detergent after each imaging experiment. 10. Unhealthy cells should be excluded for analysis.

Acknowledgments We thank Dr. Jian Zuo at St. Jude Children’s Research Hospital for the mouse Prestin construct and Dr. Takanari Inoue at Johns Hopkins University School of Medicine for the R-GECO construct. This study was supported by the Ministry of Science and Technology (MOST), Taiwan under Grant No. 108-2636-B-007003, 109-2636-B-007-003, 108-2638-B-010-001-MY2, 108-2221-E-007-041-MY3, and 108-2221-E-007-040-MY3. Additional funding consisted of a grant from National Tsing Hua University under Grant No. 109Q2511E1.

Sonogenetic Modulation of Cellular Activities

123

References 1. Robinson L (1997) Diagnostic ultrasound: a primary care-led service? Br J Gen Pract 47:293–296 2. Brown AK (2009) Using ultrasonography to facilitate best practice in diagnosis and management of RA. Nat Rev Rheumatol 5:698–706 3. Fini M, Tyler WJ (2017) Transcranial focused ultrasound: a new tool for non-invasive neuromodulation. Int Rev Psychiatry 29:168–177 4. Karaboce B (2016) Investigation of thermal effect by focused ultrasound in cancer treatment. IEEE Instrum Meas Mag 19:20–64 5. Fregni F, Pascual-Leone A (2007) Technology insight: noninvasive brain stimulation in neurology—perspectives on the therapeutic potential of rTMS and tDCS. Nat Clin Pract Neurol 3:383–393 6. Kubanek J, Shi J, Marsh J et al (2016) Ultrasound modulates ion channel currents. Sci Rep 6:1–14 7. Tyler WJ, Tufail Y, Finsterwald M et al (2008) Remote excitation of neuronal circuits using low-intensity, low-frequency ultrasound. PLoS One 3:e3511 8. Dalecki D (2004) Mechanical bioeffects of ultrasound. Annu Rev Biomed Eng 6:229–248 9. O’Brien WD (2007) Ultrasound-biophysics mechanisms. Prog Biophys Mol Biol 93:212–255 10. Speed CA (2001) Therapeutic ultrasound in soft tissue lesions. Rheumatology 40:1331–1336 11. Busse JW, Bhandari M, Kulkarni AV (2002) The effect of low-intensity pulsed ultrasound therapy on time to fracture healing: a metaanalysis. CMAJ 166:437–441 12. Kim YS, Rhim H, Min JC (2008) Highintensity focused ultrasound therapy: an overview for radiologists. Korean J Radiol 9:291–302 13. Wood AKW, Sehgal CM (2015) A review of low-intensity ultrasound for cancer therapy. Ultrasound Med Biol 41:905–928 14. O’Reilly MA, Hynynen K (2015) Emerging non-cancer applications of therapeutic ultrasound. Int J Hyperth 31:310–318 15. Piper RJ, Hughes MA, Moran CM (2016) Focused ultrasound as a non-invasive intervention for neurological disease: a review. Br J Neurosurg 30:286–293 16. Gutowski KA (2016) Microfocused ultrasound for skin tightening. Clin Plast Surg 43:577–582

17. Fry FJ, Sanghvi NT, Foster RS (1995) Ultrasound and microbubbles: their generation, detection and potential utilization in tissue and organ therapy-experimental. Ultrasound Med Biol 21:1227–1237 18. Sakakima Y, Hayashi S, Yagi Y et al (2005) Gene therapy for hepatocellular carcinoma using sonoporation enhanced by contrast agents. Cancer Gene Ther 12:884–889 19. Zolochevska O, Xia X, Williams BJ et al (2011) Sonoporation delivery of interleukin-27 gene therapy efficiently reduces prostate tumor cell growth in vivo. Hum Gene Ther 22:1537–1550 20. Alkins R, Burgess A, Ganguly M et al (2014) Focused ultrasound delivers targeted immune cells to metastatic brain tumors. Cancer Res 73:276–285 21. Lentacker I, DeCock I, Deckers R (2014) Understanding ultrasound induced sonoporation: definitions and underlying mechanisms. Adv Drug Deliv Rev 72:49–64 22. Chu PC, Liu HL, Lai HY et al (2015) Neuromodulation accompanying focused ultrasound-induced blood-brain barrier opening. Sci Rep 5:1–12 23. Fan CH, Huang YS, Huang WE et al (2017) Manipulating cellular activities using an ultrasound-chemical hybrid tool. ACS Synth Biol 6:2021–2027 24. Mitragotri S (2005) Healing sound: the use of ultrasound in drug delivery and other therapeutic applications. Nat Rev Drug Discov 4:255–260 25. Goertz DE, Todorova M, Mortazavi O et al (2012) Antitumor effects of combining docetaxel (Taxotere) with the Antivascular action of ultrasound stimulated microbubbles. PLoS One 7:e52307 26. Lin CY, Tseng HC, Shiu HR et al (2012) Ultrasound sonication with microbubbles disrupts blood vessels and enhances tumor treatments of anticancer nanodrug. Int J Nanomedicine 7:2143–2152 27. Sirsi S, Borden M (2009) Microbubble compositions, properties and biomedical applications. Bubble Sci Eng Technol 1:3–17 28. Ng A, Swanevelder J (2011) Resolution in ultrasound imaging. Contin Educ Anaesthesia Crit Care Pain 11:186–192 29. Ibsen S, Tong A, Schutt C et al (2015) Sonogenetics is a non-invasive approach to activating neurons in Caenorhabditis elegans. Nat Commun 6:8264

124

Yao-Shen Huang et al.

30. Ye J, Tang S, Meng L et al (2018) Ultrasonic control of neural activity through activation of the mechanosensitive channel MscL. Nano Lett 18:4148–4155 31. Pan Y, Yoon S, Sun J et al (2018) Mechanogenetics for the remote and noninvasive control of cancer immunotherapy. Proc Natl Acad Sci U S A 115:992–997 32. Qiu Z, Guo J, Kala S et al (2019) The mechanosensitive Ion Channel Piezo1 significantly mediates in vitro ultrasonic stimulation of neurons. iScience 21:448–457 33. Oh SJ, Lee JM, Kim HB et al (2019) Ultrasonic neuromodulation via astrocytic TRPA1. Curr Biol 29:3386–3401.e8 34. Yang Y, Pacia CP, Ye D et al (2020) Sonogenetics for noninvasive and cellular-level neuromodulation in rodent brain. bioRxiv 1–33. https://doi.org/10.1101/2020.01.28. 919910

35. Huang YS, Fan CH, Hsu N et al (2020) Sonogenetic modulation of cellular activities using an engineered auditory-sensing protein. Nano Lett 20:1089–1100 36. Wu CY, Fan CH, Chiu NH et al (2020) Targeted delivery of engineered auditory sensing protein for ultrasound neuromodulation in the brain. Theranostics 10:3546–3561 37. Zhao Y, Araki S, Wu J et al (2011) An expanded palette of genetically encoded Ca2+ indicators. Science 333:1888–1891 38. Smith DA, Porter TM, Martinez J et al (2007) Destruction thresholds of echogenic liposomes with clinical diagnostic ultrasound. Ultrasound Med Biol 33:797–809 39. Wang DS, Panje C, Pysz MA et al (2012) Cationic versus neutral microbubbles for gene delivery in cancer. Radiology 264:721–732

Chapter 8 Constructing Smartphone-Controlled Optogenetic Switches in Mammalian Cells Yuanhuan Yu, Guiling Yu, and Haifeng Ye Abstract With the increasing indispensable role of smartphones in our daily lives, the mobile health care system coupled with embedded physical sensors and modern communication technologies make it an attractive technology for enabling the remote monitoring of an individual’s health. Using a multidisciplinary design principle coupled with smart electronics, software, and optogenetics, the investigators constructed smartphone-controlled optogenetic switches to enable the ultraremote-control transgene expression. A custom-designed SmartController system was programmed to process wireless signals from smartphones, enabling the regulation of therapeutic outputs production by optically engineered cells via a far-red light (FRL)-responsive optogenetic interface. In the present study, the investigators describe the details of the protocols for constructing smartphone-controlled optogenetic switches, including the rational design of an FRL-triggered transgene expression circuit, the procedure for cell culture and transfection, the implementation of the smartphone-controlled far-red light-emitting diode (LED) module, and the reporter detection assay. Key words Mammalian synthetic biology, Optogenetics, Smartphone-controlled cells, Far-red light, Light-controlled designer cells

1

Introduction Telecommunication technology has significantly advanced the development of smartphones used for wireless mobile digital devices, which enables virtual online communication in real time, especially for those living in remote areas that are far from healthcare facilities. Since smartphones have the capability to interface with peripheral devices, smartphone-based mobile health (mHealth) systems has been applied to many biomedical areas, in which the patient health-related data obtained by the smartphone can be sent over the internet to their physician for a detailed investigation [1–3]. Recently, smartphone-based technologies have been reported for the wireless control of transgene expression in mammalian cells or animals [4].

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

125

126

Yuanhuan Yu et al.

Meanwhile, strategies based on optogenetics have recently attracted attention as versatile tools with high spatiotemporal resolution in many biomedical applications, and used for nerve stimulation [5, 6], transgene expression [7–9], genome activation [10, 11], gene editing [12, 13], and cell migration [14]. Several optogenetic switches responsive to UV (~360 nm) [15], blue light (~440 nm) [5–8], and near-infrared light (700–1400 nm) [9, 13, 16] have been developed to remotely control the transgene expression in mammalian cells or animals. Compared with short wavelengths of light sources, such as UV and blue light, far-red light (FRL, ~730 nm) is an ideal optogenetic inducer due to the high biocompatibility and deep tissue penetration (above 5 mm beneath the surface of the skin, much deeper than the penetration of UV and blue light) [17, 18]. With this regard, the investigators constructed an FRL-responsive optogenetic switch to control the transgene expression in mammalian cells. In order to achieve the ultraremote control of transgene expression, as inspired by mHealth, and to integrate telecommunication technology with optogenetics, the present smartphonecontrolled FRL-responsive genetic switch can respond to external signals from the smartphone app to induce a therapeutic output expression. This smartphone-controlled optogenetic switch contains five modules: (1) smartphone; (2) ECNU-TeleMed application (app); (3) custom-designed intelligent electronic home server box “SmartControl-Box”; (4) far-red light-emitting diode (730 nm LED) array controlled by the SmartControl-Box; (5) designer cells containing the FRL-responsive genetic switch. In order to achieve the ultraremote control of transgene expression through the smartphone, the ECNU-TeleMed app was developed to regulate the SmartControl-Box via the Global System for Mobile Communications (GSM) network. When the smartphone was connected to the internet, the designer cells, which contain a FRL-triggered genetic switch, were set as the receiver units for smartphone-transmitted FRL signals from the SmartControllBox, which translates the smartphone-controlled electronic commands into biological responses [4] (Fig. 1a). The core technology of this smartphone-controlled transgene expression system is a bacteriophytochrome-based optogenetic switch, which has three critical components: light receptor BphS (a bacterial light-activated cyclic diguanylate monophosphate [c-diGMP] synthase), YhjH (a c-di-GMP–specific phosphodiesterase to control intracellular c-di-GMP homeostasis), and BldD (a c-di-GMP–binding domain derived from Streptomyces coelicolor’s transcription factor; tetrameric c-di-GMP binds to the C-terminal domains of two BldD monomers to induce protein dimerization, which enables BldD to bind to its target DNA sequence whiG). These were fused to a transactivation domain p65-VP64 to form a hybrid transactivator p65-VP64-BldD. Upon FRL illumination,

Smartphone-Controlled Optogenetic Switches

127

Fig. 1 The smartphone-controlled transgene expression in mammalian cells. (a) The schematic of the customdesigned smartphone app for controlling the SmartControl-Box activities. The ECNU-TeleMed app was programmed to regulate the activity of the custom-made far-red LED array through the SmartControl-Box, enabling FRL-responsive designer cells to initiate the SEAP reporter expression. (b) The schematic of the FRL-triggered mammalian designer cells. The far-red light activates the engineered BphS to produce c-di-GMP, and triggers the hybrid transactivator p65-VP64-BldD dimerization, followed by the binding to its specific chimeric promoter PFRL to drive the SEAP expression. (The pictures were adapted and modified from Shao et al. [4])

the photoreceptor BphS could convert intracellular guanylate triphosphate (GTP) into c-di-GMP. The increase in cytosol c-di-GMP enabled the hybrid transactivator p65-VP64-BldD to translocate into the nucleus, and bind to its chimeric promoter (PFRL containing whiG DNA operator sequences), thereby initiating the transgene expression (Fig. 1b) [4]. In this section, the investigators will

128

Yuanhuan Yu et al.

describe the detailed procedures for constructing smartphonecontrolled genetic switches combined with telecommunication technology with optogenetics. This smartphone-controlled optogenetic device enables the adaptation of the SmartController concept toward any kind of diseases, and might boost the progress of personalized, digitalized, and globalized precision medicines into clinics.

2

Materials

2.1 Construction of the Smartphone Controlled FRL-Responsive Module

1. Intelligent remote controller: The intelligent remote controller purchased from Smart Home Studio consisted of four core modules: a wireless signal processor, an embedded microprocessor unit (MPU) STM32, six channels of independent output relay drivers, and an AC/DC converter (Fig. 2).

Fig. 2 The schematic circuit diagram for the intelligent remote controller modules

Smartphone-Controlled Optogenetic Switches

129

Fig. 3 The images for the switching power supply. The red line indicates the key modules of the switching power supply

2. Switching power supply: The switching power supply (Yunlifang Studio) contains a liquid crystal display (LCD) screen, an output voltage adjustment switch, an output current adjustment switch, an input interface, and an output interface (Fig. 3). 3. Smartphone ECNU-TeleMed app: The ECNU-TeleMed app has been developed to remotely control the activity of SmartControl-Box-connected electronic devices. This app enables the adjustment of light illumination time and intensities. 4. Far-red LED (730 nm). 5. Printed circuit board (PCB). 2.2 Construction of the Plasmids

1. PCR thermal cyclers.

2.2.1 Instruments

3. The gel imaging system.

2. The horizontal agarose gel electrophoresis system. 4. Ultramicro spectrophotometer. 5. Constant temperature incubator. 6. The electric-heated thermostatic water bath. 7. Shaking incubator. 8. Centrifuge. 9. Plastic petri dish (diameter: 9.0 cm). 10. Centrifuge tubes (0.2, 1.5, 2 and 15 mL).

2.2.2 Reagents

1. Polymerase Chain Reaction (PCR) Kit. 2. Primers (oligonucleotides, chemically synthesized by the gene synthesis company). 3. Agarose. 4. 1  TAE buffer: 24.2 g/L Tris, 5.71 mL/L glacial acetic acid, 10 mL/L 50 mM EDTA.

130

Yuanhuan Yu et al.

5. GoldView: nucleic acid dye, final concentration of usage: 0.05‰. 6. Agarose gel DNA purification kit. 7. Restriction enzyme: NheI, BamHI, KpnI, XbaI, AatII, and SbfI. 8. Plasmid: pcDNA3.1(+) and pSEAP2-control. 9. Seamless cloning and assembly kit. 10. T4 DNA ligase. 11. Escherichia coli (E. coli). 12. Lysogeny broth (LB) liquid medium: 5 g/L NaCl, 5 g/L yeast, 10 g/L tryptone. The LB medium was solidified by the addition of 1.5% (w/v) agar. 13. Ampicillin (Amp): 100 mg/mL Ampicillin in ddH2O, filtersterilized (0.2μm). Stored at 20  C for up to 6 months. The stock was diluted into the corresponding culture medium at a final concentration of 100μg/mL. 14. Plasmid DNA extraction kit. 2.3 Cell Culture and Transfection

1. Cell culture laboratory.

2.3.1 Instruments

3. Biosafety cabinet.

2. Cell culture incubator. 4. Microscope. 5. Centrifuge. 6. Cell culture dish (diameter: 10 cm). 7. 24-well cell culture plate 8. Sterile syringe (20 mL). 9. Sterile conical centrifuge tube (15 and 50 mL). 10. Filter (0.22μm).

2.3.2 Reagents

1. Human embryonic kidney cells (HEK-293). 2. Cell culture medium: Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% (v/v) fetal bovine serum (FBS) and 1% (v/v) penicillin–streptomycin solution. 3. 1μg/μL polyethyleneimine (PEI, polyethyleneimine, molecular weight 40,000). Dissolve 100 mg of PEI in 100 mL of ddH2O, and adjusted to pH 7. Filter-sterilize (0.22μm) and store at 20  C.

2.4

FRL Illumination

Custom-designed 4  6 far-red LED array (730 nm, the size matches 24-well cell culture plate).

Smartphone-Controlled Optogenetic Switches

2.5 SEAP Reporter Assay

1. Constant temperature incubator (65  C).

2.5.1 Instruments

3. 96-well assay plate (transparent).

131

2. Multimode microplate reader. 4. Multipass pipette. 5. Reagent reservoirs.

2.5.2 Reagents

1. 2  SEAP buffer: 20 mM of homoarginine, 1 mM MgCl2, 21% (v/v) diethanolamine. Adjusted to pH of 9.8 and stored at 4  C, protected from light. 2. Substrate solution: 120 mM of p-nitrophenylphosphate. Stored at 20  C, protected from light. 3. PBS buffer: 8 g/L NaCl, 0.2 g/L KCl, 0.24 g/L KH2PO4, and 1.44 g/L Na2HPO4. Adjusted the pH of 7.4.

3

Methods

3.1 Construction the Smartphone Controlled FRL-Responsive Module (SmartController 1.0)

The smartphone ECNU-TeleMed app can remotely control the SmartControl-Box via the global GSM network to automatically trigger the 4  6 far-red LED array (Fig. 4).

3.1.1 Development of the Smartphone ECNU-TeleMed app

The design and development of the ECNU-TeleMed app was conducted using the Android software development kit, which contained the different preset algorithms, enabling the smartphone to remotely send user-defined parameters, including far-red LED brightness and illumination time (Fig. 5, see Note 1).

3.1.2 Assembly of the far-red LED Array Module

1. Connect the 4  6 far-red LEDs to the PCB (Fig. 6a, see Note 2). 2. Install a master control switch to the power supply terminals of the 24 far-red LEDs to achieve manual control of the light board circuit switch (Fig. 6b). 3. Arrange the 24 LEDs at equal intervals in an array of 4 rows and 6 columns, with 2 cm per row spacing and column spacing (see Note 3), in order to fit the dimensions of a 24-well cell culture plate (each LED was centered above a single well).

3.1.3 Assembly of the SmartController 1.0

1. Connect the power supply input interface of the intelligent remote controller to the AC/DC converter (see Note 4). 2. Connect the negative pole of the power supply input interface of the 6 switching power supplies to the 6 channels output relay switch of the intelligent remote controller.

132

Yuanhuan Yu et al.

Fig. 4 The smartphone-regulated electronic control system. (a) The experimental setting of the smartphonecontrolled SmartControl-Box-driven regulation of the custom-designed 4  6 far-red LED array. The 6 display panels represent the different preset SmartControl-Box-algorithms that can be controlled by the ECNUTeleMed app. (b) The detailed electric circuit diagram for the SmartController 1.0 system. (1)–(2) The AC/DC converter enables the power supply from different types of line currents, and is capable of supporting (3) the wireless signal processor, (4) the embedded MPU chip, (5) the relay drivers, (6) the relay units, and (7) the LED arrays. (These figures were adapted from Shao et al. [4])

3. Connect the output voltage and current adjustment switch of the 6 switching power supplies to the 6 channels output driver of the intelligent remote controller.

Smartphone-Controlled Optogenetic Switches

133

Fig. 5 Screenshots of the user interface for the ECNU-TeleMed app. (The picture was adapted from Shao et al. [4])

4. Connect the power supply input interface of the switching power supply to the AC/DC power converter. 5. Connect the power input of the far-red LED circuit board to the output of the switching power supply. 6. Connect the AC/DC power converter to AC 220 V. 7. Install the ECNU-TeleMed app that matches the SmartControl-Box in the smartphone to control the output voltage of the 6 independent switching power supplies by operating the smartphone app software through 3G/4G/WiFi network (see Note 5). 3.2 Construction of the BphS-BldD Based FRL-Triggered Optogenetic Switch

3.3 Cell Culture and Transfection

The FRL-triggered genetic switch contains three critical plasmids (pWS46, pGY32 and pXY34; Fig. 7). The DNA encoding BphS [19], YhjH [20], p65-VP64-BldD [21], promoter PFRL [21], and SEAP [22] (human placental secreted alkaline phosphatase) reporter were chemically synthesized, according to previous studies (Table 1). All constructs are confirmed using Sanger sequencing (see Note 6). 1. Grow human embryonic kidney cells (HEK-293) in cell culture medium at 37  C in a humidified atmosphere containing 5% CO2 (see Note 7).

a

J2

J5

Resistances (0805 package)

V6

V5 D5

A

D6

VCC

VCC

V4 D4

V6 DZ5

VCC

VCC

V5 DZ4

DZ3

DZ6

VCC

V4

V3 DZ2

VCC

V2

V1

2 AGND

V6

2 AGND

DZ1

VCC

Far-red lights

D3

J3

1

V4 1 J4

V3 D2

VCC

V1

V2

V3

2 AGND

2 AGND

V2

J1

8

D1

VCC

1 2 3 4

GND_ZT

GND_ZT

2 AGND

AGND S

1

V1 JR

1

1

A

GND

7

Parallel Interfaces of switching power supply 2 AGND

Brightness adjustment

6

VCC

5

V5

Self-locking switch

4

VCC

3

VCC

2

1

1

J6

Slide rhostat for fine adjustment

Device selection description:

GND

GND

GND

3 D16

3 D15

3 D14

3 D25

3 D26

GND 1

GND 1

GND 1 3 D24

3 D23

3 D36

3 D35

GND 1

GND 1

GND 1

2

GND 1 3 D46

3 D45

2

2

2

5.Independent regulating resistance (0805 package)

RP46

RP45

RP44

RP43 2

GND 1

GND 1

GND 1 RP42

2

3 D44

R46

C

GND 1

GND 1

GND 1

GND

GND

GND

D46

GND

2

D45

3 D42

D46 R45

3.Parallel Interfaces of switching power supply Rectifier diode 1N5408 Forward current 3A / Reverse Voltage1000V Hole size: 1.2mm 4.Far-red lights: Clearance 2.54mm;

RP36

RP35 2

2

2

RP41

GND

2

D44

RP26 2

RP34

RP33

GND 1

GND 1 R44

3 D41

R43

D45

D44

D43 R42

D46

GND

2

2

D43

2

2 D42

RP32 2

2

D41

2.Brightness adjustment: Select the appropriate sliding rheostat;

GND

GND

GND

GND

GND D42

D41 R41

GND 1

GND 1 RP31

GND

2 1 VCC D45

RP25 2

3 D34

R36

3 D33

R35

3 D32

D36

D35 R34

D36

D36

2 D35 D44

RP16 2

RP24 2

GND 1

D43

1.Self-locking switch: Clearance 2.54mm;

GND

GND

GND 1 R33

2

2

3 D31

R32

D34

D33

D32

D31 R31

GND 1

GND 1 3 D22

GND 1 3 D21

D26

D25

D24 GND

GND

GND

GND

2 1 VCC D35

1 VCC

2

2 D33

D34 1 VCC

2

2 D32

D31 1 VCC

1 VCC

1 VCC D42

RP15 2

RP23

RP22

RP21

2

D41

3 D13

3 D12

3 D11

D16

D14

D15

D13

D12

D11 D26

D26

2 D25 D34

RP14 2

3 D43

D33

1 VCC

2

2 D24

D23 D32

1 VCC

1 VCC

2

2 D22

D21 1 VCC

1 VCC D31

RP13 2

GND 1

D25

R26

R25

R24

R23

R22

R21

2

C

RP12 2

2

GND 1

D24

D22

D16 1 VCC

D23

R16

RP11

D21

D15 1 VCC

D22

R15

2

D14

2

D13 1 VCC

1 VCC

2

2

D12 1 VCC

2

2

D11 1 VCC

D21

R14

D16

GND

D15

R13

D23

D14

1 VCC

1 VCC

1 VCC D13

R12

GND

D12

R11

GND

D11

1 VCC

1 VCC

B

1 VCC

B

Schematic 24 Far-red lights

D

D

Title Size

Number

Revision

A3 Date: File: 1

2

3

4

5

6

2016/4/15 D:\AD14share\baiyu\24led.SchDoc 7

Sheet of Drawn By: 8

b

Fig. 6 The schematic for the circuit design of the 4  6 far-red LED array. (a) The circuit principle diagram of the 4  6 far-red LED array. (b) The printed circuit board for a 4  6 far-red LED array. (The figures were adapted from Shao et al. [4])

Smartphone-Controlled Optogenetic Switches

135

Fig. 7 The schematic depicting the configuration of the vectors for the FRL-triggered gene circuit. BphS-P2A-YhjH and p65-VP64-BldD were expressed under the control of a constitutive promoter PhCMV (human cytomegalovirus immediate early promoter). The synthetic light responsive promoter PFRL consisted of three fragments: pA (SV40 polyA), three copies of BldD binding site whiG, and the minimal promoter PhCMVmin (minimal version of PhCMV) Table 1 Plasmids encoding the FRL-triggered genetic switch used in this chapter [4] Plasmid Description and cloning strategy

References

pWS46

Constitutive mammalian PhCMV-driven BphS and YhjH expression vector (PhCMV-BphS-P2A-YhjH-pA) BphS and YhjH were chemically synthesized. BphS and YhjH were cloned into the corresponding sites (NheI/BamHI) of pcDNA3.1(+) using the Seamless Cloning and Assembly Kit

[4]

pGY32

Constitutive mammalian PhCMV-driven FRL-dependent transactivator expression [4] vector (PhCMV-p65-VP64-BldD-pA). VP64-BldD and p65 were chemically synthesized, and both fragments were cloned into the corresponding sites (KpnI/XbaI) of pcDNA3.1(+) using the Seamless Cloning and Assembly Kit

pXY34

FRTA-specific FRL-inducible SEAP expression vector [PFRL-SEAP-pA; PFRL, pA-(whiG)3-PhCMVmin]

[4]

BldD Streptomyces coelicolor transcription factor regulating aerial hyphae formation, BphS engineered bacterial diguanylate cyclase, FRTA mammalian far-red light-dependent transactivator, P2A picornavirus-derived self-cleaving peptide engineered for bicistronic gene expression in mammalian cells, p65 65 kDa transactivator subunit of NF-κB, pA polyadenylation signal, PFRL FRL-v2-specific chimeric promoter, PhCMV human cytomegalovirus immediate early promoter, PhCMVmin minimal version of PhCMV, SEAP human placental secreted alkaline phosphatase, VP64 tetrameric core of herpes simplex virus-derived transactivation domain, whiG BldD-specific binding sequence, YhjH bacterial c-di-GMP phosphodiesterase

2. Seed 6  104 HEK-293 cells per well in a 24-well cell culture plate at 18 h before transfection (see Note 8). 3. Transfect cells in each well (24-well plate) with a total of 300 ng of plasmid mixture (pWS46, 100 ng; pGY32, 100 ng; pXY34, 100 ng) diluted in 50μL of FBS-free and antibiotic-free DMEM medium. 4. Add 0.9μL of PEI (1μg/μL, PEI and DNA at a ratio of 3:1) and mix adequately (see Note 9).

136

Yuanhuan Yu et al.

Fig. 8 The image of the custom-designed far-red LED array used for the FRL-controlled transgene expression in mammalian cells grown in monolayer cultures. (The picture was adapted from Shao et al. [4])

5. Incubate the 50μL of DNA-PEI mixture solution at room temperature for 15 min to allow the complex formation between the positively charged PEI (amine groups) and negatively charged pDNA (phosphate groups), and the addition dropwise to the cells. 6. Change the medium with fresh cell culture medium [DMEM containing 10% (v/v) FBS and 1% (v/v) antibiotic] at 6 h after transfection (see Note 10). 3.4

FRL Illumination

1. Place the culture plate below a custom-designed 4  6 far-red LED array at 18 h after transfection (Fig. 8, see Note 11). 2. Remotely regulate the activity of the custom-made far-red LED array, including LED brightness and illumination time, through the SmartControl-Box driven by ECNUTeleMed app. 3. Expose cells to FRL (0–5 mW/cm2) for 4 h once a day for 2 days (see Note 12).

3.5 SEAP Reporter Assay

The production of human placental SEAP in the cell culture medium was quantified using the p-nitrophenylphosphate-based light absorbance time course assay (see Note 13). 1. The cell culture supernatant (200μL) was collected and placed in 96-well assay plates (transparent) at 48 h after the first illumination. 2. Heat-inactivated cell culture supernatant at 65  C for 30 min. 3. The addition of 100μL of 2  SEAP buffer and 20μL of substrate solution to the 80 μL heat-inactivated cell culture supernatant (see Note 14).

Smartphone-Controlled Optogenetic Switches

137

4. Measurement of the light absorbance at 405 nm at 37  C for 30 min using a multimode microplate reader (see Note 15). 5. Calculate the SEAP production from the slope of the timedependent increase in light absorbance.

4

Notes 1. The wavelength of the LED light can be custom-required (730 nm in this case). 2. Each group of 6 LEDs was connected in series with each current-limiting resistor. 3. The 24 far-red LEDs was centered above a single well of the 24-well cell culture plate to ensure the uniformity of the light distribution. 4. The fuse (parameter: 3A) was connected to the negative connecting line of the AC/DC converter in series, in order to enable the intelligent multipath remote controller to work within the safe current. 5. The illumination intensity was set by adjusting the output voltage of the six independent switching power supplies through operating the ECNU-TeleMed app software, and the duration of the far-red light by powering the six independent switching power supplies. Note that over illumination intensity and time can lead to the decrease in the level of induced SEAP production due to the cell damage caused by thermal stress. 6. The sequences of all constructs are confirmed using Sanger sequencing. 7. The DMEM medium was stored at 4  C. Prior to use, the media was preheated at 37  C for 30 min. Cells were regularly tested for the absence of Mycoplasma and bacterial contamination. Mycoplasma and bacterial contamination can influence the experimental results. 8. The cells should be in individual layers and uniformly dispersed in 24-well cell culture plates. 9. When transfecting in 24-well plates, the transfection mixture was spread in drops over the whole surface of the well. After pipetting, the plate was gently shaken in a cross pattern. The investigators recommend a 1:3 DNA-to-PEI ratio to achieve high transfection efficiency. 10. It is necessary to replace the transfection medium with fresh DMEM medium at 6 h after cell transfection. Long-term incubation with the transfection mixture would cause cell death.

138

Yuanhuan Yu et al.

11. When illuminating cells, each LED of the custom-designed 4  6 far-red LED array was aligned over each well of the cell culture plate. The dark-adapted samples in parallel were covered with an aluminum foil, in order to protect these from unintended illumination. 12. In the present system, cells were illuminated with 730 nm of FRL light at 1 mW/cm2 for 4 h per day for 2 days. The appropriate conditions for photoactivation was determined in the system. 13. The investigators prefer to use the SEAP reporter. However, other reporters, including fluorescent proteins or firefly luciferase, can also be used. The SEAP reporter can be replaced by any other outputs. 14. The mixture of 2 SEAP buffer and substrate solution should be quickly added to the cell culture supernatant samples, and immediately detected. 15. If the SEAP production is too high, the samples should be diluted with PBS.

Acknowledgments We are very grateful to Dr. Meiyan Wang for revising the manuscript. This work was financially supported by the grants from the National Key R&D Program of China, Synthetic Biology Research (no. 2019YFA0904500), the National Natural Science Foundation of China (NSFC: no. 31971346, no. 31861143016), the Science and Technology Commission of Shanghai Municipality (no. 18JC1411000) to H.Y. Materials availability: All genetic components related to this chapter are available with a material transfer agreement and can be requested from H.Y. (hfye@bio. ecnu.edu.cn). References 1. Bhavnani SP, Narula J, Sengupta PP (2016) Mobile technology and the digitization of healthcare. Eur Heart J 37(18):1428–1438 2. Preechaburana P, Suska A, Filippini D (2014) Biosensing with cell phones. Trends Biotechnol 32(7):351–355 3. Vashist SK, Luppa PB, Yeo LY et al (2015) Emerging technologies for next-generation point-of-care testing. Trends Biotechnol 33 (11):692–705 4. Shao J, Xue S, Yu G et al (2017) Smartphonecontrolled optogenetically engineered cells enable semiautomatic glucose homeostasis in diabetic mice. Sci Transl Med 9(387):eaal2298

5. Boyden ES, Zhang F, Bamberg E et al (2005) Millisecond-timescale, genetically targeted optical control of neural activity. Nat Neurosci 8(9):1263–1268 6. Bi A, Cui J, Ma Y et al (2006) Ectopic expression of a microbial-type rhodopsin restores visual responses in mice with photoreceptor degeneration. Neuron 50(1):23–33 7. Wang X, Chen X, Yang Y (2012) Spatiotemporal control of gene expression by a lightswitchable transgene system. Nat Methods 9 (3):266–269 8. Ye H, Daoud-El Baba M, Peng RW et al (2011) A synthetic optogenetic transcription device

Smartphone-Controlled Optogenetic Switches enhances blood-glucose homeostasis in mice. Science 332(6037):1565–1568 9. Kaberniuk AA, Shemetov AA, Verkhusha VV (2016) A bacterial phytochrome-based optogenetic system controllable with near-infrared light. Nat Methods 13(7):591–597 10. Shao J, Wang M, Yu G et al (2018) Synthetic far-red light-mediated CRISPR-dCas9 device for inducing functional neuronal differentiation. Proc Natl Acad Sci U S A 115(29): E6722–E6730 11. Nihongaki Y, Furuhata Y, Otabe T et al (2017) CRISPR-Cas9-based photoactivatable transcription systems to induce neuronal differentiation. Nat Methods 14(10):963–966 12. Nihongaki Y, Kawano F, Nakajima T et al (2015) Photoactivatable CRISPR-Cas9 for optogenetic genome editing. Nat Biotechnol 33(7):755–760 13. Yu Y, Wu X, Guan N et al (2020) Engineering a far-red light–activated split-Cas9 system for remote-controlled genome editing of internal organs and tumors. Sci Adv 6:eabb1777 14. Weitzman M, Hahn KM (2014) Optogenetic approaches to cell migration and beyond. Curr Opin Cell Biol 30:112–120 15. Favory JJ, Stec A, Gruber H et al (2009) Interaction of COP1 and UVR8 regulates UV-Binduced photomorphogenesis and stress acclimation in Arabidopsis. EMBO J 28 (5):591–601

139

16. Pan Y, Yang J, Luan X et al (2019) Nearinfrared upconversion-activated CRISPR-Cas9 system: a remote-controlled gene editing platform. Sci Adv 5(4):eaav7199 17. Clement M, Daniel G, Trelles M (2005) Optimising the design of a broad-band light source for the treatment of skin. J Cosmet Laser Ther 7(3–4):177–189 18. Ash C, Dubec M, Donne K et al (2017) Effect of wavelength and beam width on penetration in light-tissue interaction using computational methods. Lasers Med Sci 32(8):1909–1918 19. Ryu MH, Gomelsky M (2014) Near-infrared light responsive synthetic c-di-GMP module for optogenetic applications. ACS Synth Biol 3(11):802–810 20. Ryjenkov DA, Simm R, Ro¨mling U et al (2006) The PilZ domain is a receptor for the second messenger c-di-GMP: the PilZ domain protein YcgR controls motility in enterobacteria. J Biol Chem 281(41):30310–30314 21. Bush MJ, Tschowri N, Schlimpert S et al (2015) c-di-GMP signalling and the regulation of developmental transitions in streptomycetes. Nat Rev Microbiol 13(12):749–760 22. Schlatter S, Rimann M, Kelm J et al (2002) SAMY, a novel mammalian reporter gene derived from Bacillus stearothermophilus alpha-amylase. Gene 282(1-2):19–31

Chapter 9 Constructing a Smartphone-Controlled Semiautomatic Theranostic System for Glucose Homeostasis in Diabetic Mice Guiling Yu, Yuanhuan Yu, and Haifeng Ye Abstract With the development of mobile communication technology, smartphones have been used in point-of-care technologies (POCTs) as an important part of telemedicine. Using a multidisciplinary design principle coupling electrical engineering, software development, synthetic biology, and optogenetics, the investigators developed a smartphone-controlled semiautomatic theranostic system that regulates blood glucose homeostasis in diabetic mice in an ultraremote-control manner. The present chapter describes how the investigators tailor-designed the implant architecture “HydrogeLED,” which is capable of coharboring a designer-cell-carrying alginate hydrogel and wirelessly powered far-red light LEDs. Using diabetes mellitus as a model disease, the in vivo expression of insulin or human glucagon-like peptide 1 (shGLP-1) from HydrogeLED implants could be controlled not only by pre-set ECNU-TeleMed programs, but also by a custom-engineered Bluetooth-active glucometer in a semiautomatic and glycemia-dependent manner. As a result, blood glucose homeostasis was semiautomatically maintained in diabetic mice through the smartphone-controlled semiautomatic theranostic system. By combining digital signals with optogenetically engineered cells, the present study provides a new method for the integrated diagnosis and treatment of diseases. Key words Mammalian synthetic biology, Optogenetics, Synthetic designer cells, Telemedicine, Diabetes

1

Introduction Diabetes mellitus is a chronic metabolic disease that results from hyperglycemia [1]. At present, there are more than 463 million diabetic patients worldwide, and this number is predicted to increase to 700 million by 2045 [2]. The clinical manifestation of diabetes is the endocrine disorder of carbohydrate metabolism, which is primarily caused by inadequate insulin release (type 1 insulin-dependent diabetes mellitus) or insulin insensitivity coupled with insufficient compensatory insulin release (type 2 non–insulin-dependent diabetes mellitus) [3]. Type 1 and type 2 diabetes

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

141

142

Guiling Yu et al.

are both characterized by chronic hyperglycemia, which contributes to various diabetic complications, such as nephropathy, retinopathy, neuropathy, and cardiomyopathy [4–6]. Diabetic patients need to control their diet and inject insulin or human glucagon-like peptide 1 (shGLP-1) analogs every day to maintain blood glucose stability. However, repeated injections every day over a long period of time can cause great pain to patients. Furthermore, there is strong risk of hypoglycemia [7] or other side effects [8] due to the uncontrollable release of insulin and shGLP-1. With the advent of regenerative medicine, induced pluripotent stem cells (iPSCs) can be induced and differentiated into glucose-sensitive insulinproducing β-like cells, but the insufficient differentiation efficiency and purity greatly limit its clinical application in the treatment of diabetes [9–11]. The hybrid closed-loop automated insulin pump (MiniMed 670G) [12], which designed and developed by Medtronic (Northridge, CA), can regulate the injection volume of insulin in real time, according to the blood glucose level, thereby maintaining the blood glucose level within a certain range [13]. However, insulin needs to be filled to the container every 3 days, and the needle of the insulin pump can be rapidly blocked by fibrotic cells [14, 15]. Therefore, long-term sustainable glycemic control through therapeutic intervention, which enables the tight control of insulin delivery, remains as a key challenge in diabetes therapy. With the continuous innovation of mobile communication technology, the smartphone has rapidly developed, gradually transforming from a pure communication device into a multifunctional electronic device, and integrating high-definition cameras, electronic banks and global positioning systems [16]. Recently, the smartphone has also become an important part of telemedicine, and has been used in point-of-care technologies (POCTs) [17]. For example, diabetics can quickly and accurately measure their blood glucose at home with one drop of blood, and upload the information onto the smartphone, through which the information could be shared with doctors to consult further therapeutic or preventive interventions [18]. However, the present available telemedicine was developed for blood glucose monitoring, but not for therapeutic purposes. Therefore, there is an urgent need to develop a closedloop system that combines the blood glucose monitoring unit together with the insulin or shGLP-1 delivery unit to achieve the integration of diagnosis and treatment. In the present study, taking advantages of the accuracy of electronic and software engineering in regulating the input of digital information, the investigators coupled this with optogenetics, in order to design a smartphone-controlled semiautomatic theranostic system that allows for the wireless control of engineered cell activity with the ECNU-TeleMed app on the smartphone (Fig. 1) [19]. First, the investigators constructed an far-red light (FRL)-triggered transgene expression system based on the bacterial

A Smartphone-Controlled Semiautomatic Theranostic Systems

143

Fig. 1 The schematic of the smartphone-controlled semiautomatic theranostic system. The abstract diagram shows the smartphone-controlled engineered cells, which enable the semiautomatic point-of-care for controlling blood glucose homeostasis

light-activated cyclic diguanylate monophosphate (c-di-GMP) synthase BphS, and different synthetic mammalian transactivators (FRTAs) by assembling BldD, p65 (65-kDa transactivator subunit of nuclear factor kB [NF-kB]), VP64 (tetrameric core of herpes simplex virus–derived transactivation domain), and the multiple FRTA specific chimeric promoter (PFRL2.x) variants to regulate the therapeutic outputs production in diabetic mice. A custom-designed SmartControl-Box 3.0 was programmed to process wireless signals, enabling the smartphone to regulate insulin or shGLP-1 [20] production through the optogenetically engineered cells implanted in diabetic mice via a custom-designed “ECNUTeleMed” mobile application (app). In order to control the engineered cells with blood glucose data, the investigators customengineered a Bluetooth-active glucometer that transmits blood glucose data to the SmartController 3.0, automatically triggering the different illumination intensity of the receiver LEDs in diabetic mice, according to user-defined glycemic thresholds. In order to wirelessly control the production of therapeutic outputs through optogenetically engineered cells, the investigators designed and implanted HydrogeLED, which contained optogenetically engineered cells that carry alginate hydrogel, a wirelessly powered receiver coil, a resonance capacitor, and two far-red LEDs (lightemitting diodes). The custom-designed electromagnetic emission circuit (EEC), which generates electromagnetic sine wave signals at 180 kHz, wirelessly powers the HydrogeLED via electromagnetic induction. In diabetic mice, the optogenetically engineered cells in the HydrogeLED could be remotely controlled using the ECNUTeleMed app on the smartphone, or the blood glucose measured

144

Guiling Yu et al.

by the custom-engineered Bluetooth-active glucometer. The smartphone-controlled semiautomatic theranostic system can be applied to diverse metabolic diseases, providing a new strategy for cell-based therapy.

2

Materials

2.1 Plasmids (Table 1) 2.2 Buffers and Stock Solutions

1. Lysogeny broth (LB) medium: 10 g/L tryptone, 5.0 g/L yeast extract, and 5 g/L NaCl are dissolved in deionized water and autoclaved. 2. LB medium plates: Bacteriological agar is added to the liquid LB medium at 1.5% (w/v), and autoclaved afterward. 3. 10
PBS: 80 g/L NaCl, 2 g/L KCl, 35 g/L Na2HPO4·12H2O, and 2.7 g/L KH2PO4 is dissolved in deionized water. The pH is adjusted at 7.4, and autoclaved afterward. 4. Physiological saline solution: 9 g of NaCl is dissolved in 1 L of deionized water. Then, this is autoclaved and stored at room temperature. 5. Trypsin: 2.5 g of trypsin and 0.2 g of EDTA are dissolved in 1000 mL of 1
PBS. Then, the pH is adjusted at 7.4, and sterilized with a 0.22-μm filter. 6. 2
Buffer: 20 mM homoarginine, 1 mM MgCl, and 2% diethanol amine is dissolved in deionized water. The pH is adjusted at 9.8, and stored at 4  C protected from light. 7. p-Nitrophenol phosphoric acid (pNPP): 2.226 g of pNPP is dissolved in 50 mL of 2
Buffer, and stored at 20  C protected from light. 8. MOPS: 10 mM MOPS and 0.85% (w/v) NaCl are dissolved in deionized water. The pH is adjusted at 7.2. 9. Polymerization buffer: 10 mM MOPS and 100 mM CaCl2 are dissolved in deionized water. The pH is adjusted at 7.2. 10. 1.5% (w/v) sodium alginate: 1.5 g of sodium alginate is dissolved in 100 mL of MOPS. Then, this is sterilized with a 0.22μm filter, and stored at 4  C. 11. 0.05% (w/v) Poly-L-lysine solution: 0.05 g of poly-L-lysine is dissolved in 100 mL of MOPS. Then, this is sterilized with a 0.22-μm filter (see Note 1). 12. D-Glucose: 1 g of D-glucose is dissolved in 10 mL of physiological saline solution. Then, this is sterilized with a 0.22-μm filter.

A Smartphone-Controlled Semiautomatic Theranostic Systems

145

Table 1 Plasmids designed and used in this study Plasmid

Description and cloning strategy

References

pSBtet-GP

Sleeping beauty transposon stable expression vector for doxycyclineAddgene inducible firefly luciferase and constitutive expression of GFP, rtTA, and (#60495) puromycin resistance gene (PTRE3G-luciferase-pA-PRPBSA-GFP-2A-rtTA2A-puromycin-pA)

pCMVT7SB100

PhCMV-driven SB100X transposase expression vector (PhCMV-SB100X-pA) Addgene (#34879)

pWS251

FRTA-specific far-red light (FRL)-inducible SEAP and insulin stable expression vector, containing constitutive ZeoR and EGFP expression unit (ITR-PFRL2.13a-SEAP-P2A-mINS-pA::PmPGK-ZeoR-P2A-EGFPpA-ITR)

[19]

pWS252

FRTA-specific far-red light (FRL)-inducible shGLP-1 and SEAP stable expression vector, containing constitutive ZeoR and EGFP expression unit (ITR-PFRL2.13a-shGLP-1-P2A-SEAP-pA: PmPGK-ZeoR-P2AEGFP-pA-ITR)

[19]

pYH88

Constitutive mammalian stable expression vector for BphS, YhjH, PuroR and mCherry (ITR-PhCMV-BphS-P2A-YhjH-P2A-P65-VP64-BldDP2A-mCherry-pA: PmPGK-PuroR-pA-ITR)

[19]

BldD Streptomyces coelicolor transcription factor regulating aerial hyphae formation, BphS engineered bacterial diguanylate cyclase, EGFP enhanced green fluorescent protein, FRL far-red light, FRTA mammalian far-red light-dependent transactivator, mCherry mushroom coral red fluorescence protein, mINS modified rodent insulin variant for optimal expression in HEK-293 cells, P2A picornavirus-derived self-cleaving peptide engineered for bicistronic gene expression in mammalian cells, p65 65 kDa transactivator subunit of NF-kB, pA polyadenylation signal, PCR polymerase chain reaction, PFRL2.13a (pA-(whiG)3-PhCMVmin), BldD-based FRTA-specific synthetic mammalian far-red light-inducible promoter variants, PhCMV human cytomegalovirus immediate early promoter, PhCMVmin minimal version of PhCMV, PmPGK mouse phosphoglycerate kinase gene promoter, PRPBSA a synthetic RPBSA promoter, which is made up of a fragment of the RPL13a promoter fused to a region of the RPL41 gene, to drive the selection/reporter parts of each construct, PuroR gene product that confers puromycin resistance to mammalian cells, rtTA reverse tetracycline/ doxycycline dependent mammalian transactivator, PTRE3G rtTA-specific doxycycline-inducible mammalian promoter (Clontech, CA), SB100X the Sleeping Beauty transposase, SEAP human placental secreted alkaline phosphatase, shGLP-1 short variant of human glucagon-like peptide 1, VP64 tetrameric core of Herpes simplex virus-derived transactivation domain, whiG BldD-specific binding sequence, YhjH bacterial c-di-GMP phosphodiesterase, ZeoR gene product that confers zeocin resistance to mammalian cells

13. Citric acid solution: 1.05 g of citric acid is dissolved in 50 mL of deionized water. Then, the pH is adjusted at 4.5, and sterilized with a 0.22-μm filter. 14. Sodium citrate solution: 1.47 g of sodium citrate is dissolved in 50 mL of deionized water. Then, the pH is adjusted at 4.5 and sterilized with a 0.22-μm filter. 15. Cell culture medium: Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin solution. Then, this is stored at 4  C, and warmed up to 37  C before use. 16. Polyethyleneimine (PEI, 1μg/μL). This is dissolve with ddH2O, the pH is adjusted to 7, and this is sterilized with a 0.22-μm filter. This is stored at 20  C.

146

Guiling Yu et al.

17. Plasmid extraction kit (TIANGEN). 18. Mouse Insulin ELISA Kit (Mercodia AB). 19. Secreted alkaline phosphatase (SEAP) assay kit (Roche Diagnostics). 20. High-Sensitivity GLP-1 Active ELISA kit (Merck Millipore). 21. Puromycin (1μg/mL) and zeocin (100μg/mL). Stored at 20  C. 22. Streptozotocin (Sigma). 2.3

Lab Equipment

1. Microplate reader (BioTek). 2. Conical centrifuge tubes, 15 and 50 mL. 3. Far-red LEDs. 4. Tissue culture plates (24-well plates and 12-well plates). 5. Table-top centrifuge. 6. Mouse ear puncher. 7. Cell culture incubator (37  C, 5% CO2). 8. Hemocytometer. 9. Eppendorf tubes, 1.5 mL. 10. Sterile syringe filter with a 0.22μm polyethersulfone (PES) membrane. 11. Heat lamp for use during small animal surgery (75 W infrared bulb). 12. Electric clipper for small animals. 13. Surgical instruments. 14. Sterile syringes. 15. Skin staples. 16. Shaker incubator for bacterial cell culture. 17. Inverted light microscope. 18. Acrylic barrel. 19. Contour Glucometer (Exactive Easy III, MicroTech Medical Co., Ltd.). 20. Electronic scales. 21. 5D-8B commercial glucometer (5D-8B, Yi Cheng Co., Ltd.). 22. Intelligent remote controller (Smart Home Studio) (see Note 2). 23. Switching power supply (Yunlifang Studio) (see Note 3). 24. ECNU-TeleMed app (Smart Home Studio) (Fig. 2, see Note 4).

A Smartphone-Controlled Semiautomatic Theranostic Systems

147

Fig. 2 The conversion between the different glycemic threshold (GT) procedures and FRL illumination strengths. The glucose data displayed on the SmartController 3.0 LEDs are synchronized with the smartphone’s ECNU-TeleMed app. (a) GT1: 16.8 mM. FRL-activity was set to 5 mW/cm2 when scoring one drop of blood with the custom-designed glucometer of 21.7 mM. (The picture is adapted from Shao et al. [19])

148

Guiling Yu et al.

2.4 Cells and Animals

1. Cell lines: human embryonic kidney cells HEK293 (ATCC: CRL-11268). 2. Mouse strain: 12-week-old male wild-type C57BL/6J mice and 12-week-old male db/db mice (BKS.Cg-Dock7m +/+ Leprdb/J, derived from C57BL/6J mice, Charles River Laboratory) (East China Normal University [ECNU] Laboratory Animal Center) (see Note 5).

3

Methods

3.1 Construction of the SmartController 3.0

The SmartController 3.0 includes the ECNU-TeleMed app, a SmartControl-Box 3.0, custom-designed glucometer, a customdesigned electromagnetic emission circuit (EEC) and coiledLEDs (Figs. 3 and 4). The Bluetooth receiver and liquid crystal display can simultaneously receive different types of wireless signals. The microcontroller unit can be programmed to interpret the data transmitted from a custom-designed glucometer to automatically trigger the illumination of different intensities via remotely controlled coiled-LEDs, according to user-specified glycemic thresholds. All blood glucose data are synchronized using the ECNUTeleMed app on the smartphone, in order to enable real-time surveillance and optional intervention by humans.

3.1.1 Custom-Designed Glucometer

The custom-designed glucometer includes a commercial glucometer (5D-8B, Yi Cheng Co., Ltd.), a HC-05 Bluetooth transmitter, and a 3.7 V rechargeable lithium battery (Fig. 5). 1. Connect the blood glucose data transmission interface of the 5D-8B commercial glucometer with the HC-05 Bluetooth transmitter. 2. Connect the 3.7 V rechargeable lithium battery to the power supply input interface of the HC-05 Bluetooth transmitter. 3. Connect the 3.7 V rechargeable lithium battery to the power supply input interface of the 5D-8B commercial glucometer.

3.1.2 Blood Glucose Microprocessor

The blood glucose microprocessor includes the Bluetooth receiver, the microcontroller unit (MSP430) (see Note 6) and the liquid crystal display (see Note 7). 1. Connect the Bluetooth receiver module to the microcontroller unit (MSP430). 2. Connect the liquid crystal display to the microcontroller unit (MSP430).

A Smartphone-Controlled Semiautomatic Theranostic Systems

149

Fig. 3 Electric circuit diagram for SmartController 3.0. (The figure is adapted from Shao et al. [19])

Fig. 4 A photograph of the SmartController 3.0. (The figure is adapted from Shao et al. [19])

150

Guiling Yu et al.

Fig. 5 A photograph of the custom-designed glucometer. (The figure is adapted from Shao et al. [19]) 3.1.3 SmartControl-Box 3.0

The SmartControl-Box 3.0 includes the intelligent remote controller, the blood glucose microprocessor, and the switching power supply. 1. Connect the power supply input interface of the intelligent remote controller to the ac/dc converter (see Note 8). 2. Connect the negative pole of the power supply input interface of the six switching power supplies to the six channels output relay switch of the intelligent remote controller. 3. Connect the output voltage and current adjustment switch of the six switching power supplies to the six channels output driver of the intelligent remote controller. 4. Connect the microcontroller unit (MSP430) of the blood glucose microprocessor with the intelligent remote controller, through which the blood glucose data are transmitted to the ECNU-TeleMed app. 5. Connect the power supply input interface of the switching power supply to the ac/dc converter. 6. Connect the ac/dc converter to ac 220 V. 7. Match the Bluetooth transmitter module of the customdesigned glucometer with the Bluetooth receiver module of SmartControl-Box 3.0.

3.1.4 Custom-Designed Electromagnetic Emission Circuit (EEC)

An autonomous electromagnetic emission circuit (EEC) contains an electromagnetic field regulator, a homemade transmitting circular coil, and an acrylic barrel (Fig. 6).

A Smartphone-Controlled Semiautomatic Theranostic Systems

151

Fig. 6 Photographs of the custom-designed EEC. (a) The transmitting circular coil of 20 cm in diameter. (b) The electromagnetic field regulator. (c) The transmitter coil–containing field generator in an acrylic barrel. (The figure is adapted from Shao et al. [19])

1. Using a winding machine, bend the pure copper wire into the homemade transmitting circular coil with an outer diameter of 20 cm. 2. Connect the output terminal of the electromagnetic field regulator to the homemade transmitting circular coil (see Note 9). 3. Fix two transmitting circular coils on the inner wall of the acrylic barrel with an inner diameter of 20 cm (see Note 10). 4. Connect the input terminal of the custom-designed electromagnetic emission circuit to the switching power supply of the SmartControl-Box 3.0. 3.1.5 Coiled-LEDs

The Coiled-LEDs include a receiver coil, a resonance capacitor, and two far-red LEDs (Fig. 7) (see Note 11). 1. Using a winding machine, bend the pure copper wire into the receiver coil (inductance of 50 uH, wire diameter of 0.5 mm) with an outer diameter of 15 mm. 2. Connect two far-red LEDs in series, and connect these in parallel with the receiver coil and resonance capacitor (see Note 12).

3.2 Construction of the Stable Cell Line HEKFRL-SEAP-P2A-mINS and HEKFRL-shGLP-1-P2A-SEAP

1. Grow human embryonic kidney cells (HEK-293) in Cell culture medium at 37  C in a humidified atmosphere containing 5% CO2. 2. Then, seed 5
104 HEK293 cells per well in a 24-well cell culture plate at 18 h before transfection. 3. Transfect in each well with a total of 220 ng of plasmid mixture (pYH88, 100 ng; pWS251, 100 ng; pCMV-T7-SB100, 20 ng) diluted in 50μL of FBS-free and antibiotic-free DMEM medium (see Note 13).

152

Guiling Yu et al.

Fig. 7 Characteristics of the Coiled-LEDs. (a) The schematic of the Coiled-LEDs circuit. (b) A photograph showing the dimensions of the Coiled-LEDs (15 mm in diameter). (c) A photograph showing the full functionality of the Coiled-LEDs when placed in liquid solutions. (The figure is adapted from Shao et al. [19])

4. Transfect in each well with a total of 220 ng of plasmid mixture (pYH88, 100 ng; pWS252, 100 ng; pCMV-T7-SB100, 20 ng) diluted in 50μL of FBS-free and antibiotic-free DMEM medium (see Note 14). 5. Then, add 0.66μL of PEI (polyethyleneimine, molecular weight 40,000, stock solution at 1 mg/mL in ddH2O, PEI and DNA at a ratio of 3:1) and adequately mix. 6. Incubate at room temperature for 15 min to allow for the complex formation between positively charged PEI (amine groups) and negatively charged pDNA (phosphate groups), and add dropwise to the cells [21]. 7. Incubate the DNA–PEI mixture solution with cells at 37  C in a humidified atmosphere containing 5% CO2 for 6 h, the medium is removed, and cells are digested with trypsin and transferred to a 10-cm cell culture dish. 8. After 24 h, add 1μg/mL puromycin and 100μg/mL zeocin to the cell culture dish, and incubate at 37  C for 2 weeks (see Notes 15–17). 9. Pick the surviving cell clones to 24-well cell culture plates for further cultivation randomly. The cells are digested and distributed into two 24-well plates on average when the number of cells grew to approximately 12
104 per well (see Note 18). 10. Profile for FRL-stimulated SEAP expression performance through the SmartController system (1 mW/cm2; 730 nm) for 2 days at 4 h per day. The SEAP expression in the culture supernatant is scored at 72 h after the first illumination. 11. Select cell clones with almost negligible background and robust induction activity of transgene for the following studies.

A Smartphone-Controlled Semiautomatic Theranostic Systems

3.3 Construction of the Type 1 Diabetic Mouse Model

153

1. Group ten-week-old male mice (C57BL/6J, East China Normal University Laboratory Animal Centre) by body weight, at five per cage, and fast for 16 h. 2. Mix the prepared citric acid solution and sodium citrate solution at a ratio of 1:1, and place in ice (see Note 19). 3. Weigh the streptozotocin dosage (50 mg/kg), dissolved in 200μL of the citric acid and sodium citrate mixed solution, and intraperitoneally injected into mice for 5 days (see Note 20). 4. At 2 weeks after the injection, consider mice with fasting blood glucose over 16.6 mmol/L as diabetic, and use for further experiments.

3.4 HydrogeLED Implant 3.4.1 Preparation of the hydrogeLED Implant

1. Suspend the HEKFRL-SEAP-P2A-mINS cells in the 1.5% (w/v) sodium alginate buffer to a final concentration of 4
106 cells/mL. 2. Then, pipette 500μL of this suspension onto one well of a 24-well plate, and a receiver coil–containing LED is placed at the bottom. 3. Solidify for over 10 min by adding 500 mL of polymerization buffer. This is repeated for three times (see Note 21). 4. Incubate for 10 min in the 0.05% poly-L-lysine solution (Fig. 8).

3.4.2 HydrogeLED Implantation

1. Anesthetize mice by intraperitoneal injection of pentobarbital sodium salt (30 mg/kg). 2. Implant the HydrogeLED implants into the back of mice (see Note 22). 3. Suture the incised skin with skin staples. 4. Transfer the mouse to the SmartController 3.0-driven electromagnetic emission circuit (EEC) at 1 h after implantation.

Fig. 8 A photograph of the HydrogeLED in the 3.5-cm dish. (The figure is adapted from Shao et al. [19])

154

Guiling Yu et al.

3.5 Semiautomatic Control of Blood Glucose Homeostasis in Diabetic Mice

1. Match the SmartControl-Box TeleMed app.

3.0

with

the

ECNU-

2. Connect the Bluetooth of the Custom-designed glucometer to the Bluetooth of the smartphone (see Note 23). 3. Open the smartphone network, and turn on the glucose data acquisition switch of the ECNU-TeleMed app. 4. Measure the blood glucose from the caudal vein of the mouse with the custom-designed glucometer (see Note 24). 5. Display the present blood glucose level and determine whether the blood glucose level is within the normal threshold by the ECNU-TeleMed app. 6. According to the level of the blood glucose, the ECNUTeleMed app controls the illumination intensity and time of the far-red LED through the SmartControl-Box 3.0.

3.6 The Therapeutic Efficacy of the SmartphoneRegulated Semiautomatic Theranostic System in Diabetic Mice

1. Collect blood samples at 48 or 72 h after implantation for insulin detection. The serum is prepared via centrifugation (3000
g for 10 min) of clotted blood (37  C for 0.5 h, and subsequently 4  C for 2 h). 2. Quantify the insulin in the serum of mice using the mouse insulin ELISA Kit (Mercodia AB; Cat. no. 10-1247-01), according to manufacturer’s instructions.

3.6.1 Insulin ELISA 3.6.2 Glucagon-Like Peptide-1 (GLP-1) ELISA

3.6.3 Intraperitoneal Glucose Tolerance Test (IGTT) in Mice

The shGLP-1 in the serum of mice are profiled using a HighSensitivity GLP-1 Active ELISA kit (Merck Millipore; Cat. no. EGLP-35K), according to manufacturer’s instructions. 1. Fast for 16 h after 2 days of treatment. 2. Inject each mouse with aqueous 1 g/kg of intraperitoneally.

D-glucose

3. Monitor the glycemic profile of each mouse via the tail-vein blood samples at 0, 30, 60, 90 and 120 min after glucose administration using a Contour Glucometer. 4. Use the trapezoidal rule to determine the area under the curve (AUC) for IGTT. 3.6.4 Intraperitoneal Insulin Tolerance Test (IGTT) in Mice

1. Fast for 4 h after 2 days of treatment. 2. Each mouse is intraperitoneally injected with aqueous 0.1 U/ mL of insulin. 3. Monitor the glycemic profile of each mouse via the tail-vein blood samples at 0, 30, 60, 90 and 120 min after glucose administration using a Contour Glucometer.

A Smartphone-Controlled Semiautomatic Theranostic Systems

155

4. Calculate the approximation equation for the insulin resistance index with the formula: HOMA-IR ¼ [fasting glucose (mmol/ L)
fasting insulin (mU/L)]/22.5.

4

Notes 1. Weigh poly-L-lysine should be placed at low temperature, and used immediately after mixing with MOPS. 2. The intelligent remote controller includes a GSM wireless signal processor, an embedded microcontroller unit (MPU), 6 channels of the independent output relay driver groups, and an ac/dc converter. 3. The switching power supply includes a voltage and current display liquid crystal display screen, an output voltage and current adjustment switch, an input interface, and an output interface. 4. The custom-designed ECNU-TeleMed app can control the SmartController-box 3.0 activities. The design and development of the ECNU-TeleMed app connected to the SmartController-box 3.0 is performed by using the Android software development kit, which contains different preset algorithms to regulate the electronic devices. 5. For animal breeding and experiments, the national animal welfare regulations and guidelines must be followed, and ethical permission should be granted prior to commencement. 6. The microcontroller unit (MSP430) can be programmed to interpret the data transmitted from a custom-designed glucometer. The microcontroller unit (MSP430) is programmed with four glycemic thresholds (GTs) (GT1, 16.8 mM). The microcontroller unit judges the received blood glucose data and outputs the corresponding regulatory signals of different blood glucose values to adjust the switch state of the relay and the output voltage of the driver, thereby realizing the different illumination intensity and illumination times of the far-red LED. 7. The blood glucose data (blood glucose level and test time) are transmitted to the liquid crystal display. 8. The fuse (parameter of 3A) is connected with the negative connecting line of the ac/dc converter in series to protect the SmartControl-Box 3.0 with multi-output function, and allow it to work normally within the safe current. 9. The homemade transmitting circular coil is capable of generating electromagnetic sine wave signals at 180 kHz.

156

Guiling Yu et al.

10. The effective distance of the wireless electromagnetic induction is within 20 cm around the coil, and the two transmitting circular coils form a nearly uniform wireless electromagnetic environment. 11. The receiver coil receives the electromagnetic energy of the transmitting circular coil, and supplies power to the far-red LED through the rectifying and filtering of the capacitor. 12. The resonance capacitor is a chip tantalum capacitor with a capacitance of 15 nF. 13. The HEKFRL-SEAP-P2A-mINS cell line induces the expression of SEAP and mouse insulin by far-red light. 14. The HEKFRL-shGLP-1-P2A-SEAP cell line induces the expression of shGLP-1 and SEAP by far-red light. 15. The state of the cells should be observed before adding the antibiotics, which are added only when the cells are in good condition. 16. For different types of cells, puromycin and zeocin concentration gradient tests should be performed to determine the final concentration to be used. 17. Fresh medium (containing FBS, penicillin/streptomycin, puromycin, and zeocin) should be replaced every 3 days for 2 weeks. 18. Use a pipette to gently suspend the mix cells several times, making sure the cells are evenly distributed. 19. After mixing evenly, adjust the pH at 4.5, and the whole process should be operated on ice. 20. STZ is extremely sensitive to temperature and light. Weigh STZ at low temperature and away from light, and use this immediately after mixing with the citric acid and sodium citrate solution. 21. The hydrogel solidifies completely, and wraps the receiver coil completely. 22. Turn the far-red LED side down in the hydrogel to make the light more adequate. 23. The custom-designed glucometer can transmit the glucose signal to ECNU-TeleMed app and SmartControl-Box 3.0 via Bluetooth. 24. Send the blood glucose data to the SmartControl-Box 3.0 via Bluetooth. When the custom-designed glucometer detects the blood glucose, the ECNU-TeleMed app can get the blood glucose value immediately.

A Smartphone-Controlled Semiautomatic Theranostic Systems

157

Acknowledgments We are very grateful to Dr. Ningzi Guan for revising the manuscript. This work was financially supported by the grants from the National Key R&D Program of China, Synthetic Biology Research (no. 2019YFA0904500), the National Natural Science Foundation of China (NSFC: no. 31971346, no. 31861143016), the Science and Technology Commission of Shanghai Municipality (no. 18JC1411000) to H.Y. Materials availability: All genetic components related to this paper are available with a material transfer agreement and can be requested from H.Y. (hfye@bio. ecnu.edu.cn). References 1. Zhang J, Li Y, Li H et al (2018) GDF11 improves Angiogenic function of EPCs in diabetic limb ischemia. Diabetes 67 (10):2084–2095 2. Saeedi P et al (2019) Global and regional diabetes prevalence estimates for 2019 and projections for 2030 and 2045: Results from the International Diabetes Federation Diabetes Atlas, 9(th) edition. Diabetes research and clinical practice 157:107843 3. American Diabetes Association (2020) 2. Classification and diagnosis of diabetes: standards of medical Care in Diabetes—2020. Diabetes Care 43(Supplement 1):S14–S31 4. Yang X, Ongusaha PP, Miles PD et al (2008) Phosphoinositide signalling links O-GlcNAc transferase to insulin resistance. Nature 451 (7181):964–969 5. Langlet F, Haeusler RA, Linde´n D et al (2017) Selective inhibition of FOXO1 activator/ repressor balance modulates hepatic glucose handling. Cell 171(4):824–835.e818 6. Kernan WN, Viscoli CM, Furie KL et al (2016) Pioglitazone after ischemic stroke or transient ischemic attack. N Engl J Med 374 (14):1321–1331 7. Gubitosi-Klug RA, Braffett BH, White NH et al (2017) Risk of severe hypoglycemia in type 1 diabetes over 30 years of follow-up in the DCCT/EDIC study. Diabetes Care 40 (8):1010–1016 8. Cravalho CK, Meyers AG, Mabundo L et al (2019) 1326-P: metformin increases GLP-1 concentrations and improves glycemia in youth with Type 2 diabetes. Diabetes 68(Supplement 1):1326 9. Yamaguchi T, Sato H, Kato-Itoh M et al (2017) Interspecies organogenesis generates

autologous functional islets. Nature 542 (7640):191–196 10. Zhu Z, Li QV, Lee K et al (2016) Genome editing of lineage determinants in human pluripotent stem cells reveals mechanisms of pancreatic development and diabetes. Cell Stem Cell 18(6):755–768 11. Zhou Q, Melton DA (2018) Pancreas regeneration. Nature 557(7705):351–358 12. Smalley E (2016) Medtronic automated insulin delivery device gets FDA nod. Nat Biotechnol 34(12):1220 13. Knebel T, Neumiller JJ (2019) Medtronic MiniMed 670G hybrid closed-loop system. Clin Diabetes 37(1):94–95 14. Saunders A, Messer LH, Forlenza GP (2019) MiniMed 670G hybrid closed loop artificial pancreas system for the treatment of type 1 diabetes mellitus: overview of its safety and efficacy. Expert Rev Med Devices 16 (10):845–853 15. Trevitt S, Simpson S, Wood A (2016) Artificial pancreas device Systems for the Closed-Loop Control of type 1 diabetes: what systems are in development? J Diabetes Sci Technol 10 (3):714–723 16. Quesada-Gonza´lez D, Merkoc¸i A (2017) Mobile phone-based biosensing: an emerging “diagnostic and communication” technology. Biosens Bioelectron 92:549–562 17. Wang P, Kricka LJ (2018) Current and emerging trends in point-of-care technology and strategies for clinical validation and implementation. Clin Chem 64(10):1439–1452 18. Sharp L, Farrance I, Greaves RF (2016) The application of glucose point of care testing in three metropolitan hospitals. Pathology 48 (1):51–59

158

Guiling Yu et al.

19. Shao J, Xue S, Yu G et al (2017) Smartphonecontrolled optogenetically engineered cells enable semiautomatic glucose homeostasis in diabetic mice. Sci Transl Med 9(387):eaal2298 20. Ye H, Daoud-El Baba M, Peng RW et al (2011) A synthetic optogenetic transcription device enhances blood-glucose homeostasis in mice.

Science (New York, NY) 332 (6037):1565–1568 21. Shao J, Wang M, Yu G et al (2018) Synthetic far-red light-mediated CRISPR-dCas9 device for inducing functional neuronal differentiation. Proc Natl Acad Sci U S A 115(29): E6722–E6730

Chapter 10 Construction of Caffeine-Inducible Gene Switches in Mammalian Cells Daniel Bojar Abstract Controlling gene expression in mammalian cells constitutes one of the core principles of mammalian synthetic biology. Especially for cell-based therapies, inducers of gene expression which demonstrate the highest degree of safety and patient adherence are needed. In this chapter, I describe methods to implement caffeine-controlled gene expression systems into mammalian cells. Using an array of different implementation strategies, from reconstituting transcription factors to activating endogenous signaling pathways, allows for a wide range of sensitivity and capacity of the resulting caffeine-responsive gene switches. Key words Synthetic biology, Caffeine, Receptors, Transcription factors, Inducible expression

1

Introduction The control of gene expression is an integral part of most projects in synthetic biology, which aims to use and shape cell function according to engineering principles [1]. Easily accessible genetic engineering tools have enabled the construction of gene circuits which use this control of gene expression to influence cell function, for instance by producing and secreting a therapeutic protein in response to the gene expression inducer. While synthetic biology has expanded rapidly since its conception, mammalian synthetic biology is still in its maturation phase, despite its considerable advantages for an array of applications. From biocomputing [2, 3] over the production of biopharmaceuticals [4] and up to biomedical applications [5, 6], inducible gene expression is a mainstay in mammalian synthetic biology. This makes the choice of inducer molecule so crucial for the design of a project.

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

159

160

Daniel Bojar

1.1 Inducible Gene Expression Systems in Mammalian Synthetic Biology

Mammalian synthetic biology began by importing principles of gene expression regulation from bacterial synthetic biology. The bacterial transcription factor TetR (Tet repressor) was fused to the viral transactivating peptide VP16 to yield tTA (tetracyclinecontrolled transactivator protein), a transcription factor which can be turned off by the addition of the antibiotic doxycycline [7]. The control of transcription factor binding to their DNA binding site (their operator) is a general principle in the regulation of gene expression and by now has many more examples, such as control by the small molecules vanillic acid [8], benzoate [9], or phloretin [10]. Even environmental factors such as light have been used to control gene expression in this manner [11]. Later efforts have begun to make use of the natural signaling capacities of mammalian cells by designing receptors and/or capitalizing on signaling pathways which could be for instance activated by small molecules. Examples of the former include synNotch receptors [12], chimeric antigen receptors (CAR) [13], and generalized extracellular molecule sensors (GEMS) [14] which all modify transmembrane receptors with a customizable antibody fragment and activate gene expression via signaling pathways after inducer binding. Synthetic biologists have also been able to capitalize on existing signaling pathways by designing synthetic promoters [15] which make the expression of a transgene dependent on the activation of a pathway, for instance promoters regulated by the Ca2+-NFAT (nuclear factor of activated T-cells) pathway [16].

1.2 Using Caffeine in Mammalian Synthetic Biology

While much effort has been invested into finding orthogonal inducers of gene expression, there are considerable benefits in using an inducer which is already familiar to the human body. Continuous intake of caffeine in high doses has been monitored in countless studies over decades and no considerable side effects have been identified [17, 18]. Additionally, caffeine is inexpensive to synthesize [19] and only present in a small set of beverages, in which its presence is labelled. This makes it an ideal inducer for cell-based therapies, in which the control of gene expression is paramount and can be tailored to the consumption habits of a patient. This shaping of a therapy to lifestyle is likely to ameliorate the looming issue of patient nonadherence [20]. Capitalizing on a caffeine-binding nanobody which homodimerizes in the presence of caffeine [21], we recently constructed caffeine-inducible gene switches in mammalian cells via several routes [22]. This could then be used for a coffee-controlled cellbased treatment for type-2 diabetes in several mouse models. Next to traditional receptor engineering, we also demonstrated the potential of using this for nontraditional means of cellular engineering by controlling total cellular translation via caffeine with a PKR (protein kinase R)–nanobody fusion construct [23].

Construction of Caffeine-Inducible Gene Switches in Mammalian Cells

161

Caffeine Receptor

Signaling Pathway

Reporter gene

Needs to be activated by dimerization (EpoR, IL4R/IL13R)

Signal amplification determines sensitivity (e.g. STAT/MAPK/Ca2+) Number of transcription factor binding sites crucial for sensitivity

Fig. 1 Schematic of caffeine-responsive signaling pathways for inducible gene expression in mammalian cells. Any receptor which is activated by dimerization (e.g., EpoR or the interleukin 4 receptor–interleukin 13 receptor heterodimer) can be fused to the caffeine-dimerizable nanobody aCaffVHH. Depending on the used receptor and cell line, intracellular signaling mediators such as STAT3 do not necessarily need to be exogenously introduced. However, there is a broad range of possible signaling mediators (STAT6, MAPK, Ca2+/ NFAT etc.), some of which do need transfection of additional genes for full functionality. The choice of signaling pathway determines the kinetic parameters of the expression system and is a major factor in determining inducer sensitivity thanks to signal amplification. Lastly, the number of transcription factor binding sites controlling transgene expression also has a strong influence on inducer sensitivity

In this chapter, I describe in detail the construction and validation of caffeine-inducible gene switches in mammalian cells. This will comprise several different routes of arriving at caffeineinducible gene expression, with a focus on the EpoR-IL6ST (interleukin 6 signal transduction domain)/STAT3 (signal transducer and activator of transcription 3) receptor system used in our coffee-controlled diabetes treatments. However, in principle, any dimerization-dependent activation of signaling pathways or gene expression can be utilized in concert with the caffeine nanobody. The flexibility and robustness of the caffeine nanobody in protein engineering setups [24] makes it a promising candidate in the toolbox of mammalian synthetic biology, especially for biomedical applications (Fig. 1).

2 2.1

Materials Cell Culture

1. Dulbecco’s Modified Eagle’s Medium (DMEM) with 10% fetal bovine serum (FBS) and 1% penicillin–streptomycin. 2. 0.05% (v/v) Trypsin–EDTA solution). 3. Cell line (HEK-293T, human embryonic kidney cells).

162

Daniel Bojar

4. CO2 cell culture incubator. 5. Cell counting device (e.g., CASY® Cell Counter and Analyzer Model TT). 6. 1 mg/mL polyethyleneimine (PEI) HCl MAX, Linear, Mw 40,000 solution. 7. Plasmids. (a) Caffeine-sensitive receptor based on the EpoR-IL6ST/ STAT3 receptor system (PhCMV-aCaffVHH-EpoRIL6ST-pA, pDB306) [22]. (b) Reporter plasmid for STAT3 signaling activity (PSTAT3SEAP-pA, pLS13) [25]. (c) Signaling potentiator (PhCMV-STAT3-pA, pLS15) [25]. (d) pcDNA3.1(+) (PhCMV-MCS-pA) (Invitrogen). (e) pSEAP2-Control Note 1).

(PSV40-SEAP-pA)

(Clontech)

(f) PhCMV-driven scFv-EpoR-IL6ST expression (PhCMV-scFv-EpoR-IL6ST-pA, pLeo730).

(see

vector

(g) γ-butyrolactone (SCB1)-repressible SEAP expression vector (PSPA-SEAP-pA, pWW124). 8. Oligonucleotides. (a) oDB456 (50 - CAAGGATCCGGCTCTCAGGTTCAATT GGTGGAATCTGGAGGGGGTCTCGTACAGGCAGG CGGTTCTCTCCGACTGAGTTGCACAGCCTCCGG TAGGACTGGGACCAT-30 ). (b) oDB457 (50 - TCCATGTAGTATGTGATCCCAGAACT CCAACCTACAGTGGCAAGAAACTCTCTTTCTTTG CCTGGGGCCTGGCGAAACCAGGCCATTGAGTAG ATGGTCCCAGTCCTACC-30 ). (c) oDB458 (50 - CTGGGATCACATACTACATGGATTCA GTTAAAGGAAGATTCACTATCAGCCGAGATAAAG GGAAAAATACTGTGTACCTCCAGATGGACTCTCT GAAACCGGAGGACACG-30 ). (d) oDB459 (50 - TAAGAATTCGCTAGAGACGGTTACCT GTGTTCCTTGCCCCCAGTAGTCATACCCTACGGA GTAGGCGCGGGTGGCTGTACAGTAGTAGACAGC CGTGTCCTCCGGTTTCA-30 ). (e) OLS40 (50 -ACTTCGAAGCTTGCCACCATGGCCCAA TGGAATCAGCTAC-30 ). (f) OLS41 (50 - ACTTCGCTCGAGTCACATGGGGGAGG TAGCGCAC-30 ). (g) OLS10 (50 - ACTTCGGACGTCGTCGACATTTCCCG TAAATCGTCGAGTCGACATTTCCCGTAAATCGTC GACCTGCAGGTCGAGCTCGGTACCCGGGTC-30 ).

Construction of Caffeine-Inducible Gene Switches in Mammalian Cells

163

(h) OLS19 (50 - ACTTCGGAATTCCCGCGGAGGCTG GATCGGTC-30 ). 9. Restriction enzymes: AatII, BamHI, EcoRI, HindIII, XhoI. 10. Plasmid purification kit. 11. Sterile cell culture plasticware (100 mm petri dishes; 24-well plates). 12. Tabletop centrifuge. 13. Caffeine. 2.2 SEAP Reporter Gene Expression Assay

1. SEAP buffer (2x): 20 mM homoarginine, 1 mM MgCl2, 21% (v/v) diethanolamine, pH 9.8. 2. Substrate for SEAP: 120 mM para-nitrophenyl phosphate (pNPP) in 2 SEAP buffer. 3. 96-well plates. 4. Microplate reader.

3

Methods

3.1 Design Principles for Constructing a Caffeine-Inducible Gene Expression System

1. Currently, the most convenient manner to construct a caffeineinducible gene expression system relies on the caffeine nanobody aCaffVHH which homodimerizes in the presence of caffeine (see Note 2). Therefore, any signaling pathway or transcription factor activated by caffeine should be activatable by dimerization. Further, systems relying on heterodimerization are certainly possible in principle, yet they will have a lower efficiency with aCaffVHH as unproductive homodimers will be formed as well. 2. The choice of signaling system will also determine the caffeine sensitivity of the resulting gene expression system. While the affinity of aCaffVHH to caffeine will not change in different fusion proteins, the fraction of receptors which need to be dimerized for productive signaling is different for different signaling pathways. Coupling aCaffVHH to a receptor in a signaling pathway with a high degree of signal amplification will result in a gene expression system which requires lower caffeine concentrations for the induction of transgene expression. 3. The choice of synthetic promoter architecture will influence both inducer sensitivity and protein production. The number of binding elements of the transcription factor which is ultimately activated is a major factor in determining the needed inducer concentrations and the final ON/OFF ratio (see Note 3). Additional features, such as the choice of minimal promoter, the addition of a Kozak sequence, and the choice of

164

Daniel Bojar

polyadenylation site, will also influence protein production, mostly by varying the total amount of gene expression. 3.2 Construct Plasmids for a Caffeine-Inducible Gene Expression System

For this exemplary construction of a caffeine-inducible gene expression system, plasmids for the caffeine receptor, signaling amplifier, and reporter gene are described. 1. Construct a plasmid encoding the caffeine receptor [22] (see Note 4). We cloned aCaffVHH-EpoR-IL6ST, a caffeinedimerizable receptor activating JAK/STAT3 signaling upon stimulation under the control of tshe constitutive human cytomegalovirus promoter (PhCMV-aCaffVHH-EpoR-IL6ST-pA, pDB306). For this, aCaffVHH was PCR-assembled from the overlapping oligonucleotides oDB456, oDB457, oDB458 and oDB459. This product was then restricted with BamHI/EcoRI and cloned into the corresponding sites (BamHI/EcoRI) of pLeo730. 2. Construct a plasmid encoding the signaling amplifier STAT3 [25] (PhCMV-STAT3-pA; see Note 5). For this, human STAT3 was PCR-amplified using the oligonucleotides OLS40 and OLS41. The product was then restricted with HindIII/XhoI and ligated into the corresponding sites (HindIII/XhoI) of pcDNA3.1(+). 3. Construct a STAT3 reporter plasmid to monitor signaling induction by caffeine (see Note 6). To enable quantification of gene expression with the reporter gene secreted placental alkaline phosphatase (SEAP), two STAT3 binding sites were combined with a minimal promoter to drive SEAP expression (PSTAT3-SEAP-pA). These binding sites were PCR-amplified using oligonucleotides OLS10 and OLS19. The product was restricted using AatII/EcoRI and ligated into the corresponding sites (AatII/EcoRI) of pWW124.

3.3 Cell Cultivation and Transfection

While cell culture and transfection conditions have to be optimized for every cell line, this protocol is optimized for HEK-293T cells. 1. Cultivate HEK-293 T cells in 10 cm dishes filled with 10 mL of DMEM (+ 10% FBS and 1% penicillin/streptomycin) in a humidified atmosphere containing 5% CO2 at 37  C. 2. At about 80% confluency, remove the medium completely and add 0.8 mL of 0.05% (v/v) trypsin/EDTA solution. Incubate cells for 3 min at 37  C. 3. Carefully detach the cells by gentle pipetting, transfer them into a conical 15 mL Falcon tube, and centrifuge for 3 min at 450  g. 4. Discard supernatant and resuspend cells in 5 mL of DMEM. 5. Count cell concentration using a cell counter and seed wells of a 24-well plate with 0.5 mL of 2.5  105 cells/mL in DMEM

Construction of Caffeine-Inducible Gene Switches in Mammalian Cells

165

per well (see Note 7). Incubate cells for 12–24 h at 37  C and 5% CO2. 6. For each well to be transfected, combine 500 ng plasmid DNA in microtube (see Note 8) and fill to 50 μL with FBS-free DMEM. Add 2.5 μL 1 mg/mL PEI, gently mix, and incubate for 15–30 min at room temperature (see Note 9). 7. Exchange medium of seeded cells with the same volume of fresh medium. 8. After incubation, add DNA-PEI mixture dropwise to cells and mix by gently rocking the plate back and forth. 9. Place cells back into incubator at 37  C and 5% CO2. Replace medium with fresh medium after 8–12 h (see Note 10). 10. Incubate for 24–48 h at 37  C and 5% CO2 before assaying protein expression. 3.4 Testing the Caffeine-Inducible Gene Expression System

1. For best results, I advise the transfection of 5 ng pDB306 (PhCMV-aCaffVHH-EpoR-IL6ST-pA), 100 ng pLS15 (PhCMV-STAT3-pA), 150 ng pLS13 (PSTAT3-SEAP-pA), and 245 ng of pcDNA3.1(+) per well of a 24-well plate. 2. Eight hours after transfection, when replacing the medium, add caffeine in concentrations ranging from 0 to 100 μM to the medium (see Note 11). For the most robust results, aim for at least biological triplicates for each concentration. 3. Collect cell culture supernatant 24 h after caffeine addition. Transfer 100 μL per well into 96 well plates (technical triplicates for each biological sample), seal plate, and heat-inactivate endogenous phosphatases by incubation at 65  C for 30 min. 4. Let plate cool down to room temperature and spin down at 14,000  g for 2 min to precipitate cell debris. 5. Transfer 80 μL of each well into a fresh 96 well plate and add 100 μL 2 SEAP buffer as well as 20 μL of pNPP substrate solution (see Note 12). 6. Measure absorbance at 405 nm every 30 s for 30 min with a microplate reader and quantify SEAP protein levels using Lambert-Beer’s law, E ¼ a  c  d (E ¼ increase in absorbance per minute, a ¼ molar extinction coefficient of the product p-nitrophenolate (a ¼ 18,600 M1 cm1), c ¼ increase in pnitrophenolate (M min1), d ¼ length of the light path in the sample (usually 0.5 cm for a 96-well plate). To arrive at SEAP units/liter (U/L), use the formula c  106  (200/80) (dilution factor).

166

4

Daniel Bojar

Notes 1. Constitutive expression of SEAP with a plasmid such as pSEAP2-Control (PSV40-SEAP-pA) is an excellent positive control to optimize transfection and assay conditions as well as compare to inducible gene expression results. 2. Alternatives to construct a caffeine-inducible gene expression system which are based on the combination of a caffeine demethylase with a theophylline-responsive RNA switch have only been tested in Saccharomyces cerevisiae yet and are characterized by a lower sensitivity with regard to caffeine [26]. 3. While an increased number of transcription factor binding sites will likely increase both inducer sensitivity as well as final protein production levels, an elevated level of background activity is also a frequent consequence. In practice, the acceptable amount of leakiness as well as the ON/OFF-induction ratio has to be set in the context of each specific project and its requirements. 4. In various projects, aCaffVHH has been fused to multiple proteins, demonstrating its flexibility and modularity. Both N- as well as C-terminal fusions are compatible with caffeineinduced dimerization. Even intra-protein-fusions, in which aCaffVHH was sandwiched between two proteins, have performed well. Both short, rigid as well as more loose linkers can be paired with aCaffVHH. 5. Several cell lines, such as HEK-293T cells, endogenously express STAT3. Therefore, transfection and overexpression of this gene is not necessarily needed for signaling. However, even in cases in which cells endogenously expressed STAT3, transfection of additional STAT3 led to increased SEAP production and improved ON/OFF ratios, indicating the bottleneck role of STAT3 in this signaling setup. 6. While SEAP is used here as an exemplary reporter gene, in principle any gene capable of being expressed in mammalian cells could be inserted here. 7. It is important that the indicated cell number only contains viable cells. If only a certain fraction of the cells in question are viable, increase the seeded cell number accordingly to reach the indicated number of viable cells. 8. The ratio of DNA and PEI is paramount for transfection efficiency. Therefore, even if the necessary amount of DNA for a given experiment in a 24-well plate well would be lower than 500 ng, it is strongly recommended to add a filler plasmid such as pcDNA3.1(+) to the DNA mix to ensure optimal transfection efficiency.

Construction of Caffeine-Inducible Gene Switches in Mammalian Cells

167

9. While HEK-293T cells and various other cell lines can be transfected easily with PEI, other cells such as embryonic stem cells and primary cells in general can be very refractory toward transfection with PEI. For these cells, other routes of gene transfer need to be taken, such as lentiviral transduction or plasmid electroporation. 10. For several cell types, prolonged incubation with PEI can be toxic. However, HEK-293 T cells seem to be resistant to PEI toxicity. Therefore, HEK-293 T cells can exhibit high viability even if the DNA-PEI mixture is never removed from transfected cells. 11. Especially with novel caffeine-inducible gene expression systems, it is advisable to cover a large range of caffeine concentrations in the beginning, as different caffeine receptor setups have shown sensitivity differences spanning several orders of magnitude. Caffeine is nontoxic to mammalian cells up to high micromolar concentrations, enabling a very broad dynamic range. 12. The SEAP substrate pNPP is light-sensitive and should therefore be kept at 20  C and wrapped in aluminum foil. However, SEAP itself is very stable and cell culture supernatant samples containing SEAP can be kept at 20  C for many months before measurement. References 1. Black JB, Perez-Pinera P, Gersbach CA (2017) Mammalian synthetic biology: engineering biological systems. Annu Rev Biomed Eng 19:249–277. https://doi.org/10.1146/ annurev-bioeng-071516-044649 2. Kim H, Bojar D, Fussenegger M (2019) A CRISPR/Cas9-based central processing unit to program complex logic computation in human cells. Proc Natl Acad Sci 116:7214–7219. https://doi.org/10.1073/ pnas.1821740116 3. Ausl€ander D, Ausl€ander S, Pierrat X, Hellmann L, Rachid L, Fussenegger M (2017) Programmable full-adder computations in communicating three-dimensional cell cultures. Nat Methods 15:57–60. https://doi. org/10.1038/nmeth.4505 4. Tastanova A, Schulz A, Folcher M, Tolstrup A, Puklowski A, Kaufmann H, Fussenegger M (2016) Overexpression of YY1 increases the protein production in mammalian cells. J Biotechnol 219:72–85. https://doi.org/10. 1016/j.jbiotec.2015.12.005 5. Kojima R, Bojar D, Rizzi G, Hamri GC-E, El-Baba MD, Saxena P, Ausl€ander S, Tan KR,

Fussenegger M (2018) Designer exosomes produced by implanted cells intracerebrally deliver therapeutic cargo for Parkinson’s disease treatment. Nat Commun 9:1305. https://doi.org/10.1038/s41467-01803733-8 6. Liu Y, Bai P, Woischnig A-K, Charpin-El Hamri G, Ye H, Folcher M, Xie M, Khanna N, Fussenegger M (2018) Immunomimetic designer cells protect mice from MRSA infection. Cell 174:259–270.e11. https://doi.org/10.1016/j.cell.2018.05.039 7. Gossen M, Bujard H (1992) Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc Natl Acad Sci U S A 89:5547–5551 8. Gitzinger M, Kemmer C, Fluri DA, Daoud El-Baba M, Weber W, Fussenegger M (2012) The food additive vanillic acid controls transgene expression in mammalian cells and mice. Nucleic Acids Res 40:e37. https://doi.org/10. 1093/nar/gkr1251 9. Xie M, Ye H, Hamri GC-E, Fussenegger M (2014) Antagonistic control of a dual-input mammalian gene switch by food additives.

168

Daniel Bojar

Nucleic Acids Res 42:e116. https://doi.org/ 10.1093/nar/gku545 10. Gitzinger M, Kemmer C, El-Baba MD, Weber W, Fussenegger M (2009) Controlling transgene expression in subcutaneous implants using a skin lotion containing the apple metabolite phloretin. Proc Natl Acad Sci U S A 106:10638–10643. https://doi.org/10. 1073/pnas.0901501106 11. Yamada M, Suzuki Y, Nagasaki SC, Okuno H, Imayoshi I (2018) Light control of the Tet gene expression system in mammalian cells. Cell Rep 25:487–500 e6. https://doi.org/10. 1016/j.celrep.2018.09.026 12. Morsut L, Roybal KT, Xiong X, Gordley RM, Coyle SM, Thomson M, Lim WA (2016) Engineering customized cell sensing and response behaviors using synthetic notch receptors. Cell 164:780–791. https://doi.org/10.1016/j. cell.2016.01.012 13. Abate-Daga D, Davila ML (2016) CAR models: next-generation CAR modifications for enhanced T-cell function. Mol Ther Oncolytics 3:16014. https://doi.org/10.1038/mto. 2016.14 14. Scheller L, Strittmatter T, Fuchs D, Bojar D, Fussenegger M (2018) Generalized extracellular molecule sensor platform for programming cellular behavior. Nat Chem Biol 14:723–729. https://doi.org/10.1038/s41589-018-0046-z 15. Saxena P, Bojar D, Fussenegger M (2017) Design of Synthetic Promoters for gene circuits in mammalian cells. Methods Mol Biol 1651:263–273. https://doi.org/10.1007/ 978-1-4939-7223-4_19 16. Xie M, Ye H, Wang H, Charpin-El Hamri G, Lormeau C, Saxena P, Stelling J, Fussenegger M (2016) β-cell-mimetic designer cells provide closed-loop glycemic control. Science 354:1296–1301. https://doi.org/10.1126/ science.aaf4006 17. Wikoff D, Welsh BT, Henderson R, Brorby GP, Britt J, Myers E, Goldberger J, Lieberman HR, O’Brien C, Peck J, Tenenbein M, Weaver C, Harvey S, Urban J, Doepker C (2017) Systematic review of the potential adverse effects of caffeine consumption in healthy adults, pregnant women, adolescents, and children. Food Chem Toxicol 109:585–648. https://doi.org/ 10.1016/j.fct.2017.04.002

18. Poole R, Kennedy OJ, Roderick P, Fallowfield JA, Hayes PC, Parkes J (2017) Coffee consumption and health: umbrella review of meta-analyses of multiple health outcomes. BMJ 359:j5024. https://doi.org/10.1136/ bmj.j5024 19. Zajac MA, Zakrzewski AG, Kowal MG, Narayan S (2003) A novel method of caffeine synthesis from uracil. Synth Commun 33:3291–3297. https://doi.org/10.1081/ SCC-120023986 20. Hugtenburg JG, Timmers L, Elders PJ, Vervloet M, van Dijk L (2013) Definitions, variants, and causes of nonadherence with medication: a challenge for tailored interventions. Patient Prefer Adherence 7:675–682. https://doi.org/10.2147/PPA.S29549 21. Franco EJ, Sonneson GJ, DeLegge TJ, Hofstetter H, Horn JR, Hofstetter O (2010) Production and characterization of a genetically engineered anti-caffeine camelid antibody and its use in immunoaffinity chromatography. J Chromatogr B 878:177–186. https://doi. org/10.1016/j.jchromb.2009.06.017 22. Bojar D, Scheller L, Hamri GC-E, Xie M, Fussenegger M (2018) Caffeine-inducible gene switches controlling experimental diabetes. Nat Commun 9:2318. https://doi.org/10. 1038/s41467-018-04744-1 23. Bojar D, Fuhrer T, Fussenegger M (2019) Purity by design: reducing impurities in bioproduction by stimulus-controlled global translational downregulation of non-product proteins. Metab Eng 52:110–123. https:// doi.org/10.1016/j.ymben.2018.11.007 24. Bojar D, Fussenegger M (2019) The role of protein engineering in biomedical applications of mammalian synthetic biology. Small 16:1903093. https://doi.org/10.1002/smll. 201903093 25. Schukur L, Geering B, Charpin-El Hamri G, Fussenegger M (2015) Implantable synthetic cytokine converter cells with AND-gate logic treat experimental psoriasis. Sci Transl Med 7:318ra201. https://doi.org/10.1126/ scitranslmed.aac4964 26. Michener JK, Smolke CD (2012) Highthroughput enzyme evolution in Saccharomyces cerevisiae using a synthetic RNA switch. Metab Eng 14:306–316. https://doi.org/10.1016/j. ymben.2012.04.004

Part III Precise Genome Engineering Techniques Using CRISPR-Cas Systems

Chapter 11 Multiplexed Genome Engineering with Cas12a Niels R. Weisbach, Ab Meijs, and Randall J. Platt Abstract Genome engineering technologies based on CRISPR-Cas systems are fueling efforts to study genotype–phenotype relationships in a high-throughput and multiplexed fashion. While many genome engineering technologies exist and provide a means to efficiently manipulate one or a few genes in a singular contextknockout, inhibition, or activation in a constitutive, conditional, or inducible manner-progress towards engineering complex cellular programs has been hampered by the lack of technologies that can integrate these functions within a unified framework. To address this challenge, our lab created single transcript CRISPR-Cas12a (SiT-Cas12a), which enables conditional, inducible, orthogonal, and massively multiplexed genome engineering of dozens, to potentially hundreds, of genomic targets in eukaryotic cells simultaneously-providing a novel way to interrogate and engineer complex genetic programs. In this chapter, we outline the utility of SiT-Cas12a in human cells and describe experimental procedures for executing massively multiplexed genome engineering experiments-including strategies for designing and assembling customized multiplexed CRISPR guide RNA arrays as well as validating and analyzing CRISPR guide RNA array processing and genome engineering outcomes. Key words Genome engineering, CRISPR, Cas12a, CRISPR-Cas12a, Multiplexed, Orthogonal genome engineering, CRISPR array synthesis, Gene editing, Transcriptional regulation

1

Introduction Genome engineering technologies are valuable for interrogating complex cellular processes and linking genotype to phenotype. Current platforms, such as RNA-guided CRISPR–Cas enzymes have greatly expanded the genome engineering toolbox-containing programmable nucleases, transcription factors, epigenetic modifiers, transposases, base editors, prime editors, and many others [1–3]-which collectively enable the manipulation of many individual features of a cell. So far, these technologies are generally only used to make one or a few manipulations within a cell at a time. However, cells orchestrate their behaviors through a complex architecture of gene interactions, networks, and feedback-making it desirable, if not necessary, to make multiple perturbations at once to control and understand a cellular processes [4]. Over the past

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

171

172

Niels R. Weisbach et al.

few years, the field has moved towards multiplexed genome engineering to provide increased control over cellular phenotypes [5]. Here, we briefly summarize efforts to multiplex CRISPR-Cas systems and also highlight our recently developed technology termed single transcript Cas12a (SiT-Cas12a), which provides a scalable platform for massively multiplexed and orthogonal genome engineering in mammalian cells [6]. The first CRISPR-Cas gene editing technology was based on the CRISPR-associated (Cas) enzyme Cas9 from Streptococcus pyogenes (SpCas9, here referred to as Cas9). Cas9 is a DNA endonuclease that uses either a dual RNA guide-composed of a transactivating RNA (tracrRNA) and a CRISPR RNA (crRNA)-or a single guide RNA (sgRNA)-composed of an engineered fusion of tracrRNA and crRNA-to identify target sites via Watson-Crick base pairing and induce DNA double stranded breaks in human cells [7– 10]. Multiplexed genome engineering with Cas9 can be achieved using one of three general strategies. The first strategy mimics the natural CRISPR interference process wherein the crRNA and tracrRNA are individually expressed under polymerase III promoters while the Cas9 nuclease is expressed under a polymerase II promoter (Fig. 1a) [8]. The crRNA and tracrRNA form a complex through hybridization of the direct repeat region; thus, for each additional target a different crRNA has to be expressed individually. Nevertheless, this approach resulted in suboptimal editing efficiency in mammalian cells. The second strategy is based on the coexpression of crRNA and tracrRNA as sgRNA, driven by a single polymerase III promoter. To achieve multiplexed capabilities, each sgRNA is expressed under an individual polymerase III promoter (Fig. 1b) [8, 9]. The third strategy is based on RNA processing enzymes or self-cleaving sequences, such as Csy4 or ribozymes, respectively (Fig. 1c) [11]. This can either be achieved through coexpression of exogeneous proteins or by utilizing endogenous proteins [11]. However, the necessity for multipromoter systems, additional RNA processing enzymes, or the integration of multiple long and highly structured elements within the crRNA array, exacerbate further scaling ambitions while maintaining proficient editing efficiency in mammalian cells, paving the way for novel approaches to achieve efficient multiplexing capabilities. Since the discovery of Cas9, several additional RNA-guided Cas nucleases have been discovered and leveraged for genome engineering [3, 12]. One enzyme that shows particular promise for multiplexed genome engineering is Cas12a (formerly known as Cpf1) [13–15]. Cas12a possesses a dual DNase/RNase function, which enables RNA-guided DNA cleavage and the maturation of its own pre-crRNA array into mature single crRNAs without coexpression of additional RNA processing enzymes [13, 16]. Furthermore, the corresponding crRNA of Cas12a is shorter and lessstructured compared to Cas9 sgRNAs making them easier to

Multiplexed Genome Engineering with Cas12a

173

Fig. 1 Multiplexed genome editing with Cas9. (a) Schematic representation of multiplexed genome editing based on Cas9 (blue), tracrRNA (purple), and a pre-crRNA array (colored rectangles and gray diamonds). Transcription of the tracrRNA and a pre-crRNA array are driven by individual polymerase III promoters, enabling the hybridization of a crRNA–tracrRNA complex. Cas9 binds to the crRNA–tracrRNA complex and induces a DNA double strand break at the genomic target site. (b) Schematic representation of multiplexed genome editing based on Cas9 and chimeric single guide RNAs (sgRNAs). Polymerase III driven expression of sgRNAs-fusions of crRNAs and tracrRNAs-facilitates direct interaction with Cas9 and leads to targeted genome editing. (c) Schematic representation of multiplexed genome editing based on Cas9 and multiple sgRNAs achieved through RNA processing. Expression of an array of sgRNAs-driven by a polymerase II or polymerase III promoter-where each sgRNA is flanked by RNA processing sequences (e.g., tRNA, ribozyme, splice sites, Csy4 recognition sequences) enables the production of multiple mature sgRNAs. Cas9 recognizes the mature sgRNAs and subsequently facilitates genome editing

synthesize and encode within a compact multiplexed array. Yet the first multiplexed implementations of Cas12a had an approach similar to Cas9 systems, which included the necessity for multiple promoters to transcribe the nuclease and the crRNA array [13]. To harness the full potential of Cas12a for multiplexed genome engineering, our lab developed a single transcript (SiT) version, which enables expression of Cas12a from a single polymerase II promoter together with an almost limitless number of crRNAs [6]. The SiT-Cas12a platform is composed of (1) Cas12a or catalytic inactive Cas12a fused to an effector domain (e.g., transcriptional regulator), (2) crRNA array, and (3) a triplex sequence to stabilize the Cas12a mRNA transcript after crRNA processing (Fig. 2a, b). Because we use a polymerase II promoter, we can take full advantage of the plethora of regulatable expression systems, such as conditional and inducible promoters, that largely do not exist for polymerase III promoters. Another advantage of this technology lies in the capacity to perform orthogonal genome engineering wherein catalytically active Cas12a is fused to a transcriptional activation or repression domain and depending on the length of the crRNA and where it targets a gene either gene knockout (full-length crRNA targeting an exon) or transcriptional

174

Niels R. Weisbach et al.

Fig. 2 Multiplexed orthogonal genome editing and transcriptional regulation using SiT-Cas12a. (a) Schematic representation of the SiT-Cas12a architecture for genome editing. The SiT-Cas12a transcript encodes for Cas12a (yellow); Triplex structure (purple); crRNA array consisting of direct repeat sequences (gray diamonds), spacer sequences (colored rectangles), and a poly(A) (red). (b) Schematic of a modified SiT-Cas12a platform for transcriptional modulation enabled by effector domain (ED) protein fusions—facilitating either transcriptional activation [Activ] or repression [Repr]. The SiT-ddCas12a-ED transcript encodes for DNase-dead Cas12a (ddCas12a, yellow) fused to an effector domain (ED, gray). (c) Schematic of a SiT-Cas12a platform for orthogonal genome editing and transcriptional modulation. Catalytically active Cas12a-[ED] can be utilized for either genome editing, through binding of a full length crRNA (23 nt) targeting coding sequences, or for transcriptional control, through binding of a truncated crRNA (15 nt) targeting upstream of a transcriptional start site

modulation (truncated crRNA targeting upstream of the transcriptional start site) can be achieved (Fig. 2c). Together, this highlights the scalability and flexibility of the SiT-Cas12a platform, which in contrast to other CRISPR-based technologies does not rely on numerous promoters and components to implement sophisticated genome engineering programs. In the future, the unique capabilities of SiT-Cas12a will fuel efforts toward interrogating and engineering complex cellular programs. In this chapter, we describe the generation and utilization of SiT-Cas12a for multiplexed genome engineering of human cells. Using one of our SiT-Cas12a plasmids as a starting point, we provide detailed instructions for the construction of customized orthogonal SiT-Cas12a systems. Then, we describe how to design crRNAs and explain various strategies to synthesize and construct crRNA arrays within SiT-Cas12a plasmids. Finally, we describe protocols for performing and analyzing multiplexed gene editing and transcriptional modulation experiments with SiT-Cas12a. In summary, the instructions and guidelines discussed in this chapter enable the general usage of SiT-Cas12a platforms for multiplexed genome engineering of mammalian cell lines and facilitate the creation of customized genome engineering technologies.

Multiplexed Genome Engineering with Cas12a

2 2.1

175

Materials Plasmids

2.2 Molecular Reagents

The SiT-Cas12a plasmids are depicted in Table 1 and are available from Addgene (www.addgene.org/browse/article/28203590). 1. UltraPure DNase/RNase-free distilled water. 2. Custom DNA oligos. 3. gBlock Gene Fragments (dsDNA fragment) (IDT). 4. Custom Gene Synthesis (plasmid synthesis) (IDT). 5. Dithiothreitol (DTT), 1 M. 6. Stbl3 chemically competent E. coli. 7. LB (Luria Broth) media. 8. LB (Luria Broth) plates. 9. Ethanol. 10. Ampicillin sodium salt. 11. SOC (Super Optimal Broth) media. 12. Triton X-100. 13. UltraPure Tris–HCl, 1 M. 14. UltraPure 0.5 M EDTA, pH 8.0. 15. Proteinase K. 16. Phusion Flash High-Fidelity PCR Master Mix. 17. E-Gel 48 agarose gels, 2%. 18. 4–20% TBE polyacrylamide gels. 19. SYBR Gold Nucleic Acid Gel Stain (10,000
Concentrate). 20. qScript cDNA SuperMix. 21. EvaGreen qPCR Master Mix.

2.3

Kits

1. Mini preparation Kit (Qiagen, cat. no.: 27106). 2. Quick RNA Miniprep Plus kit (Zymo Research, cat. no.: R1058).

Table 1 SiT-Cas12a plasmids available at Addgene as genome engineering toolbox Plasmid #

Plasmid description

Addgene ID

pCI108

EF1a-loxp-stop-loxp-SiT-Cas12a

128407

pCI118

minCMV-SiT-Cas12 under Tetracycline-inducible promoter

128406

pCI152

CMV-SiT-Cas12a in lentiviral backbone

128405

pCE046

EF1a-SiT-Cas12a couple with repressor domain

128132

pCE048

EF1a-SiT-Cas12a

128124

pCE059

EF1a-SiT-Cas12a coupled with activator domain

128136

176

Niels R. Weisbach et al.

3. DNA Clean & Concentrator (Zymo Research, cat. no.: D4004). 4. SURVEYOR® Mutation Detection Kit (IDT, cat no.: 72993204/706020). 5. QIAquick Gel Extraction Kit (Qiagen, cat no.: 28706). 6. Gibson Assembly Cloning Kit (New England BioLabs, cat no.: E5510S). 2.4

Enzymes

1. BamHI (Fast Digest, Thermo Fisher Scientific). 2. Esp3I (BsmBI) (Fast Digest, Thermo Fisher Scientific). 3. LguI (SapI) (Fast Digest, Thermo Fisher Scientific). 4. FastAP-Thermosensitive Alkaline Phosphatase (Thermo Fisher Scientific). 5. T7 DNA Ligase. 6. T4 DNA Ligase.

2.5 Human Cell Culture

1. Cell line: Human embryonic kidney 293T cell line. 2. Culture medium: Dulbecco’s Eagle’s Medium (DMEM) with 10% FBS and 10 mM HEPES. 3. TrypLE. 4. PBS, pH 7.4. 5. Lipofectamine 2000 Transfection Reagent. 6. Opti-MEM I Reduced Serum Medium.

2.6 Special Equipment 2.7 Sequencing Primers

1. E-Gel Electrophoresis System. 2. Real-Time PCR Thermocycler. The sequencing primers in Table 2 can be used to validate cloning products for all SiT-Cas12a plasmids (see Table 1).

Table 2 Sequencing primers for verification of SiT-Cas12a plasmids by Sanger sequencing Name

Sequence

Cas12a forward primer

GAGAAGATGCTGAACGCGAAGCTGAAGGATCAG

Triplex reverse primer

GCCAAAAAGCAAAACCTGAGAAAA

Triplex forward primer

TTTTCTCAGGTTTTGCTTTTTGGC

Multiplexed Genome Engineering with Cas12a

3

177

Methods The single transcript Cas12a (SiT-Cas12a) platform enables a broad range of applications in mammalian cells. To further expand the scope of usage, individual adjustments of SiT-Cas12a plasmids (see Table 1) can be achieved by changing the effector domain (see Subheading 3.1) and by incorporating individual crRNA arrays of different sizes (see Subheading 3.2). Validation of SiT-Cas12abased genome engineering events in mammalian cells is described in detail for genome editing (see Subheading 3.4) and transcriptional regulation (see Subheading 3.5).

3.1 Construction of Customized SiT-Cas12a Effector Plasmids

SiT-Cas12a plasmids can be utilized for multiplexed genome engineering and transcriptional regulation in mammalian cells and are available from Addgene (see Table 1). To leverage the SiT-Cas12a platform for multiplexed orthogonal genome engineering in a customized way, an effector domain (e.g., base editor, epigenetic regulators) can be inserted at the 30 end of Cas12a. Below, we describe in detail how to synthesize a customized SiT-Cas12a plasmid platform with the addition of an effector domain. 1. Construction of the SiT-Cas12a plasmid backbone. All SiT-Cas12a plasmids listed in Table 1 harbor a BamHI restriction site downstream of Cas12a, which allows the incorporation of the customized effector domain along with an appropriate linker peptide. Digest the SiT-Cas12a plasmid backbone using BamHI and add thermosensitive alkaline phosphatase (FastAP) to prevent self-ligation. X μl

SiT-Cas12a plasmid (2 μg)

2 μl

BamHI (FastDigest)

1 μl

FastAP

3 μl

10
FastDigest buffer

Y μl

H2O

30 μl

Total

Incubate the reaction mixture with the following thermocycler parameters to achieve complete SiT-Cas12a plasmid backbone digestion at optimal enzymatic conditions and subsequently inactivate the enzymes. 37 ˚C

60 min

80 ˚C

10 min

4 ˚C

Hold

178

Niels R. Weisbach et al.

Purify the digested SiT-Cas12a plasmid backbone mixture with any DNA purification kit (e.g., Zymo Research) and elute the DNA in 20 μl Elution Buffer. 2. Preparation of the customized effector domain. To successfully clone the effector domain into the SiT-Cas12a plasmid backbone, add two BamHI restriction sites (“GGATCC”) at the 50 and 30 ends of the effector domain. Design and order the construct as a dsDNA fragment (e.g., gBlock (IDT)). Digest the construct using BamHI. X μl

Effector domain (2 μg)

2 μl

BamHI (FastDigest)

3 μl

10
FastDigest buffer

Y μl

H2O

30 μl

Total

Incubate the reaction mixture in a thermocycler with the following program to obtain a dsDNA fragment with suitable overhang sequences for cloning into the SiT-Cas12a plasmid backbone. 37 ˚C

60 min

80 ˚C

10 min

4 ˚C

Hold

Purify the digested dsDNA fragment of the effector domain with any DNA purification kit (e.g., Zymo Research) and elute the DNA fragment in 20 μl Elution Buffer. Purified dsDNA fragments can be stored at 20  C or directly used as an insert for the ligation reaction. 3. Ligation of the customized SiT-Cas12a effector plasmid. Ligate the digested SiT-Cas12a plasmid backbone (Vector) and the digested effector domain (Insert) in the following ratio to obtain the customized SiT-Cas12a effector plasmid. X μl

Vector (50 fmol)

Z μl

Insert (150 fmol)

10 μl

2
T7 reaction buffer

1 μl

T7 DNA ligase

Y μl

H2O

20 μl

Total

Multiplexed Genome Engineering with Cas12a

179

Incubate the reaction mixture at room temperature for at least 30 min before transforming into competent cells. 4. Transformation and plasmid preparation of the SiT-Cas12a effector plasmid. Add 5 μl of the ligation mixture into a thawed shot of competent E. coli Stbl3 cells and keep the cell suspension on ice for 20 min. Subsequently, heat shock the cell mixture for 30 s at 42  C and immediately chill the cells on ice for 5 min. Mix the transformed cells with 1 volume of prewarmed SOC medium (37  C) and spread the cell suspension on a LB agar plate containing 100 μg/ml ampicillin followed by overnight incubation at 37  C. Inoculate single colonies in LB media containing 100 μg/ml ampicillin overnight at 37  C and 300 rpm. Isolate the plasmid DNA on the next day using any DNA purification kit (e.g., Qiagen Mini preparation kit). 5. Verification of the SiT-Cas12a effector plasmid by Sanger sequencing. Verify the customized SiT-Cas12a effector plasmid with Sanger sequencing (any Sanger sequencing service) by using the Cas12a forward primer or the Triplex reverse primer, listed in Table 2. 3.2 Generation and Incorporation of crRNA Arrays into SiT-Cas12a Plasmids

The SiT-Cas12a plasmids enable orthogonal genome engineering in mammalian cells and require the cloning of crRNA arrays to realize multiplexed capabilities. The optimal method to assemble crRNA arrays depends on the goal, as well as the size, and composition of the crRNA array (see Notes 1–3). We describe three methods that can be utilized in detail below.

3.2.1 Assembly and Incorporation of Small crRNA Arrays into SiT-Cas12a Plasmids

The assembly of small crRNA arrays is limited to a maximum of 3 spacers, separated by direct repeat (DR) sequences, and is based on dsDNA synthesis, such as Gene Fragments Synthesis (IDT) (see Note 4). Synthesized crRNA arrays can be directly incorporated into any SiT-Cas12a plasmid shown in Table 1. 1. Construction of the SiT-Cas12a plasmid backbone. SiT-Cas12a plasmids harbor Type IIS restriction sites (BsmBI for pCI108, pCI118, and pCI152 or SapI for pCE059, pCE046, and pCE048) between two DR sequences which facilitate easy incorporation of crRNA arrays (Fig. 3a). SiT-Cas12a plasmids need to be digested with the respective restriction enzyme. For plasmid pCE046, pCE048, or pCE059, the following composition of the reaction mixture is recommended.

180

Niels R. Weisbach et al.

Fig. 3 SiT-Cas12a platform for systematic crRNA array incorporation. (a) Schematic representation of the SiT-Cas12a plasmid backbone for incorporating crRNA arrays. The SiT-Cas12a transcript encodes for Cas12a (yellow), Triplex structure (purple), Direct repeat sequence (gray diamond), Stuffer sequence (dashed rectangle), and poly(A) (red). The stuffer sequence contains Type IIS restriction sites which enable staggered cleavage of adjacent direct repeat sequences. (b) Cloning strategy for small crRNA arrays (up to 3 spacers) into the SiT-Cas12a plasmid backbone. Digestion of the SiT-Cas12a plasmid and synthesized crRNA array with Type IIS restriction enzymes results in complementary overhangs that facilitate ligation

X μl

SiT-Cas12a plasmid (2 μg)

2 μl

SapI (FastDigest)

2 μl

10
FastDigest buffer

Y μl

H2O

20 μl

Total

For plasmid pCI108, pCI118, or pCI152, supplement the reaction mixture with 1 mM DTT according to the manufacturer’s protocol. X μl

SiT-Cas12a plasmid (2 μg)

2 μl

BsmBI (FastDigest)

0.2 ul

DTT (100 mM)

2 μl

10x FastDigest buffer

Y μl

H2O

20 μl

Total

To achieve complete digestion of the SiT-Cas12a plasmid backbone, incubate the reaction mixture with the following parameters in a thermocycler. 37 ˚C

60 min

80 ˚C

10 min

4 ˚C

Hold

Multiplexed Genome Engineering with Cas12a

181

Purify the digested SiT-Cas12a plasmid backbone with any DNA purification Kit (e.g., Zymo Research) and elute the DNA in 20 μl Elution Buffer. 2. Design of small crRNA arrays. The digested SiT-Cas12a plasmid backbone harbors a region of insertion that contains DR sequences with overhangs at the 50 and 30 ends and enables the incorporation of crRNA arrays (Fig. 3b). Design and order the crRNA arrays as dsDNA fragments, containing up to 3 spacer sequences, separated by DR sequences (50 -Spacer1-DR-Spacer2-DR-Spacer3-30 ), with suitable 50 and 30 overhang sequences along with Type IIS restriction sites (see Note 4). It is essential that the 50 and 30 overhang sequences align with the Type IIS restriction sites of the digested SiT-Cas12a plasmid backbone. 3. Preparation of dsDNA fragments for restriction cloning. Digest the synthesized dsDNA fragment with the respective Type IIS restriction enzyme based on the SiT-Cas12a plasmid backbone. For SiT-Cas12a plasmid backbones based on plasmids pCE046, pCE048, or pCE059, the following composition of the reaction mixture is recommended. X μl

dsDNA fragment (200 ng)

2 μl

SapI (FastDigest)

2 μl

10
FastDigest buffer

Y μl

H2O

20 μl

Total

For SiT-Cas12a plasmid backbones based on plasmids pCI108, pCI118, or pCI152, supplement the reaction mixture with 1 mM DTT according to the manufacturer’s protocol. X μl

dsDNA fragment (200 ng)

2 μl

BsmBI (FastDigest)

0.2 μl

DTT (100 mM)

2 μl

10
FastDigest buffer

Y μl

H2O

20 μl

Total

Incubate the reaction mixture in a thermocycler with the following parameters to obtain a dsDNA fragment with suitable overhang sequences to ligate into the SiT-Cas12a plasmid backbone.

182

Niels R. Weisbach et al.

37 ˚C

60 min

80 ˚C

10 min

4 ˚C

Hold

Purify the digested dsDNA fragment with any DNA purification kit (e.g., Zymo Research) and elute the DNA in 20 μl Elution Buffer. 4. Ligation of SiT-Cas12a plasmid and verification by Sanger sequencing. Mix the digested and purified dsDNA fragment with the digested and purified SiT-Cas12a plasmid backbone in the following ratio. X μl

SiT-Cas12a plasmid backbone (50 fmol)

Z μl

dsDNA fragment (150 fmol)

10 μl

2
T7 reaction buffer

1 μl

T7 DNA ligase

Y μl

H 2O

20 μl

Total

Ligate the reaction at room temperature for 30 min and store the mixture at 20  C. Transform 5 μl of the ligation mixture as detailed in Subheading 3.1 (step 4) and verify the SiT-Cas12a plasmid by Sanger sequencing as detailed in Subheading 3.1 (step 5) using the Triplex forward primer (see Table 2). 3.2.2 Assembly and Incorporation of Medium crRNA Arrays into SiT-Cas12a Plasmids

The assembly process of medium crRNA arrays-harboring up to 10 crRNAs-can be accomplished by repeated PCR elongation of a customized dsDNA template, consisting of two spacers separated by a single DR sequence. After multiple rounds of PCR elongations, the crRNA array can be cloned into the SiT-Cas12a plasmid backbone by Gibson Assembly. 1. Construction of the SiT-Cas12a plasmid backbone. Digest the SiT-Cas12a plasmid with the respective restriction enzyme as described in Subheading 3.2.1 (step 1). For plasmid pCE046, pCE048, or pCE059 use the following composition of the reaction mixture. X μl

SiT-Cas12a plasmid (2 μg)

2 μl

SapI (FastDigest)

2 μl

10
FastDigest buffer

Y μl

H2O

20 μl

Total

Multiplexed Genome Engineering with Cas12a

183

For plasmid pCI108, pCI118, or pCI152, supplement the reaction mixture with 1 mM DTT according to the manufacturer’s protocol. X μl

SiT-Cas12a plasmid (2 μg)

2 μl

BsmBI (FastDigest)

0.2 μl

DTT (100 mM)

2 μl

10
FastDigest buffer

Y μl

H2O

20 μl

Total

To achieve complete digestion of the SiT-Cas12a plasmid backbone, incubate the reaction mixture with the following parameters in a thermocycler. 37 ˚C

60 min

80 ˚C

10 min

4 ˚C

Hold

Purify the digested SiT-Cas12a plasmid backbone with any DNA purification kit (e.g., Zymo Research) and elute the DNA in 20 μl Elution Buffer. 2. Construction of customized dsDNA templates. Design a dsDNA template that consists of two spacers (see Notes 1–3), separated by a DR sequence, and order the complementary strands as ssDNA oligos. To assemble the customized dsDNA template, mix 1 μl of each DNA oligo (100 μM) with 1 μl 10
Annealing Buffer (500 mM NaCl, 100 mM MgCl2, 400 mM Tris–HCl, pH 7.9) and dilute the reaction with H2O to a total volume of 10 μl. Incubate the reaction in a thermocycler with the following parameters. 95 ˚C

10 min

95 ˚C to 85 ˚C

1.34  C per s

85 ˚C to 25 ˚C

0.2  C per s

4 ˚C

Hold

Use 100-fold diluted dsDNA templates for assembly of the crRNA array. 3. PCR-based elongation of crRNA array. To construct the crRNA array, design forward and reverse primers that bind to the flanking spacer sequences of the customized dsDNA template. Imbed one to two additional spacer sequences and DR sequences within each primer. Extend the

184

Niels R. Weisbach et al.

dsDNA template with the designed primer pair by PCR using Phusion Flash High-Fidelity PCR Master Mix according to manufacturer’s instructions. To further extend the crRNA array, repeat the PCR-based elongation by using 1 μl PCR reaction mixture as template with a complementary flanking primer pair. Repeat the process until the desired crRNA array composition is achieved. To finalize the crRNA array in preparation for SiT-Cas12a plasmid incorporation, design the final pair of elongation primers that bind to the flanking spacer sequences and encode specific Gibson Assembly overhang sequences from approximately 30 nucleotides, matching the overhang sequences from the digested SiT-Cas12a plasmid backbone sequence (Fig. 4a). 4. Visualization and purification of the crRNA array by gel electrophoresis. To visualize the successful crRNA array synthesis by PCR elongation, load the assembled crRNA array on a 2% E-Gel and extract the corresponding DNA band. Purify the crRNA array with any DNA gel extraction kit (e.g., Qiagen) and elute the DNA in 30 μl Elution Buffer.

Fig. 4 Assembly strategies for massively multiplexed SiT-Cas12a platforms. (a) Schematic representation of the assembly process for medium sized crRNA arrays (up to 10 crRNAs) by PCR-based elongation and subsequent incorporation into the SiT-Cas12a plasmid backbone by Gibson Assembly. The medium sized crRNA array is assembled through PCR-based elongation of a dsDNA fragment-consisting of two spacer sequences separated by one direct repeat sequence-repetitively. Each extension cycle elongates the crRNA array with one or two additional crRNAs on each side depending on the primer design. The final elongation step adds complementary sequences to the SiT-Cas12a plasmid backbone to be used by Gibson Assembly. (b) Schematic representation of the assembly strategy for 10 or more crRNAs into the SiT-Cas12a plasmid backbone by Golden Gate cloning. The strategy is based on utilizing the complementary Type IIS restriction sites and Golden Gate cloning by using a SiT-Cas12a plasmid and additional plasmids encoding medium sized crRNAs arrays (encoding three to six crRNAs). Thus, the crRNA arrays are extracted from the plasmids and simultaneously incorporated into a SiT-Cas12a plasmid backbone

Multiplexed Genome Engineering with Cas12a

185

5. Assembly of SiT-Cas12a plasmid by Gibson Assembly cloning and verification by Sanger sequencing. Clone the crRNA array into the plasmid backbone by using the Gibson Assembly cloning kit. X μl

Vector (50 fmol)

Z μl

crRNA array (150 fmol)

10 μl

2
Gibson Assembly Master Mix

Y μl

H2O

20 μl

Total

Incubate the reaction at 50  C for 60 min and store the mixture at 20  C. Transform 5 μl ligation mixture as detailed in Subheading 3.1 (step 4) and verify the SiT-Cas12a plasmid by Sanger sequencing as detailed in Subheading 3.1 (step 5) using the Triplex forward primer (see Table 2). 3.2.3 Assembly and Incorporation of Large crRNA Arrays into SiT-Cas12a Plasmids

The assembly of large crRNA arrays-harboring 10 or more crRNAs-can be accomplished by Custom Gene Synthesis (e.g., IDT) of several medium sized crRNA arrays provided on separate plasmids and concurrent cloning into SiT-Cas12a plasmids by Golden Gate cloning. 1. Custom gene synthesis of medium-sized crRNA arrays. Design customized gene plasmids containing up to 6 crRNAs which are compatible for Golden Gate cloning into the SiT-Cas12a plasmid. To enable a directed and scarless assembly of a large crRNA array, incorporate truncated DR sequences, flanked by Type IIS restriction sites, on the 50 and 30 end of the crRNA array. The Type IIS restriction sites need to be complementary to each other and to the SiT-Cas12a backbone plasmid (Fig. 4b). 2. Assembly of SiT-Cas12a plasmids by Golden Gate cloning and verification by Sanger sequencing. Clone the crRNA array by Golden Gate cloning using all synthesized customized gene plasmids together with any SiT-Cas12a plasmid (see Table 1). X μl

SiT-Cas12a plasmid (20 fmol)

Z μl

Customized gene plasmid (60 fmol)

1 μl

10
T4 ligase buffer

0.75 μl

BsmBI/SapI (FastDigest)

0.25 μl

T7 ligase

Y μl

H2O

10 μl

Total

186

Niels R. Weisbach et al.

To assemble the full crRNA array and simultaneously incorporate the crRNA array into the SiT-Cas12 plasmid, incubate the reaction mixture using the following program. 37 ˚C

5 min

16 ˚C

5 min
30 cycle

37 ˚C

15 min

55 ˚C

20 min

4 ˚C

Hold

Transform 5 μl of the reaction mixture as detailed in Subheading 3.1 (step 4) and verify the SiT-Cas12a plasmid by Sanger sequencing as detailed in Subheading 3.1 (step 5) using the Triplex forward primer (see Table 2). 3.3 Maintenance and Transfection of HEK 293T Cells

The SiT-Cas12a system can be used for genome engineering in mammalian cells. To demonstrate successful genome editing, transcriptional regulation, or orthogonal engineering, HEK 293T cells were maintained and transfected with various SiT-Cas12a plasmids. 1. Cell culture. Culture HEK 293T cells in antibiotic free Dulbecco’s modified Eagle’s Medium (DMEM), supplemented with 10% FBS and 10 mM HEPES, at 37  C and 5% CO2. Maintain the cells by passaging continuously every 48 hours and keep the confluency below 70%. 2. Seeding cells prior to transfection. At 16–24 h prior to transfection, collect HEK 293T cells by washing with PBS (pH 7.4) and disassociating them with TrypLE. Subsequently, plate 5
104 cells into 24-well plates with 500 μl cultivation medium. Spread the cells by gently shaking and incubate the plate at 37  C and 5% CO2 overnight. 3. Transfection of HEK 293T cells. Depending on the experimental setup, dilute 0.6 to 1 μg of total DNA per well in 50 μl Opti-MEM and mix the solution gently by pipetting. Additionally, dilute 2 μl Lipofectamine 2000 transfection reagent with 48 μl Opti-MEM in a separate tube and mix the solution by pipetting. Incubate both solutions at room temperature for 5 min. Then, transfer the diluted DNA solution into the Lipofectamine solution and shake the mixture gently. To facilitate the formation of DNA–Lipofectamine complexes, incubate the mixture at room temperature for 20 min. Finally, add the DNA–Lipofectamine solution to the

Multiplexed Genome Engineering with Cas12a

187

cells in a dropwise manner and incubate the plate at 37  C and 5% CO2 overnight. Exchange the media after 4–12 h and cultivate the cells for 48–72 h to allow SiT-Cas12a–mediated genome engineering. 3.4 Analysis of SiT-Cas12a– mediated Genome Editing Efficiency

SiT-Cas12a plasmids with incorporated crRNA arrays enable genome editing of mammalian cells in a multiplexed fashion. Genome editing events can be detected and quantified with the Surveyor assay. 1. Extraction of genomic DNA from transfected cells. Transfect 5
104 HEK 293T cells with up to 1 μg of DNA plasmid. 72 h after transfection, wash and collect the transfected cells by trypsinization. Spin the cell suspension for 5 min at 500
g and discard the supernatant. To extract the genomic DNA, resuspend and lyse the cell pellet in QE Buffer (1 mM CaCl2, 3 mM MgCl2, 1% Triton X-100, 10 mM Tris pH 7.4, 1 mM EDTA, 0.2 mg ml1 Proteinase K) in a thermocycler with the following parameters. 65 ˚C

15 min

68 ˚C

15 min

98 ˚C

10 min

4 ˚C

Hold

The extracted genomic DNA can be stored at 20  C or directly used for targeted amplification of the genomic region. 2. Amplification of the targeted genomic region. Use the extracted genomic DNA as a template for PCR-based amplification of the targeted genomic region by using Phusion Flash High-Fidelity PCR Master Mix together with a designed primer pair, which bind at a distance of at least 200 bp from the expected cleavage site and amplify a fragment of less than 1500 bp. Purify the PCR reaction with any DNA purification kit (e.g., Zymo Research) and quantify the concentration by using a Nanodrop UV spectrophotometer. 3. Forming of heteroduplex DNA fragments. The formation of heteroduplex DNA fragments can be achieved by mixing 250 ng purified amplicons from the amplified genomic region with 2 μl of 10x Annealing Buffer (500 mM NaCl, 100 mM MgCl2, 400 mM Tris–HCl, pH 7.9) and dilute the reaction with H2O to a total volume of 20 μl. Incubate the reaction in a thermocycler with the following parameters.

188

Niels R. Weisbach et al. 95 ˚C

10 min

95 ˚C to 85 ˚C

1.34  C per s

85 ˚C to 25 ˚C

0.2  C per s

4 ˚C

Hold

The hybridized heteroduplex DNA fragments can be stored at 20  C or directly used as template for the Surveyor assay. 4. Surveyor Assay. Cleavage events of the hybridized heteroduplex DNA fragments are quantified by using the SURVEYOR® Mutation Detection kit (IDT). In brief, add MgCl2, Surveyor enhancer and Surveyor nuclease according to manufacturer’s instructions and incubate the reaction in a heating block or thermocycler for 60 min at 42  C. The Surveyor mixture can be stored at 20  C or directly used for visualization and quantification of genome editing efficiency. 5. Visualization and quantification of genome editing efficiency. Separate the Surveyor mixture by gel electrophoresis using a 2% E-Gel for quick analysis or by using polyacrylamide gels for a higher resolution of the digested heteroduplex DNA fragments. Visualize the digested heteroduplex DNA fragments using Gel DOC EZ imager and calculate the genome editing efficiency by gel-band intensity through applying the following formula.      b Indel ð%Þ ¼ 1  1 
0:5
100 ða þ b Þ with a as the integrated band intensity of the uncut DNA fragments and b as the sum of the integrated band intensity of the cut heteroduplex DNA fragments. 3.5 Validation of SiT-Cas12a– Mediated Transcriptional Regulatory Efficiency

Multiplexed transcriptional regulation of genes in mammalian cells can be achieved by using SiT-Cas12a plasmids with incorporated transcriptional regulator domains and crRNA arrays. Gene expression levels can be validated by quantitative PCR (qPCR) as described below. 1. RNA Isolation from transfected cells. Transfect 5
104 HEK 293T cells with up to 1 μg of DNA plasmid and cultivate transfected cells for 48 h. Subsequently, wash and collect transfected cells by trypsinization and wash the cell suspension with PBS (pH 7.4). Spin the cell suspension for 5 min at 500
g and discard the supernatant. Extract the RNA with any RNA extraction kit (Quick RNA Miniprep Plus

Multiplexed Genome Engineering with Cas12a

189

kit, Zymo Research) by following the manufacturer’s instructions and quantify the RNA. Purified RNA can be stored at 80  C or directly used as template for cDNA synthesis. 2. Synthesis of cDNA. Transcribe 500 ng purified RNA into cDNA by using qScript cDNA SuperMix. 4 μl

5
qScript cDNA SuperMix

X μl

500 ng RNA

Y μl

H2O

20 μl

Total

Incubate the cDNA reaction mixture in a thermocycler with the following program. 25 ˚C

5 min

42 ˚C

30 min

85 ˚C

5 min

4 ˚C

Hold

cDNA samples can be stored at 20  C or directly used as template for qPCR. 3. Quantification of RNA expression by qPCR. Perform qPCR on the cDNA template by using Fast Plus EvaGreen® qPCR Master Mix with target specific qPCR primers according to manufacturer’s instructions. Normalize the RNA expression of the target gene based on the RNA expression of a control gene for HEK 293T cells, such as glyceraldehyde 3-phosphate dehydrogenase (GAPDH), and quantify the gene expression with the comparative CT method [17].

4

Notes 1. Identification of CRISPR-Cas12a targeting sites Efficient Cas12a-mediated genome engineering requires the identification of optimal genomic target sites. The selection requirements are: (1) selection of the target genomic locus for genome editing (see Note 2) or transcriptional regulation (see Note 3), (2) presence of protospacer adjacent motif (PAM) sequences compatible with AsCas12a (“TTTV”), and (3) identification of the target sequence downstream of the PAM. Optimal crRNAs can be predicted in silico by using Cas12aspecific tools, such as CINDEL [18], or more advanced deep learning tools, such as DeepCpf1 [19]. To optimize the

190

Niels R. Weisbach et al.

efficiency, especially in a multiplexed setup, it is necessary to validate various spacers in the chosen experimental background before combining the most effective spacers within a single crRNA array. 2. Choosing crRNA targeting sites for gene editing Genome editing by SiT-Cas12a systems require a DNaseactive Cas12a variant (see Table 1) and a preselected crRNA guide. In addition to the mentioned requirements above (see Note 1), crRNAs for optimal genome editing require a spacer sequence length from 20 to 23 nt to enable Cas12a-mediated dsDNA cleavage of a specific genomic target. Cas12a-mediated DNA cleavage in the genome of mammalian cells results in the activation of the nonhomologous end joining (NHEJ) pathway, which not only repairs the cleaved DNA but also can induce insertions or deletions (indels) of single or multiple base pairs. Consequently, indel formation can disrupt gene function through frameshifts in the coding frame of a gene. For editing a gene of interest, identify the target spacer sequences within the coding region and validate the spacers with the Surveyor assay (see Subheading 3.4). To enhance gene editing efficiency, utilize several crRNAs targeting the same gene within one crRNA array to achieve synergistic effects. Nevertheless, an increased number of DNA cleavage events within mammalian cells can lead to decreased fitness, increased number of chromosomal rearrangement events, and potentially cell death. Hence, it is necessary to balance the possible enhanced gene editing effect against the possible drawbacks. 3. Choosing crRNA targeting sites for gene regulation The SiT-Cas12a system enables orthogonal gene editing and transcriptional regulation with the DNase-active Cas12a variant fused with a transcriptional regulation domain (see Table 1). Besides the previously mentioned requirements for crRNA design (see Note 1), crRNAs for transcriptional regulation in an orthogonal SiT-Cas12a system require a defined spacer sequence length of 15 nt to enable target binding without Cas12a-mediated DNA cleavage. To facilitate transcriptional regulation of target genes, crRNAs should target the promoter region proximal to the transcriptional start site (TSS). We recommend validating the crRNAs individually by using a qPCR-based method (see Subheading 3.5) before incorporation into crRNA arrays. 4. Design and order of dsDNA fragments or synthesized plasmids To leverage the full potential of the SiT-Cas12a platform, it is necessary to assemble and incorporate crRNA arrays of certain sizes. Through modern DNA synthesis technologies, the production of linear and circular DNA fragments can be realized on an industrial scale by various suppliers and enable the

Multiplexed Genome Engineering with Cas12a

191

crRNA array synthesis to a certain level. However, the quantity and the quality among the suppliers varies and the current limitation of synthesis lies in the complexity of long crRNA array sequences, especially with regard to the number of repetitive DR sequences within a single crRNA array. Facing this limitation, the synthesis of linear crRNA arrays is currently restricted to two DR sequences (gBlock synthesis by IDT) and the synthesis of crRNA arrays within plasmids to five DR sequences (Gene synthesis by IDT). The exact number of DRs that can be incorporated in a single DNA fragment will vary between suppliers. Therefore, depending on the number of targets, choose the strategy-explained in this chapter-that facilitates crRNA array synthesis toward the desired number of genomic targets. References 1. Hsu PD, Lander ES, Zhang F (2014) Development and applications of CRISPR-Cas9 for genome engineering. Cell 157:1262–1278. https://doi.org/10.1016/j.cell.2014.05.010 2. Doudna JA, Charpentier E (2014) The new frontier of genome engineering with CRISPR-Cas9. Science 346:1258096. https://doi.org/10.1126/science.1258096 3. Anzalone AV, Koblan LW, Liu DR (2020) Genome editing with CRISPR–Cas nucleases, base editors, transposases and prime editors. Nat Biotechnol 38:824–844. https://doi. org/10.1038/s41587-020-0561-9 4. Steinmetz LM, Davis RW (2004) Maximizing the potential of functional genomics. Nat Rev Genet 5:190–201. https://doi.org/10.1038/ nrg1293 5. Nissim L, Perli SD, Fridkin A et al (2014) Multiplexed and programmable regulation of gene networks with an integrated RNA and CRISPR/Cas toolkit in human cells. Mol Cell 54:698–710. https://doi.org/10.1016/j. molcel.2014.04.022 6. Campa CC, Weisbach NR, Santinha AJ et al (2019) Multiplexed genome engineering by Cas12a and CRISPR arrays encoded on single transcripts. Nat Methods 16:887–893. https:// doi.org/10.1038/s41592-019-0508-6 7. Haurwitz RE, Jinek M, Wiedenheft B et al (2010) Sequence- and structure-specific RNA processing by a CRISPR endonuclease. Science 329:1355–1358. https://doi.org/10.1126/ science.1192272 8. Cong L, Ran FA, Cox D et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339:819–823. https://doi.org/ 10.1126/science.1231143

9. Mali P, Yang L, Esvelt KM et al (2013) RNA-guided human genome engineering via Cas9. Science 339:823–826. https://doi.org/ 10.1126/science.1232033 10. Jinek M, East A, Cheng A et al (2013) RNA-programmed genome editing in human cells. eLife 2:1–9. https://doi.org/10.7554/ eLife.00471 11. McCarty NS, Graham AE, Studena´ L, Ledesma-Amaro R (2020) Multiplexed CRISPR technologies for gene editing and transcriptional regulation. Nat Commun 11:1281. https://doi.org/10.1038/s41467020-15053-x 12. Makarova KS, Wolf YI, Iranzo J et al (2020) Evolutionary classification of CRISPR–Cas systems: a burst of class 2 and derived variants. Nat Rev Microbiol 18:67–83. https://doi. org/10.1038/s41579-019-0299-x 13. Zetsche B, Heidenreich M, Mohanraju P et al (2017) Multiplex gene editing by CRISPR–Cpf1 using a single crRNA array. Nat Biotechnol 35:31–34. https://doi.org/ 10.1038/nbt.3737 14. Adiego-Pe´rez B, Randazzo P, Daran JM et al (2019) Multiplex genome editing of microorganisms using CRISPR-Cas. FEMS Microbiol Lett 366:1–19. https://doi.org/10.1093/ femsle/fnz086 15. Zhao Y, Boeke JD (2020) CRISPR–Cas12a system in fission yeast for multiplex genomic editing and CRISPR interference. Nucleic Acids Res 48:5788–5798. https://doi.org/ 10.1093/nar/gkaa329 16. Fonfara I, Richter H, Bratovicˇ M et al (2016) The CRISPR-associated DNA-cleaving enzyme Cpf1 also processes precursor CRISPR

192

Niels R. Weisbach et al.

RNA. Nature 532:517–521. https://doi.org/ 10.1038/nature17945 17. Schmittgen TD, Livak KJ (2008) Analyzing real-time PCR data by the comparative CT method. Nat Protoc 3:1101–1108. https:// doi.org/10.1038/nprot.2008.73 18. Kim HK, Song M, Lee J et al (2017) In vivo high-throughput profiling of CRISPR–Cpf1

activity. Nat Methods 14:153–159. https:// doi.org/10.1038/nmeth.4104 19. Kim HK, Min S, Song M et al (2018) Deep learning improves prediction of CRISPR–Cpf1 guide RNA activity. Nat Biotechnol 36:239–241. https://doi.org/10.1038/nbt. 4061

Chapter 12 Highly Multiplexed Analysis of CRISPR Genome Editing Outcomes in Mammalian Cells Soh Ishiguro and Nozomu Yachie Abstract CRISPR–Cas-based genome editing has enabled efficient genetic engineering of a range of organisms and sparked revolutions in many fields of biology. After Streptococcus pyogenes Cas9 was first demonstrated for mammalian genome editing, many CRISPR-associated (Cas) protein variants have been isolated from different species and adopted for genome editing. Furthermore, various effector domains have been fused to these Cas proteins to expand their genome-editing abilities. Although the number of genomeediting tools has been rapidly increasing, the throughput of cell-based characterization of new genomeediting tools remains limited. Here we describe a highly multiplexed genome editing and sequencing library preparation protocol that allows high-resolution analysis of mutation outcomes and frequencies induced by hundreds to thousands of different genome-editing reagents in mammalian cells. We have successful experiences of developing several key genome-editing tools using this protocol. The protocol also is designed to be compatible with robotic liquid handling systems for further scalability. Key words Genome editing, CRISPR–Cas9, Base editing, High-throughput sequencing, Amplicon sequencing

1

Introduction A number of CRISPR genome-editing systems have been developed. Streptococcus pyogenes Cas9 (SpCas9) was the first to be demonstrated for mammalian genome editing [1, 2] and it has been the most widely used in a range of fields, including genome-wide gene knockout screens and development of genome editing-based therapeutic approaches [3]. In CRISPR–Cas9 genome editing, a chimeric single guide RNA (gRNA) recruits Cas9 to the target genomic sequence that is adjacent to a 30 protospacer adjacent motif (PAM) through sequence complementarity (Fig. 1). The two endonuclease domains of Cas9 then induce a double-stranded DNA break (DSB) at the target genomic region. This has been used to induce either mutations (mostly deletions) in target genes through error-prone nonhomologous end-joining

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

193

194

Soh Ishiguro and Nozomu Yachie

Cas9 M PA

3 5 3

3 5

gRNA

5

Target-AID H2 O

C

ABEmax H 2O

NH3

H 2O

NH3

A

U

C

I

nCas9 1

A 3 5

5

gRNA

5 3

nCas9 1

3

A CD 3 5

Ta dA 5

gRNA

5 3

C A

3

M PA

3

I

UGI

M PA

C

NH3

A

U

nCas9

M PA

5 3

H 2O

NH3

UGI

UGI A CD

Target-ACEmax

3 5

Ta dA 5

gRNA

Fig. 1 CRISPR–Cas9 genome editing and base editing. Conceptual diagrams of genome editing by wild-type Cas9 and three base editors. The base editing tools, Target-AID [11], ABEmax [17], and Target-ACEmax [35], convert C·G!T·A, A·T!G·C, and simultaneously C·G!T·A and A·T!G·C, respectively

DNA repair or transgene insertions through homology-directed DNA repair in eukaryotic cells. While wild-type SpCas9 has been adopted in diverse applications, many protein engineering efforts have further enhanced its genome-editing ability. For example, recent studies have expanded the targeting scope of SpCas9 by relaxing its wild-type PAM restriction (50 -NGG-30 or 50 -NGA-30 ). Such PAM-altered Cas9 variants include SpCas9-NG [4] and xCas9-3.7 [5] for 50 -NG-30 PAM, and near PAM-less SpG and SpRY [6]. Other efforts have focused on minimizing undesired off-target edits that usually occur through mismatch-tolerant annealing of gRNA to nontargeting genomic regions, and several high-fidelity Cas9 variants, such as SpCas9-HF1 [7], eCas9(1.1) [8], and evoCas9 [9], have been derived. The DSB-based genome editing by wild-type SpCas9 is particularly effective in gene knockout, but has a drawback in precision genome editing, where mutation outcomes are not always predictable. Induction of a single-base substitution has been proposed with the transgene insertion approach to replace target sequence with a donor template sequence by homology-directed DNA repair, but its efficiency is limited. Furthermore, Cas9-derived DSBs are cytotoxic in cells [10, 11]. Harnessing a nucleoside deaminase domain to catalytically inactive Cas9 (dCas9) or nickase Cas9 (nCas9), base editing technologies have enabled targeted

High-Throughput Analysis of CRISPR Genome Editing Outcomes

195

nucleotide substitutions to be induced without DSB [12]. The current base editors are categorized as cytosine base editors (CBEs), for targeted C·G!T·A substitution, and adenine base editors (ABEs), for targeted A·T!G·C substitution (Fig. 1). The CBEs use a cytidine deaminase such as rAPOBEC1 or PmCDA1 to convert cytidines into uridines in the nontargeting DNA strand of gRNA and a uracil glycosylase inhibitor to prevent base excision repair of uracil bases [11, 13]. Following DNA replication, uracils are converted into thymines. The ABEs instead use an evolved tRNA adenosine deaminase (TadA) that converts adenosines to inosines on single-stranded DNA (ssDNA). Although none of the natural adenosine deaminases have been found to have catalytic activity for ssDNA, a directed protein evolution approach derived the evolved TadA heterodimer for A·T!G·C base editing [14]. The CBEs and ABEs both have narrow base editing windows within the gRNA target site and are suited for targeted sequence alternation. The idea of base editing has been rapidly applied to the engineered SpCas9 variants to expand their targeting scopes [15] and enhance their targeting fidelities [16]. Furthermore, several studies have reported that codon optimization, linker optimization, and other accessory domains, such as nuclear localization signals and ssDNA-binding domains, can be used to derive more efficient base editors if they are fused in an optimal order [17– 20]. Non-specific C!U and A!I edits of the cellular transcriptome have been a concern for CBEs using rAPOBEC1 and ABEs using TadA, the original wild-types of which have catalytic activity for RNA (this has been reported not to be the case for PmCDA1based CBEs). However, protein engineering approaches also have derived deaminase variants that minimize such nonspecific RNA off-target activities [21–24]. Although the delivery efficiency of genome-editing reagents to cells and nuclei is important, SpCas9 is 1358 aa long, which limits the delivery efficiency of SpCas9 to cells by, for example, adenoassociated viral vectors. To resolve this issue, other compact Cas9 orthologues have been explored and reported for genome editing, such as SaCas9 from Staphylococcus aureus (1053 aa) [25] and CjCas9 from Campylobacter jejuni (984 aa) [26]. These Cas9 orthologues also have been fused to deaminase domains to obtain compact base editing tools [27, 28]. Furthermore, other CRISPR– Cas systems from outside the CRISPR–Cas9 family have been characterized for efficient genome editing [29]. For example, LbCas12a from Lachnospiraceae bacterium (formally characterized as LbCpf1) catalyzes site-specific cleavage of DNA with T-rich 50 -TTTV-30 PAM adjacent to the 50 -end of the gRNA-targeting sequence [30]. rAPOBEC1 and TadA also have been fused to catalytically inactive LbCas12a for base editing [31].

196

Soh Ishiguro and Nozomu Yachie

Accordingly, a range of Cas proteins, mutations, and different effector and linker domains have been reported for genome editing and expanding the design space to develop new genome-editing tools. However, the number of possible design combinations to derive a new tool has encountered a combinatorial explosion, making it difficult to derive an optimal-best tool for a specific target of application because the editing outcome patterns of each tool need to be thoroughly evaluated for a range of genomic on-target and predicted off-target sites in cells. Recently, along with three other similar studies [32–34], we have developed a novel dualfunction base editor Target-ACEmax for the simultaneous introduction of C·G!T·A and A·T!G·C substitutions by fusing both cytidine and adenosine deaminases to nCas9 [35]. We tested genome-editing outcome patterns of 47 endogenous target sites for three dual-function base editor candidates, six single-function base editors, and four single-function base editor mixes, as well as a non-functional enzyme control in triplicates by amplicon sequencing (a total of 1833 assays). In this methodology article, we describe a modified protocol that enables genome-editing outcome assays of 48 different gRNA target sites in human embryonic kidney 293 T (HEK293T) cells for genome-editing tools as well as a control enzyme in triplicate (Fig. 2). The protocol has five sections: (1) screening of gRNA target sites and target amplification primers, (2) preparation of a gRNA expression plasmid library, (3) culturing of cells, (4) massively parallel transfection of genome-editing reagents, (5) template genomic DNA preparation, and (6) generation of a multiplexed amplicon sequencing library. This protocol produces a single multiplexed amplicon sequencing library for hundreds to thousands of genome-editing assay samples and enables analysis of their genome-editing outcomes by one-shot massively parallel sequencing (Fig. 3). This basic protocol can easily be scaled up with increased numbers of gRNA on-target and predicted off-target sites. It can also be modified for different cell lines with optimal cell culture and transfection protocols. (We have successful experiences of performing similar genome-editing assays for HeLa, HCC827, and mouse embryonic stem cells with similar library-scale protocols.) Furthermore, we designed many of the process modules in the protocol to be compatible with robotic liquid handling systems for more systematic characterization of a large number of genome-editing tools. We believe that this step-bystep protocol will be of great use for researchers to facilitate the rapid development of new genome-editing tools in mammalian cells.

High-Throughput Analysis of CRISPR Genome Editing Outcomes

197

Fig. 2 Overview of the highly multiplexed amplicon sequencing protocol to analyze CRISPR genome editing outcomes and frequencies. First, the genome-editing enzyme plasmids and gRNA plasmids are assembled to produce all possible combinatorial transfection reagents. Then, highly parallel genome-editing assays are performed in triplicates by cell transfection. Next, cell samples are lysed and direct PCR is used to amplify the gRNA target regions with primers that have common adapter sequences (PS1.0 and PS2.0). The first-round PCR products are then pooled for each genome-editing enzyme of each replicate and the second-round PCR is performed to attach custom Illumina index adapters. After sample normalization, the amplicon sequencing libraries are pooled into a single multiplexed library and subjected to an Illumina sequencing

2

Materials

2.1 General Considerations

All the experimental steps described here use PCR-grade ultrapure distilled water (not autoclaved distilled water) to minimize the potential risks of nucleic acid and DNase/RNase contaminations that may affect the genome-editing assays and the quality of the sequencing libraries. We also recommend performing all the experiments using filtered tips on a clean bench. In the following sections, we provide a basic protocol to measure mutation outcomes of given genome-editing tools and a nonfunctional control enzyme (EGFP) condition for 48 different endogenous gRNA targeting sites of HEK293T cells in triplicates (hundreds to thousands of genomeediting assays). However, this protocol can be easily scaled up for an increased number of gRNAs and target sites, and the cell line also can be replaced with others with their own cell culture and transfection protocols.

Outcome patterns

Soh Ishiguro and Nozomu Yachie

EMX1

Outcome patterns

NEAT1

198

A T G C

Target

PAM

Fig. 3 Data analyses of two arbitrary selected genome-editing assays in triplicates multiplexed with a total of nearly two thousand assays. Base editing of NEAT1 and EMX1 encoding regions by Target-ACEmax 2.2 Screening of gRNA Target Sites and Target Amplification Primers

1. DNase/RNase-free ultrapure distilled water. 2. 8-strip individually capped PCR tubes. 3. Hard-Shell® 96-Well PCR Plates, low profile, thin wall, skirted, white/clear (Bio-Rad HSP9601). 4. Microseal® ‘C’ PCR Plate Sealing Film (Bio-Rad MSC1001). 5. 5 Phusion HF Buffer (NEB B0518S). 6. Phusion DNA Polymerase (NEB M0530S). 7. dNTPs (25 mM each). 8. 1/100 diluted SYBR Green: mix 10 μL of SYBR™ Green I Nucleic Acid Gel Stain (Invitrogen S7563) and 990 μL of 100% dimethyl sulfoxide. Wrap with aluminum foil to protect from light and store at 4  C. 9. 6 loading dye. 10. Target amplification of forward primers with PS1.0 adapter sequence: 50 -TAACTTACGGAGTCGCTCTACG XXXXXXX XXXXXXXXXXXXX -30 (Xs: annealing sequence of forward primer for target amplification). 11. Target amplification of reverse primers with PS2.0 adapter sequence: 50 -GGATGGGATTCTTTAGGTCCTG XXXXXXX XXXXXXXXXXXXX -30 (Xs: annealing sequence of reverse primer for target amplification).

High-Throughput Analysis of CRISPR Genome Editing Outcomes

2.3 Preparation of gRNA Expression Plasmid Library

199

1. DNase/RNase-free ultrapure distilled water. 2. DH5-alpha Competent E. coli. 3. Super Optimal broth with Catabolite repression (SOC) outgrowth medium. 4. T4 DNA Ligase (NEB). 5. 10 T4 Ligase Buffer with 10 mM ATP (NEB). 6. T4 Polynucleotide Kinase (NEB). 7. BsmBI (NEB R0580S). 8. 1 mg/mL BSA: mix 10 μL of 10 mg/mL BSA and 90 μL of ultrapure distilled water. Store at 20  C. 9. gRNA cloning backbone plasmid (pSI-356 v2; available at Addgene https://www.addgene.org/158431/) (see Note 1). 10. DNA miniprep kit. 11. 100-mL disposable DNase/RNase-free reagent reservoirs. 12. 96-well DNase/RNase-free 2-mL round bottom deep-well plates. 13. 96-well DNase/RNase-free PCR plates. 14. Microseal® ‘C’ PCR Plate Sealing Films (Bio-Rad MSC1001) or any alternative that can perfectly seal up the wells. 15. Lysogeny broth (LB) medium: add 25 g of LB powder to distilled water, top up to 1 L, autoclave and store at 4  C. 16. LB-Ampicillin medium: add 100 μL of 100 mg/mL ampicillin to 100 mL of LB medium. 17. LB-Ampicillin agar plate: add 15 g of agar, and 25 g of LB powder to distilled water in a glass bottle, top up to 1 L, autoclave, cool down to 55  C, then add 1000 μL of 100 mg/mL ampicillin, and pour to bacterial petri dishes. 18. 15-mL bacterial culture tubes. 19. Sanger sequencing primer to validate gRNA spacer sequences. Use 50 - TTTCCCATGATTCCTTCATATTT -30 for the gRNA cloning backbone plasmid above (compatible with any human U6 promoter-gRNA expression plasmid).

2.4

Culturing of Cells

1. Human embryonic kidney cells 293T (HEK293T cells). 2. 0.05 w/v% Trypsin-0.53 mmol/L EDTA-4Na Solution with Phenol Red. 3. Heat-inactivated fetal bovine serum (FBS): thaw FBS (Invitrogen 10437-028) at 4  C overnight, incubate for 30 min in 56  C water bath, and aliquot to 50-mL tubes. Store at 20  C.

200

Soh Ishiguro and Nozomu Yachie

4. 1 Phosphate-Buffered Saline (PBS): mix 100 mL of 10 D-PBS() and 900 mL of DNase/RNase-free ultrapure distilled water in a 1 L glass bottle. Store at 4  C. 5. 100 penicillin–streptomycin. 6. Cell culture medium: mix 500 mL of DMEM, 50 mL of heatinactivated FBS, and 5 mL of 100 penicillin–streptomycin. Store at 4  C. 7. 5 mM Acetic Acid (AcOH): add 500 μL of 1 M AcOH to 99.50 mL of DNase/RNase-free ultrapure distilled water in 200 mL glass bottle and autoclave. Store at 4  C. 8. Collagen-I solution: mix 40 μL of Collagen Type I and 11.96 mL of 5 mM AcOH in a 15-mL tube. Store at 4  C. 9. 96-well cell culture plates. 10. 10-cm cell culture dishes. 2.5 Massively Parallel Transfection of Genome-Editing Reagents

1. Genome-editing enzyme expression plasmids. For example, PpX165 is a wild-type SpCas9 nuclease expression plasmid for mammalian genome editing (Addgene 48137). A range of base editor plasmids also can be obtained from Addgene (https:// www.addgene.org/crispr/base-edit/). 2. EGFP expression plasmid as non-functional enzyme control (pLV-eGFP, Addgene 36083, or any alternative). 3. Plasmid DNA midiprep Kit. 4. Plasmid DNA miniprep Kit. 5. 96-well DNase/RNase-free 1.1 mL round-bottom deep well plates. 6. Microseal® ‘C’ PCR Plate Sealing Film (Bio-Rad MSC1001). 7. 1 mg/mL Polyethylenimine (PEI) MAX solution: dissolve 10 mg PEI MAX (Polysciences 49553-93-7) in 10 mL of ultrapure distilled water, filter through a 0.22-μm filter, and aliquot 1000 μL into each 1.5-mL microcentrifuge tube with screw cap. Store at 20  C for up to 12 months.

2.6 Template Genomic DNA Preparation

1. 50 mM NaOH: add 500 μL of 5 N NaOH to 49.5 mL of DNase/RNase-free ultrapure distilled water in a 50-mL tube. Store at 4  C (see Note 2). 2. 1 M Tris–HCI, pH 8.0. 3. Hard-Shell® 96-Well PCR Plates, low profile, thin wall, skirted, white/clear (Bio-Rad HSP9601). 4. Microseal® ‘C’ PCR Plate Sealing Film (Bio-Rad MSC1001).

High-Throughput Analysis of CRISPR Genome Editing Outcomes

2.7 Generation of a Multiplexed Amplicon Sequencing Library and Sequencing

201

1. DNase/RNase-free ultrapure distilled water. 2. 8-strip individually capped PCR tubes. 3. Hard-Shell® 96-Well PCR Plates, low profile, thin wall, skirted, white/clear (Bio-Rad HSP9601). 4. Microseal® ‘C’ PCR Plate Sealing Film (Bio-Rad MSC1001). 5. 5 Phusion HF Buffer (NEB B0518S). 6. Phusion DNA Polymerase (NEB M0530S). 7. dNTPs (25 mM each). 8. 1/100 diluted SYBR Green: mix 10 μL of SYBR™ Green I Nucleic Acid Gel Stain (Invitrogen S7563) and 990 μL of 100% dimethyl sulfoxide. Wrap with aluminum foil to protect from light and store at 4  C. 9. 6 loading dye. 10. Agencourt AMPure XP (Beckman Coulter A63881). 11. 1.5 mL DNA LoBind tubes (Eppendorf 0030122348). 12. 70% Ethanol: add 30 mL of 100% Ethanol to 70 mL of ultrapure distilled water. Store at 4  C. 13. 1 Tris-EDTA (TE) buffer. 14. KAPA Library Quantification Kits (Illumina) (KAPA Biosystems KK4824, or any alternative). 15. Illumina MiSeq v3 600 cycles kit (Illumina MS-102-3003) or Illumina MiSeq v2 Micro 300 cycles kit (Illumina MS-103-1002). 16. Illumina PhiX control v3 (Illumina FC-110-3001). 17. Target amplification forward primers with PS1.0 adapter sequence: 50 -TAACTTACGGAGTCGCTCTACG XXXXXXX XXXXXXXXXXXXX -30 (Xs: annealing sequence of forward primer for target amplification). 18. Target amplification reverse primers with PS2.0 adapter sequence: 50 -GGATGGGATTCTTTAGGTCCTG XXXXXXX XXXXXXXXXXXXX -30 (Xs: annealing sequence of reverse primer for target amplification). 19. Custom Illumina P5 index primer: 50 -AATGATACGGCGAC CACCGAGATCTACACTCTTTCCCTACAC GACGCTCTTCCGATCT NNNNN YYYYYYYYY T AACTTACGGAGTCGCTCTACG-30 (Ns: random sequence for better flow cell clustering; Ys: 9-bp custom index). 20. Custom Illumina P7 index primer: 50 -CAAGCAGAAGACGG CATACGAGATCGGTCTCGGCATTCCTGCT GAACCGCTCTTCCGATCT NNNNN YYYYYYYYY G GATGGGATTCTTTAGGTCCTG-30 (Ns: random sequence for better flow cell clustering; Ys: 9-bp custom index).

202

2.8

Soh Ishiguro and Nozomu Yachie

List of Equipment

1. Micropipettes (0.2–2 μL, 2–20 μL, 20–200 μL, and 100–1000 μL). 2. 12-channel micropipettes 20–200 μL).

(0.2–2

μL,

2–20

μL,

and

3. 5–50-mL pipete. 4. Bench-top centrifuge. 5. Centrifuge with 96-well plate and 50-mL tube rotors. 6. Heat block. 7. 96-well aluminum block. 8. 1.5-mL tube aluminum block. 9. Thermal cycler machine. 10. Agarose gel imager. 11. Agarose gel electrophoresis bath. 12. Quantitative PCR machine. 13. Vortex mixer. 14. Bacterial incubator. 15. Shaking incubator. 16. Human cell culture incubator (including CO2 controller and aspirator). 17. Cell culture hoods (bacterial and human cell culture). 18. Automated cell counter. 19. Water baths (37  C for human cells and 42  C for bacterial transformation). 20. Plate seal roller. 21. Spectrophotometer. 22. Magnet stand (1.5-mL tube or 96-well plate format). 23. Illumina sequencer.

3

Methods

3.1 Screening of gRNA Target Sites and Amplification Primers

Because target genomic regions need to be stably amplified by PCR and the following genome editing assay procedures need to align with such target regions, we first screen two or three primer pairs for each candidate gRNA target region (Fig. 4), and discard candidate gRNAs with no stable PCR amplification with any of the tested primer pairs. To secure a certain number of gRNAs to be used in the genome editing assays, primer pairs for an excess number of gRNA target sites can be screened in this step. The PCR efficiencies and expected band sizes of primer pairs are evaluated by qPCR and agarose gel electrophoresis. This section describes screening of three primer pairs for 48 genomic gRNA target sites (a total of 144 primer pairs).

High-Throughput Analysis of CRISPR Genome Editing Outcomes

203

Fig. 4 Example of PCR primer screening for stable amplification of gRNA target regions. PCR products with the expected band sizes are marked with white dots 3.1.1 Primer Preparation

1. Design three PCR primer pairs for each of 48 selected gRNA targeting regions and order DNA oligos (144 forward and 144 reverse primer oligos). See Subheading 2.2 for the adapter sequences added to the target amplification primers. Using Primer3 or any alternative, design primer pairs each to have a melting temperature of 55–65  C without homopolymer or palindrome sequences and to produce a PCR product of 150–230 bp for the pooled library preparation (see Subheading 3.6.1 for details). 2. Mix 5 μL of 100 μM forward primer and 5 μL of 100 μM reverse primer to prepare a 50 μM primer mix in each well of fresh 96-well plates using a 12-channel pipette. 3. Prepare 10 mL ultrapure distilled water in a disposable regent reservoir. 4. Transfer 40 μL of ultrapure distilled water from the reservoir to each sample well of the primer mix plates using a 12-channel pipette to obtain 10 μM primer mixes. 5. Seal the primer mix plates with optically clear adhesive PCR films (we recommend the Bio-Rad films described in Subheading 3 throughout the study for good sealability). The primer mix plates can be stored at 20  C.

3.1.2 Template Genomic DNA Preparation

1. Place a DMEM bottle in a 37  C water bath to warm up. 2. Thaw one vial of HEK293T cells in a 37  C water bath and transfer the cells into 5 mL of the prewarmed DMEM in a 15-mL sample tube and centrifuge the cells at 200  g for 5 min. 3. Aspirate and remove the supernatant. 4. Add 10 mL of the prewarmed DMEM into the sample tube. 5. Gently mix the cells by pipetting several times.

204

Soh Ishiguro and Nozomu Yachie

6. Transfer the cells into a 10-cm cell culture dish. 7. Incubate the cell culture dish for 2–3 days in a cell culture incubator at 37  C and 0.05% CO2 to allow cell proliferation to 80–90% confluency (see Note 3). 8. Remove the media using an aspirator and apply 1000 μL of 1 PBS to the cell culture dish. 9. Remove the reagent using an aspirator and apply 1000 μL of 0.05% trypsin-EDTA to the cell culture dish. 10. Incubate the cell culture dish for 4 min in a cell culture incubator at 37  C and 0.05% CO2. 11. Remove the reagent using an aspirator and add 1000 μL of 50 mM NaOH to the cell culture dish for cell lysis. 12. Transfer the sample to a 2-mL tube and incubate at 95  C for 15 min on a heat block, place on ice for 5 min, and neutralize the sample with 100 μL of 1 M Tris–HCl (pH 8.0). This cell lysate sample can be stored at 20  C. 3.1.3 Primer Screening by qPCR

1. To perform the following quantitative PCR (qPCR) for each of the 144 primer pairs, prepare 180 qPCR master mix in a 5 mL sample tube (see Note 4): 1 reaction unit Component

Final concentration

Volume (μL)

5 Phusion HF buffer

1

4

2 mM dNTPs

200 μM

2

10 μM primer mix

500 nM each

2

DNA template (lysed cell sample)

2

Phusion DNA polymerase

0.2

1/100 diluted SYBR green

0.06

Ultrapure distilled water

9.74

Total

20

180 qPCR master mix Component

Volume (μL)

5 Phusion HF buffer

720

2 mM dNTPs

360

DNA template (lysed cell sample)

360

Phusion DNA polymerase

36 (continued)

High-Throughput Analysis of CRISPR Genome Editing Outcomes

205

Component

Volume (μL)

1/100 diluted SYBR green

10.8

UltraPure distilled water

1753.2

Total

3240

2. Transfer the qPCR master mix into a disposable reagent reservoir placed on ice. 3. Using a 12-channel pipette, transfer 18 μL of the qPCR master mix from the reservoir to each well of fresh 96-well PCR plates. 4. Using a 12-channel pipette, add candidate primer pair mixes, 2 μL each, to the qPCR reaction plates. 5. Seal the qPCR reaction plates with optically clear adhesive PCR films. 6. Centrifuge the qPCR reaction plates at 1000  g for 2 min to spin down and remove air bubbles. 7. Perform qPCR with the following thermal cycle conditions: Step

Temperature

Time

1

98  C

0’30

2



0’10



98 C

3

60 C

0’10

4

72  C

1’00

Go to 2 6 7

(29 times) 

72 C 

4 C

5’00 1

8. Save the qPCR results with Ct values for the primer pairs. 9. Centrifuge the qPCR reaction plates at 1000  g for 2 min and gently remove the plate seals. 10. Prepare 6 loading dye in a disposable reagent reservoir and transfer 4 μL to each sample well of the qPCR reaction plate using a 12-channel pipette and mix well. 11. Run a gel for the PCR products, 5 μL each, using 2% agarose gel for 20 min with 135 V. 12. Take gel images and evaluate PCR band sizes and amplification efficiencies with Ct values. 13. Select the best primer pair for each of the gRNA target sites and rearray the curated 10 μM primer pair mixes into a fresh 96-well PCR plate.

206

Soh Ishiguro and Nozomu Yachie

14. Seal the curated primer mix plate with optically clear adhesive PCR films. 15. Centrifuge the curated primer mix plate at 1000  g for 2 min and store at 20  C. 3.2 Preparation of gRNA Expression Plasmid Library

After determining the gRNA target sites and their corresponding primers, gRNA expression plasmids are prepared according to the following procedure. Two ssDNA oligos encoding the gRNA spacer region are prepared for each target site, annealed, and phosphorylated (Fig. 5a). The dsDNA insert is then ligated into a digested plasmid backbone. The assembled DNA is transformed into bacterial cells, and the plasmid clones are obtained by colony isolation (Fig. 5b), followed by miniprep and validated by Sanger sequencing. This gRNA library preparation protocol allows fast preparation of 48–96 gRNA expression plasmids within a week. gRNA libraries can be prepared for other CRISPR–Cas systems by changing the dsDNA insert design and plasmid backbone.

3.2.1 Preparation of gRNA Spacer Inserts

1. Spin down forward and reverse ssDNA oligo plates (100 μM each) (see Note 5). The ssDNA pairs need to be designed to produce sticky 50 (CACC), and 30 (AAAC) ends after annealing. 2. Add 10 μL of forward oligo and 10 μL of reverse oligo into a fresh 96-well PCR plate using a 12-channel pipette. The ssDNA mix plate can be stored at 20  C. 3. To perform the following annealing and phosphorylation reactions for 48 samples, prepare 50 reaction master mix. 1 reaction unit Component

Volume (μL)

ssDNA oligo mix (50 μM each)

2.0

10 T4 ligase buffer with 10 mM ATP

0.5

T4 polynucleotide kinase

0.25

Ultrapure distilled water

2.25

Total

5.0

50 reaction mix Component

Volume (μL)

10 T4 ligase buffer with 10 mM ATP

25

T4 polynucleotide kinase

12.5

Ultrapure distilled water

112.5

Total

150

High-Throughput Analysis of CRISPR Genome Editing Outcomes

207

Fig. 5 Cloning of gRNA expression plasmids. (a) A dsDNA insert encoding a gRNA spacer is prepared by annealing of ssDNA oligos and cloned to a plasmid backbone. (b) Colony isolation to obtain clones of the assembled gRNA products

4. Aliquot 3 μL of the reaction mix into each well of a fresh 96-well PCR plate. 5. Add the ssDNA mixes, 2 μL each, to the reaction plate and assemble with the reaction mix. 6. Seal the reaction plate with an optically clear adhesive PCR film. 7. Centrifuge the reaction plates at 1000  g for 30 s. 8. Incubate the reaction plate in a thermal cycler with the following conditions. Step

Temperature

Time

1

37  C

30’00



2

95 C

5’00

3

95  C, 1  C/cycle

0’12

Go to 3 4

(69 times) 

25 C

1

9. Prepare 10 mL ultrapure distilled water in a disposable reagent reservoir. 10. Remove the optically clear adhesive PCR films. 11. Add 95 μL of ultrapure distilled water to each reaction well using a 12-channel pipette and mix well. 12. Seal the plate with an optically clear adhesive PCR film. 13. Proceed to the next step, or store the dsDNA insert plate at 20  C. 3.2.2 Ligation Assembly

1. To perform the ligation assembly for each dsDNA insert, prepare 50 reaction mix.

208

Soh Ishiguro and Nozomu Yachie

1 reaction unit Component

Volume (μL)

1/10 diluted annealed oligo

2.5

10 T4 ligase buffer with 10 mM ATP

0.625

1 mg/mL BSA

0.31

T4 DNA ligase

0.2

BsmBI

0.2

25 ng/μL backbone plasmid (pSI-356 v2)

0.25

Ultrapure distilled water

2.165

Total

6.25

50 reaction mix Component

Volume (μL)

10 T4 ligase buffer with 10 mM ATP

31.25

1 mg/mL BSA

15.5

T4 DNA ligase

10.0

BsmBI

10.0

25 ng/μL backbone plasmid (pSI-356 v2)

12.5

Ultrapure distilled water

108.25

Total

187.5

2. Add 2.5 μL of insert dsDNA each into a fresh 96-well PCR plate. 3. Add 3.75 μL of the ligation assembly mix into each well and slowly mix with the insert dsDNA by pipetting up and down. 4. Seal the reaction plate with an optically clear adhesive PCR film. 5. Spin down the reaction plate. 6. Incubate the reaction plate in a thermal cycler with the following conditions. Step

Temperature 

Time

1

37 C

5’00

2

20  C

5’00

Go to 1

(14 times) 

3

55 C

30’00

4

4 C

1

7. Proceed to the next step, or store the reaction plate 20  C.

High-Throughput Analysis of CRISPR Genome Editing Outcomes 3.2.3 Bacterial Transformation

209

1. Thaw chemically competent E. coli cells on ice (a total volume of 480 μL for 48 reactions). 2. Aliquot 10 μL of the competent cells into a fresh 96-well PCR plate. 3. Put a 96-well aluminum block on ice to cool down. 4. Transfer 2 μL of the ligation assembly product into each sample well of the transformation reaction plate using a 12-channel pipette. 5. Seal the transformation reaction plate with an optically clear adhesive PCR film. 6. Place the transformation reaction plate on the ice-chilled 96-well aluminum block and incubate on ice for 30 min. 7. Heat-shock the transformation samples at 42  C for 30 s by floating the reaction plate on a preheated water bath. 8. Immediately place the transformation reaction plate on the ice-chilled aluminum block and incubate for 2 min. 9. Remove the optically clear adhesive PCR film. 10. Add 50 μL SOC outgrowth medium into each sample well of the reaction plate using a 12-channel pipette. 11. Seal the reaction plate with an optically clear adhesive PCR film. 12. Place the reaction plate in a 37  C bacterial incubator and incubate for 10 min. 13. During the incubation, airdry 5–6 LB+Amp plates on a clean bench. 14. Remove the optically clear adhesive PCR film and briefly mix the samples by pipetting up and down several times using a 12-channel pipette. 15. Spot 10 μL of the transformation sample each on an LB+Amp agar plate (we usually spot 12 samples per 10-cm petri dish). 16. Incubate the LB+Amp sample plates overnight in a 37  C bacterial incubator. 17. The bacterial plates should be stored at 4  C until the obtained clones are validated in the following quality check procedure.

3.2.4 Plasmid Purification and Sanger Sequencing

1. Isolate two to three colonies for each gRNA and inoculate each to 5 mL LB+Amp liquid media in a 15-mL culture tube (see Note 6). 2. Incubate the sample tubes overnight at 37  C using a tube rotator or shaker. 3. Purify plasmid DNAs by miniprep.

210

Soh Ishiguro and Nozomu Yachie

4. Validate the inserts by Sanger sequencing. The sequencing primer sequence is provided in Subheading 2.3. 5. Store the validated plasmid DNAs at 20  C. 3.3

Culturing of Cells

3.3.1 Collagen Coating of Cell Culture Plates

Here we describe the culturing of HEK293T cells that are used for the genome-editing assay (described in Subheading 3.4). HEK293T cells are expanded and seeded into the wells of 96-well cell culture plates. One and a half 96-well cell culture plates are required for each genome-editing enzyme (or control enzyme) to perform 48 genome-editing assays with unique gRNAs in triplicates. 1. Prepare a 20-mL collagen-I coating solution in a disposable reagent reservoir. 2. Apply 50 μL of collagen-I coating solution into each well of fresh 96-well cell culture plates using a 12-channel pipette. 3. Incubate 30 min in a cell culture incubator at 37  C and 0.05% CO2. 4. Remove the solution from the cell culture plates using a 12-channel pipette. 5. Add 50 μL 1 PBS to each well of the cell culture plates to wash the collagen-I coating solution and remove the reagent using a 12-channel pipette. 6. Proceed to the next step, or the collagen-coated 96-well cell culture plates can be stored for at least 1 week at room temperature, wrapped with aluminum foil.

3.3.2 Seeding Cells into 96-Well Collagen-Coated Cell Culture Plates

1. Place a DMEM bottle in a 37  C water bath to warm up. 2. Thaw 2-mL HEK293T cell stock vials in a 37  C water bath. Make sure to have enough cells for a target screening space. 3. Transfer the cells from each stock vial to 5 mL of the prewarmed DMEM in a 15-mL sample tube and centrifuge at 200  g for 5 min. 4. Aspirate and remove the supernatant. 5. Add the prewarmed DMEM, 10 mL each, into the sample tubes. 6. Gently mix the cells by pipetting several times. 7. Transfer the cells from each sample to a 10-cm cell culture dish. 8. Incubate the cell culture dishes for 2–3 days in a cell culture incubator at 37  C and 0.05% CO2 to allow cell proliferation to 80–90% confluency (see Note 3). 9. Remove the medium using an aspirator.

High-Throughput Analysis of CRISPR Genome Editing Outcomes

211

10. Apply 1 PBS, 1000 μL each, to the cell culture dishes. 11. Remove the reagent from the cell culture dishes using an aspirator. 12. Apply 0.05% trypsin-EDTA, 1000 μL, each to the cell culture dishes. 13. Incubate the cell culture dishes for 4 min in a cell culture incubator at 37  C and 0.05% CO2. 14. Apply DMEM, 1000 μL each, to the cell culture dishes and slowly mix by pipetting to obtain a cell suspension. 15. Transfer the cells from each cell culture dish into a 15-mL tube containing 5 mL of DMEM. 16. Quantify cell concentrations using a cell counter and adjust to 25 cells/μL using DMEM. 17. Transfer the cells from the sample tubes into a sterile disposable reagent reservoir. 18. Transfer cell sample, 200 μL each (5000 cells), to the collagencoated 96-well cell culture plate wells using a 12-channel pipette. For each of the genome-editing or control enzymes, three four-row units (i.e., three units of rows A–D or E–H of 96-well cell culture plates) are required to allow genome-editing assays with 48 gRNAs in triplicates. 19. Incubate the assay plates for 12–15 h in a cell culture incubator at 37  C and 0.05% CO2 and immediately proceed with the next step (see Note 7). 3.4 Massively Parallel Transfection of Genome-Editing Reagents

3.4.1 gRNA Reagent Preparation

Here we describe massively parallel transfections of genome-editing reagents to HEK293T cells in the 96-well plates prepared as described in Subheading 3.3 with an efficient strategy for assembling different gRNAs and different genome-editing enzymes using a 12-channel pipette (Fig. 6). Each reaction transfects 120 ng of a genome-editing (or control) enzyme expression plasmid and 40 ng of a gRNA expression plasmid to the cells using PEI Max transfection reagent. After transfection, the cells are incubated for 3 days to induce genome editing. The assay conditions may need to be modified for other cell lines by changing input plasmid amounts, transfection reagent, cell density, and/or incubation time. 1. Measure the DNA concentration of each gRNA expression plasmid using a spectrophotometer immediately before the transfection assay. 2. Adjust concentrations of gRNA plasmids to 25 ng/μL using ultrapure distilled water and assemble, 100 μL each, in a fresh 96-well PCR plate. Fill rows A–D of the PCR plate for

212

Soh Ishiguro and Nozomu Yachie

Fig. 6 Assembly and transfection of genome editing reagents for each enzyme

48 gRNAs, so that the well positions are in concordance with the well positions of the target amplification primers in the curated primer mix plate. 3. Seal the gRNA library plate with an optically clear adhesive PCR film. 4. Centrifuge the gRNA library plate at 1000  g for 2 min. The plate can be stored long term at 20  C. 3.4.2 Transfection

1. Prepare genome-editing enzyme plasmids and a control enzyme plasmid by midiprep. 2. Measure and adjust the DNA concentrations of each enzyme plasmid to 500 ng/μL (see Note 8). 3. To perform genome-editing assays for 48 gRNAs in triplicates, prepare 200 enzyme mix in a 15-mL sample tube for each of the genome-editing and control enzymes. Note that a sufficient excess of the reagent mix is necessary for the following two layers of aliquoting procedures. 1 reaction unit Component

Volume (μL)

500 ng/μL genome-editing or control plasmid

0.24

25 ng/μL gRNA plasmid

1.6

1 mg/mL PEI Max

0.48

1 PBS

50.0

Total

52.32

High-Throughput Analysis of CRISPR Genome Editing Outcomes

213

200 reaction mix Component

Volume (μL)

500 ng/μL genome-editing or control plasmid

48

1 mg/mL PEI max

320

1 PBS

10,000

Total

10,368

4. Mix the enzyme mixes by vortexing. 5. For each genome-editing (or control) enzyme, prepare a fresh 96-well round-bottom deep-well plate as the transfection mix plate and aliquot 800 μL of the enzyme mix to every well in row H (H1–H12). 6. Take gRNA expression plasmids, 5.5 μL each, from the gRNA library plate (48 samples) and apply to the corresponding positions of the transfection mix plates (rows A–D) for each genome-editing (or control) enzyme using a 12-channel pipette. 7. For each genome-editing (or control) enzyme, using a 12-channel pipette, take 180 μL of the enzyme mix from every well of row H in the transfection mix plate and apply it to row A of the same transfection mix plate and mix well by pipetting. Repeat the same procedure for destination rows B–D. 8. Seal the transfection mix plates with optically clear adhesive PCR films and incubate for 15 min at room temperature. 9. Retrieve the assay plates prepared in Subheading 3.3.2 from the cell culture incubator. 10. Apply transfection mixes, 52.32 μL each, to the designated wells of the assay plates using a 12-channel pipette. Note that each genome-editing (or control) enzyme assay requires three four-row units of the assay plates for genome-editing assays with 48 unique gRNAs in triplicates (i.e., a four-row unit of a transfection mix is stamped three times to one and a half 96-well assay plates). 11. Incubate the assay plates for 72 h in a cell culture incubator at 37  C and 0.05% CO2. 3.5 Template Genomic DNA Preparation

Here we describe genomic DNA template preparation for the amplicon sequencing procedure. The genome-editing treated cell samples in the 96-well assay plates are lysed with NaOH and boiling. and neutralized by Tris–HCl. This simple cell lysis is sufficient for the downstream amplicon sequencing library preparation procedure. Unlike column-based genomic DNA extraction protocols, this protocol processes one assay sample plate (96 samples) in 20 min with minimal hands-on time and also can be implemented for use with robotic liquid handling systems for further scalability.

214

Soh Ishiguro and Nozomu Yachie

1. Remove cell culture medium (about 250 μL each) from the assay plates using a 12-channel pipette (see Note 9). 2. Put 20 mL of freshly prepared 50 mM NaOH in a disposable reagent reservoir. 3. Using a 12-channel pipette, transfer 50 μL of the NaOH solution to every sample well of the assay plates. 4. Using a 12-channel pipette, carefully and gently transfer the entire lysed cell samples to fresh 96-well PCR plates. The cell lysate will be sticky. Slowly pipet the samples. 5. Seal the sample PCR plates with optically clear adhesive PCR films. 6. Centrifuge the sample PCR plates at 1050  g for 2 min. 7. Incubate the sample PCR plates at 95  C for 15 min, then cool to 4  C using a thermal cycler. 8. Centrifuge the sample PCR plates at 1050  g for 2 min. 9. Prepare 10 mL of 1 M Tris–HCl (pH 8.0) in a disposable reagent reservoir. 10. Remove the adhesive PCR films and carefully add 5 μL of the Tris–HCl solution into every sample well of the sample PCR plates for neutralization using a 12-channel pipette. The Tris– HCl volume should be 10% of the input NaOH volume. 11. Seal the sample PCR plates again with optically clear adhesive PCR films (see Note 10). 12. Centrifuge the sample PCR plates at 1050  g for 2 min to spin down the remaining cell lysates on well walls and to remove air bubbles. 13. Proceed to the next step, or the cell lysate plates can be store at 20  C for a few months or 80  C for a long time. 3.6 Generation of a Multiplexed Amplicon Sequencing Library and Sequencing

In this final section, we describe parallel amplification of genomeediting target sites, pooling and indexing of the PCR products for each genome-editing (or control) enzyme, and generation of a multiplexed amplicon sequencing library with quality controls for one-shot Illumina sequencing to evaluate hundreds to thousands of genome-editing samples. Target genomic regions of genomeedited samples are PCR amplified using the corresponding primers screened in Subheading 3.1. Then, PCR products of similar sizes are all pooled for each genome-editing enzyme of each assay replicate and subjected to magnetic bead-based purification. The pooled PCR samples are reamplified by a second-round PCR to add Illumina sequencing adapters with unique indices. The indexed sequencing libraries are separately size-selected by agarose gel electrophoresis and quantified by qPCR (Fig. 7). Equal molar quantities of the indexed libraries are further pooled as a single multiplexed library and analyzed using an Illumina sequencer.

High-Throughput Analysis of CRISPR Genome Editing Outcomes P5 index

P5

P7 index

PS1.0

Target region

PS2.0

Read1

Lib 1

Lib 2

215

P7 Read2

Lib 3

Lib 4

Lib 5

Lib 6

Lib 7

Lib 8

Lib 9

500bp 300bp

Fig. 7 Preparation of amplicon sequencing libraries. The first-round PCR product pools of different assay enzymes of different replicates are reamplified by a second-round PCR. The PCR products are then sizeselected by agarose gel electrophoresis. The yellow boxes show the typical area for the size selection

After sequencing, sequencing reads can be demultiplexed into individual genome-editing assays according to their custom indices and endogenous target sequences. Sequencing reads of each assay sample can be used for mutation outcome pattern analyses. 3.6.1 PCR Amplification of the Target Regions

1. Thaw the curated primer mix plates (48 primer pairs) established in Subheading 3.1 at room temperature and centrifuge at 1000  g for 2 min. 2. Retrieve the cell lysate plates established in Subheading 3.5 (thaw if frozen) and centrifuge at 1000  g for 2 min. 3. To perform the following target amplification PCRs, prepare 180 PCR master mix for each genome-editing (or control) enzyme for a total of triplicate samples (scale the master mix volume with the number of genome-editing and control enzymes used). 1 reaction unit Component

Final concentration

Volume (μL)

5 Phusion HF buffer

1

4

2 mM dNTPs

200 μM

2

10 μM primer mix

500 nM each

2

DNA template (cell lysate)

2

Phusion DNA polymerase

0.2

Ultrapure distilled water

9.8

Total

20

216

Soh Ishiguro and Nozomu Yachie

180 PCR master mix Component

Volume (μL)

5 Phusion HF buffer

480

2 mM dNTPs

240

Phusion DNA polymerase

24

Ultrapure distilled water

1764

Total

2508

4. Transfer the PCR reaction mix into a disposable reagent reservoir placed on ice. 5. Prepare the same number of fresh 96-well PCR plates as the assay plates prepared in Subheading 3.3.2. Label the PCR plates to correspond to the assay plates. Using a 12-channel pipette, transfer the PCR master mix, 17 μL each, from the reservoir to all the wells of the PCR plates. 6. Using a 12-channel pipette, take 2 μL each from the four-row unit of the curated primer mix plate (rows A–D) and apply them to the top four-row unit (rows A–D) of the first PCR plate. Repeat the same procedure until both of the top and bottom four-row units of the all the PCR plates are filled up. 7. Using a 12-channel pipette, take 2 μL each from the cell lysate plates and apply them to the corresponding well positions of the PCR plates. Gently mix the samples by pipetting. 8. Seal the PCR plates with optically clear adhesive PCR films. 9. Centrifuge the PCR plates at 1000  g for 2 min. 10. Perform the first-round target amplification PCR with the following thermal cycle conditions. Step

Temperature

Time

1

98  C

0’30

2

98  C

0’10

3 4



0’10



1’00

60 C 72 C

Go to 2 5 6

(29 times) 

72 C 

4 C

5’00 1

11. Centrifuge the first-round PCR sample plates at 1000  g for 2 min.

High-Throughput Analysis of CRISPR Genome Editing Outcomes

217

12. Proceed to the bead-based DNA purification step, or store the PCR sample plates at 20  C. 3.6.2 Purification of the First PCR Products

1. Retrieve AMPure XP beads from a fridge, settle to room temperature, and vortex thoroughly before use (see Note 11). 2. Centrifuge the first-round PCR sample plates at 1000  g for 2 min. 3. Gently remove the plate seals. 4. For each genome-editing (or control) enzyme of each assay replicate, pool and combine the first-round PCR products, 3 μL each, in a 1.5-mL tube. This yields 144-μL PCR sample pools each containing amplicon products for 48 gRNA targeting regions. 5. Add 259.2 μL of AMPure XP beads (1.8 volume) to each first-round PCR sample pool and thoroughly mix by pipetting up and down several times without making bubbles. 6. Incubate the sample tubes at room temperature for 5 min to allow PCR products to bind to the beads. 7. Place the sample tubes on a magnetic stand and wait for 1 min to allow magnetic separation of beads. 8. Without removing the sample tubes from the magnetic stand, carefully remove all of the clear supernatants. 9. Add 500 μL of 70% EtOH to each sample tube on the magnetic stand and wait for 30 s. 10. Without removing the sample tubes from the magnetic stand, carefully remove all of the clear supernatants. 11. Repeat steps 9 and 10 one more time. Make sure no EtOH remains in the sample tubes. 12. Remove the sample tubes from the magnetic stand and airdry for 1–3 min. 13. Apply 50 μL of ultrapure distilled water and gently mix with the magnetic beads by pipetting. 14. Incubate the sample tubes at room temperature for 2 min to allow PCR products to elute in the water solvent. 15. Place the sample tubes on the magnetic stand and wait for 2 min to allow magnetic separation of beads. 16. Without removing the samples tubes from the magnetic stand, transfer the clear supernatants each to a fresh 1.5-mL tube. 17. Measure DNA concentrations using a spectrophotometer. 18. Adjust sample concentrations to 10 ng/μL with ultrapure distilled water.

218

Soh Ishiguro and Nozomu Yachie

19. Proceed with the next pooled indexing PCRs, or store the firstround PCR pool samples at 20  C. 3.6.3 Pooled Indexing PCR for Multiplexed Sequencing

1. Vortex the first-round PCR pool samples and spin down well using a benchtop centrifuge. 2. To perform the following second-round indexing PCR for each pooled sample, prepare 1.1 second-round PCR master mix. Scale up the volume of the master mix with the number of the first-round PCR pool samples (the number of genome-editing or control enzymes multiplied by three for assay replicates). 1 reaction unit Component

Final concentration

Volume (μL)

5 Phusion HF buffer

1

8

2 mM dNTPs

200 μM

4

10 μM P5 index primer

500 nM

2

10 μM P7 index primer

500 nM

2

First-round PCR pool

1

Phusion DNA polymerase

0.4

Ultrapure distilled water

22.6

Total

40

1.1 master mix Component

Volume (μL)

5 Phusion HF buffer

8.8

2 mM dNTPs

4.4

Phusion DNA polymerase

0.44

Ultrapure distilled water

24.86

Total

38.5

3. Aliquot the PCR master mix, 35 μL each, to individually capped eight-strip PCR tubes. 4. Apply unique combinations of custom Illumina P5 and P7 index primers to the PCR reaction tubes. See Subheading 2.7 for the design of the custom Illumina index primers. 5. Apply the first-round PCR sample pools 1 μL each to the PCR reaction tubes and mix gently by pipetting. Make sure unique index pairs are assigned to the first-round PCR sample pools. 6. Spin down the PCR reaction tubes using a benchtop centrifuge.

High-Throughput Analysis of CRISPR Genome Editing Outcomes

219

7. Perform PCR with the following thermal cycle conditions. Step

Temperature

Time

1

98  C

0’30

2



0’10



98 C

3

60 C

0’10

4

72  C

2’00

Go to 2 5 6

(14 times) 

72 C 

4 C

5’00 1

8. Perform gel electrophoresis analysis with 2% agarose gel to check the rough yields and size distributions of the reamplified PCR products. 9. For each reamplified PCR product, cut the gel to obtain the PCR product in the expected size range (Fig. 7) and purify the amplicon sequencing libraries using a PCR purification kit (see Note 12). 10. Elute the size-selected samples into 20 μL of ultrapure distilled water and collect the eluate in a 1.5-mL LoBind DNA tube. 11. Check the concentration for each amplicon sequencing library using a spectrophotometer (see Note 13). 12. Proceed to the next step below, or store the amplicon sequencing libraries at 20  C. 3.6.4 Multiplexed Amplicon Sequencing

1. Retrieve the amplicon sequencing libraries (and thaw if frozen). 2. Vortex the sample tubes well and spin down using a benchtop centrifuge. 3. Using KAPA Library Quantification Kits (Illumina), quantify the absolute concentrations of the amplicon sequencing libraries according to the manufacturer’s instruction (see Note 14). 4. Pool amplicon sequencing libraries in a 1.5-mL LoBind DNA tube so that each library is represented equally. 5. Using KAPA Library Quantification Kits (Illumina), quantify the absolute concentration of the multiplexed library according to the manufacturer’s instruction. 6. Analyze the multiplexed amplicon sequencing library with 20% PhiX spike-in using an Illumina sequencer (see Note 15). The library can be stored long term at 20  C (see Note 16).

220

4

Soh Ishiguro and Nozomu Yachie

Notes 1. We used pSI-356 v2 in our previous studies [4, 35]. pSI-356 v2 encodes the human U6 promoter and 2-kb filler sequence sandwiched by two BsmBI cloning sites followed by the gRNA scaffold sequence. The filler DNA fragment is replaced with a target spacer-encoding fragment to obtain a gRNA expression plasmid. 2. Prepare a fresh NaOH solution immediately before the experiment every time. 3. For better transfection efficiency, use low passage HEK293T cells that have undergone no more than 10 passages since their establishment. 4. PCR efficiency might be affected by the amount of cell lysate. The template cell lysate concentration may need to be optimized. 5. To make the gRNA construction easy with a 12-channel pipette, order forward and reverse ssDNA oligos in the 96-well format. Obtain corresponding forward and reverse ssDNA oligos in the same positions of two separate plates. We also recommend obtaining the ssDNA oligos dissolved in 100 μM water or TE instead of a dry shipping option that requires the additional steps of dissolving and normalizing the ssDNA oligo samples. 6. Two to three colonies are usually sufficient to obtain a correct plasmid clone. 7. Before proceeding to the next step, use a microscope to confirm that the seeded cells are uniformly distributed in the well and adhered to the flat well bottom. 8. Midiprep from a 150–200 mL bacterial culture with 12–15 h incubation at 37  C should be sufficient to obtain a genomeediting enzyme expression plasmid for transfection assays with 48 gRNAs in triplicates several times. Note that some genomeediting enzyme expression plasmids are toxic for bacterial growth and may need further optimization in this step. 9. The experiment can be stopped here. After completely removing the culture medium, the assay plates can be stored at 20  C. 10. Cell lysate samples are sticky. PCR films need to be removed gently to avoid cross-well sample contaminations. 11. AMPure XP beads are sufficient to remove residual primers and primer dimers from the first-round PCR products. We recommend size-selection using agarose gel electrophoresis when strong nonspecific band signals are observed in a gel. The

High-Throughput Analysis of CRISPR Genome Editing Outcomes

221

purity of a first-round PCR product greatly affects the result of the downstream second-round PCR reamplification with custom Illumina index adapters. 12. The second-round PCR products usually show smear bands on a gel in a range of 50 bp from the expected product size. According to our amplicon sequencing data, we assume this may be caused by indel mutations induced by genome editing. We recommend to size-select a wider area of the PCR bands than what is for usual for molecular cloning. 13. If the measured concentration is more than 10 ng/μL by spectrophotometry, adjust the sample concentration to 5–10 ng/μL with ultrapure distilled water so that the sample quantity is within the range of the of standards in the downstream qPCR. 14. Because the size distribution of the amplicon sequencing library can be expected by design, we prefer to use qPCR for library quantification and normalization rather than other quantification methods, such as TapeStation or BioAnalyzer. 15. The amplicon sequencing library should be sequenced (pairedend) with 20%–30% PhiX spike-in because the library complexity for each sequencing cycle is relatively low, and this is known to restrict sequencing cluster identification, at least slightly, in an Illumina sequencer. From 20,000 to 50,000 sequencing reads for each genome-editing assay is usually sufficient for further data analyses. We usually choose MiSeq Micro (total output five million reads) for less than 200 genome editing assays or MiSeq v3 (total output 25 million reads) for larger assays. 16. DNA binds to the tube wall even if LoBind tubes are used. The sequencing library should be quantified by qPCR no more than 2–3 days before the sequencing run.

Acknowledgments We thank Hideto Mori for optimizing the protocol in concordance with data analysis pipelines and Mamoru Tanaka for optimizing the detailed experimental steps. This study was funded by the Japan Agency for Medical Research and Development (AMED) Platform Project for Supporting Drug Discovery and Life Science Research (to N.Y.), the New Energy and Industrial Technology Development Organization (NEDO) and AMED PRIME program (17gm6110007) to N.Y., and the Japan Society for the Promotion of Science (JSPS) Grant-in-Aid for Scientific Research (16 J06287) to S.I.

222

Soh Ishiguro and Nozomu Yachie

References 1. Cong L, Ran FA, Cox D et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339:819–823 2. Mali P, Yang L, Esvelt KM et al (2013) RNA-guided human genome engineering via Cas9. Science 339:823–826 3. Hsu PD, Lander ES, Zhang F (2014) Development and applications of CRISPR-Cas9 for genome engineering. Cell 157:1262–1278 4. Nishimasu H, Shi X, Ishiguro S et al (2018) Engineered CRISPR-Cas9 nuclease with expanded targeting space. Science 361:1259–1262 5. Hu JH, Miller SM, Geurts MH et al (2018) Evolved Cas9 variants with broad PAM compatibility and high DNA specificity. Nature 556:57–63 6. Walton RT, Christie KA, Whittaker MN et al (2020) Unconstrained genome targeting with near-PAMless engineered CRISPR-Cas9 variants. Science 368:290–296 7. Kleinstiver BP, Pattanayak V, Prew MS et al (2016) High-fidelity CRISPR-Cas9 nucleases with no detectable genome-wide off-target effects. Nature 529:490–495 8. Slaymaker IM, Gao L, Zetsche B et al (2016) Rationally engineered Cas9 nucleases with improved specificity. Science 351:84–88 9. Casini A, Olivieri M, Petris G et al (2018) A highly specific SpCas9 variant is identified by in vivo screening in yeast. Nat Biotechnol 36:265–271 10. Kuscu C, Parlak M, Tufan T et al (2017) CRISPR-STOP: gene silencing through baseediting-induced nonsense mutations. Nat Methods 14:710–712 11. Nishida K, Arazoe T, Yachie N et al (2016) Targeted nucleotide editing using hybrid prokaryotic and vertebrate adaptive immune systems. Science 353:aaf8729 12. Rees HA, Liu DR (2018) Base editing: precision chemistry on the genome and transcriptome of living cells. Nat Rev Genet 19:770–788 13. Komor AC, Kim YB, Packer MS et al (2016) Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533:420–424 14. Gaudelli NM, Komor AC, Rees HA et al (2017) Programmable base editing of A·T to G·C in genomic DNA without DNA cleavage. Nature 551:464–471 15. Huang TP, Zhao KT, Miller SM et al (2019) Circularly permuted and PAM-modified Cas9

variants broaden the targeting scope of base editors. Nat Biotechnol 37:626–631 16. Rees HA, Komor AC, Yeh WH et al (2017) Improving the DNA specificity and applicability of base editing through protein engineering and protein delivery. Nat Commun 8:15790 17. Koblan LW, Doman JL, Wilson C et al (2018) Improving cytidine and adenine base editors by expression optimization and ancestral reconstruction. Nat Biotechnol 36:843–846 18. Zafra MP, Schatoff EM, Katti A et al (2018) Optimized base editors enable efficient editing in cells, organoids and mice. Nat Biotechnol 36:888–893 19. Wang X, Li J, Wang Y et al (2018) Efficient base editing in methylated regions with a human APOBEC3A-Cas9 fusion. Nat Biotechnol 36:946–949 20. Zhang X, Chen L, Zhu B et al (2020) Increasing the efficiency and targeting range of cytidine base editors through fusion of a singlestranded DNA-binding protein domain. Nat Cell Biol 22:740–750 21. Zhou C, Sun Y, Yan R et al (2019) Off-target RNA mutation induced by DNA base editing and its elimination by mutagenesis. Nature 571:275–278 22. Grunewald J, Zhou R, Garcia SP et al (2019) Transcriptome-wide off-target RNA editing induced by CRISPR-guided DNA base editors. Nature 569:433–437 23. Grunewald J, Zhou R, Iyer S et al (2019) CRISPR DNA base editors with reduced RNA off-target and self-editing activities. Nat Biotechnol 37:1041–1048 24. Rees HA, Wilson C, Doman JL et al (2019) Analysis and minimization of cellular RNA editing by DNA adenine base editors. Sci Adv 5:eaax5717 25. Ran FA, Cong L, Yan WX et al (2015) In vivo genome editing using Staphylococcus aureus Cas9. Nature 520:186–191 26. Kim E, Koo T, Park SW et al (2017) In vivo genome editing with a small Cas9 orthologue derived from campylobacter jejuni. Nat Commun 8:14500 27. Li X, Qian X, Wang B et al (2020) Programmable base editing of mutated TERT promoter inhibits brain tumour growth. Nat Cell Biol 22:282–288 28. Kim YB, Komor AC, Levy JM et al (2017) Increasing the genome-targeting scope and precision of base editing with engineered Cas9-cytidine deaminase fusions. Nat Biotechnol 35:371–376

High-Throughput Analysis of CRISPR Genome Editing Outcomes 29. Anzalone AV, Koblan LW, Liu DR (2020) Genome editing with CRISPR-Cas nucleases, base editors, transposases and prime editors. Nat Biotechnol 38:824–844 30. Zetsche B, Gootenberg JS, Abudayyeh OO et al (2015) Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Cas system. Cell 163:759–771 31. Kleinstiver BP, Sousa AA, Walton RT et al (2019) Engineered CRISPR-Cas12a variants with increased activities and improved targeting ranges for gene, epigenetic and base editing. Nat Biotechnol 37:276–282 32. Li C, Zhang R, Meng X et al (2020) Targeted, random mutagenesis of plant genes with dual

223

cytosine and adenine base editors. Nat Biotechnol 38:875–882 33. Grunewald J, Zhou R, Lareau CA et al (2020) A dual-deaminase CRISPR base editor enables concurrent adenine and cytosine editing. Nat Biotechnol 38:861–864 34. Zhang X, Zhu B, Chen L et al (2020) Dual base editor catalyzes both cytosine and adenine base conversions in human cells. Nat Biotechnol 38(7):856–860 35. Sakata RC, Ishiguro S, Mori H et al (2020) Base editors for simultaneous introduction of C-to-T and A-to-G mutations. Nat Biotechnol 38(7):865–869

Chapter 13 Optical Control of Genome Editing by Photoactivatable Cas9 Takahiro Otabe, Yuta Nihongaki, and Moritoshi Sato Abstract The CRISPR-Cas9 system offers targeted genome manipulation with simplicity. Combining the CRISPRCas9 with optogenetics technology, we have engineered photoactivatable Cas9 to precisely control the genome sequence in a spatiotemporal manner. Here we provide a detailed protocol for optogenetic genome editing experiments using photoactivatable Cas9, including that for the generation of guide RNA vectors, light-mediated Cas9 activation, and quantification of genome editing efficiency in mammalian cells. Key words Genome editing, CRISPR-Cas9, Guide RNA, Optogenetics, T7E1 assay, Genomic PCR, NHEJ, HDR

1

Introduction The CRISPR-Cas9 system derived from S. pyogenes provides a way of simple and efficient genome editing in diverse organisms [1]. The CRISPR-Cas9 consists of Cas9 protein and a single guide RNA (sgRNA). The Cas9-sgRNA complex scans genomic DNA to find and cleave DNA that is complementary to 20 nucleotides target sequence at the 50 -end of sgRNA. Because the target sequence of sgRNA can be readily programmed, Cas9 can target any user-defined sequence as long as a protospacer adjacent motif (PAM) in the form of NGG presents next to the 30 -end of the 20 nucleotides target sequence. Cas9-mediated double-strand breaks at target genomic site induce endogenous DNA repair pathways in the cell. Nonhomologous end joining (NHEJ) is an error-prone repair, which introduces small DNA insertions and deletions (indels) at the repaired target site, resulting in gene knockout by frameshift indels. In contrast, homology-directed repair (HDR) enables precise DNA repair. By introducing exogenous DNA donor with point mutation, genome modification with desired mutation can be achieved by HDR-mediated genome editing.

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

225

226

Takahiro Otabe et al.

a pCMV N713-pMag

NLS

N713

GS16

pMag

GS16

pCMV nMag-C714

GS16

nMag

GS16

C714

NLS

b

Fig. 1 Construction and schematic of the photoactivatable Cas9 (paCas9). (a) Constructs for paCas9. N713fragment and C714-fragment of Cas9 are fused with the Magnet system (pMag and nMag) to construct N713pMag and nMag-C714, respectively. (b) paCas9 is inactivated in the dark condition, and upon blue light illumination it is activated as a result of dimerization of the split Cas9 fragments by the Magnet system. Active paCas9 complexed with sgRNA can bind to the targeted DNA sequence. DNA double-strand break induced by paCas9 is repaired by NHEJ or HDR in mammalian cells. (Reproduced from Ref. 4 with permission from Springer Nature)

For further expanding application of Cas9 technology, many efforts of protein engineering have been converging on Cas9 to generate engineered Cas9 variants [2, 3]. We and other groups have developed optogenetic Cas9 tools which enable light control of Cas9-mediated genome perturbation with spatiotemporal resolution [4–12]. Photoactivatable Cas9 (paCas9) that we have engineered consists of a pair of split Cas9 fragments fused with lightinducible dimerization system, named Magnet system [4, 12]. The split Cas9 fragments are inactivated in the dark, and upon blue light illumination the Magnet system is heterodimerized, subsequently resulting in active Cas9 reconstitution to induce DNA doublestrand break targeted by sgRNA (Fig. 1). In this chapter, we describe the protocol for optogenetic genome editing in mammalian cell lines using paCas9. We outline the generation of mammalian expression vector for sgRNA. We also introduce plasmid HDR assay, which can measure paCas9 activity by bioluminescence reporter. In optogenetic genome editing

Optical Control of Genome Editing by Photoactivatable Cas9

227

experiment, NHEJ-mediated indel mutagenesis and HDR-mediated point mutation with single-stranded oligonucleotide donor are described with assay for quantifying the efficiency of genome editing outcome.

2

Materials

2.1 Media and Buffers

1. Lennox L Broth Base (LB) plus Amp: LB containing 100μg/ ml ampicillin. 2. Culture medium: Dulbecco’s Modified Eagle Medium (DMEM) with 10% fetal bovine serum (FBS), 100 unit/ml penicillin, 100μg/ml streptomycin. 3. Phenol red free culture medium: Dulbecco’s Modified Eagle Medium (DMEM) with 10% FBS, 100 unit/ml penicillin, 100μg/ml streptomycin. 4. 10 annealing buffer: 500 mM Tris–HCl (pH 7.5), 100 mM MgCl2, 10 mM dithiothreitol, 1000 mM NaCl. 5. 10 M buffer: 100 mM Tris–HCl (pH 7.5), 100 mM MgCl2, 10 mM dithiothreitol, 500 mM NaCl.

2.2 Molecular Biology

1. Hind III. 2. Bbs I. 3. T7 Endonuclease I. 4. D-luciferin: 500μM solution in HBSS. 5. Agarose gel. 6. DNA polymerase. 7. DNA dye. 8. DNA ligase. 9. Gel/PCR extraction kit. 10. Plasmid DNA miniprep kit. 11. Genomic DNA extraction mini kit. 12. E. coli: Mach1 competent cell or other suitable competent cell for general cloning. 13. Oligonucleotides for sgRNA sequence: 50 -CACCG(N)20-3 and 50 -AAAC(N)20C-30 . 14. 96-well black-walled cell culture plate. 15. 24-well clear-walled cell culture plate. 16. 0.2 ml PCR tube. 17. Milli-Q Water.

228

2.3

Takahiro Otabe et al.

Plasmids

1. pX330-U6-Chimeric_BB-CHh-hSpCas9 (Addgene Plasmid #42230). 2. pSPgRNA (Addgene Plasmid #47108). 3. pcDNA 3.1/V5-HisA (Invitrogen). 4. pGL4.31 vector (Promega). 5. pCold I vector (Clontech). 6. pCMV-N713-pMag (N-fragment of paCas9, see Note 1). 7. pCMV-nMag-C714 (C-fragments of paCas9, see Note 1). 8. pCMV-stopFluc reporter (see Note 1). 9. Luciferase donor plasmid (see Note 1).

2.4 Cell and Transfection Reagents

1. HEK293T.

2.5

1. Centro XS3 LB 960 microplate luminometer (BERETHOLD TECHNOLOGIES).

Instruments

2. Lipofectamine 2000 (Invitrogen). 3. SF cell line 4D-nucleofector X Kit S (Lonza).

2. 4D-nucleofector (Lonza). 3. E-shot II gel imaging system (ATTO). 4. 470 nm  20 nm LED light source (CCS Inc.). 5. Agarose gel electrophoresis equipment. 6. Thermal cycler.

3

Methods

3.1 Generation of Single-Guide RNA (sgRNA) Expression Vector (Fig. 2)

1. Synthesize a pair of oligonucleotides for sgRNA sequence. The top DNA strand is in the form of 50 -CACCG(N)20-30 , where (N)20 is user-defined target sequence. The bottom DNA strand is in the form of 50 -AAAC(N)20C-30 , where (N)20 is the reverse complement of the target sequence in the top strand (see Notes 2 and 3). The DNA oligos are resuspended at a concentration of 100μM in Milli-Q water. 2. Mix 1μl of the top DNA strand and 1μl of the bottom DNA strand together at 1:1 molar ratio in a 0.2 ml PCR tube along with 1μl of 10 annealing buffer and 7μl of Milli-Q water to make the final concentration of each oligo to equal 10μM. 3. Place the PCR tube in a thermal cycler and start the annealing program: (95  C, 10 min; 90–15  C, 1  C/1 min). 4. After running the annealing program, store the sample at 4  C. 5. Digest 1μg of pSPgRNA vector with Bbs I enzyme at 37  C for 60 min.

Optical Control of Genome Editing by Photoactivatable Cas9

229

Fig. 2 sgRNA cloning workflow. The pSPgRNA vector can be digested using the type IIS restriction enzyme Bbs I. The top DNA strand and the bottom DNA strand are designed based on the targeting locus sequence. (Append CACCG to the 50 -end of the top DNA strand, Append AAAC to the 50 -end of the bottom DNA strand). After Bbs I digestion, the annealed oligos can be ligated seamless into digested pSPgRNA vector

6. Purify the digested pSPgRNA vectro using Gel/PCR Extraction Kit. 7. DNA ligations are performed by incubating annealing oligos with appropriately linearized pSPgRNA vector using DNA ligation kit. 8. Transform the ligation product into chemical competent E. coli cells and plate onto standard LB plates containing 100μg/ml ampicillin. 9. Pick 2–3 colonies for isolating plasmids by Plasmid DNA miniprep kit. Select plasmids with correct target sequence by Sanger sequencing. Store at 20  C.

230

Takahiro Otabe et al.

Fig. 3 Experimental overview and functional characterization of paCas9. (a) Experimental overview of luciferase plasmid HDR assay. In this assay, the CMV promoter-driven luciferase containing an in-frame stop codon is cleaved by paCas9, and full length luciferase expression is then recovered through HDR with luciferase donor vector (lacking a promotor). Bioluminescence by luciferase expression indicates successful HDR event. (b) Characterization of DNA targeting specificity of paCas9. Activities of Cas9 and paCas9 targeting StopFluc-1 with a set of sgRNAs harboring single-nucleotide Watson-Crick transversion mutations. sgRNAs mutated in 50 -end G to C were not tested because 50 -end G is necessary for efficient expression from the U6 promoter. The positions of point mutations in sgRNA are indicated at the top of each panel. This result shows that the DNA targeting specificity is not significantly different between paCas9 and Cas9. (Reproduced from ref. 4 with permission from Springer Nature)

3.2 Luciferase Plasmid HDR Assay (Fig. 3)

1. Seed HEK293T cells at 2.0  104 cells/well into 96-well blackwalled cell culture plate and culture the cells for 24 h at 37  C in 5% CO2. 2. The cells are then transfected with Lipofectamine 2000 according to the manufacturer’s protocols. Plasmids encoding N713pMag, nMag-C714, sgRNA targeting stop codon, StopFluc reporter and luciferase donor plasmid are transfected at a 2.5:2.5:5:1:4 ratio. The total amount of DNA is 0.2μg/well. 3. After 24 h posttransfection, incubate the samples at 37  C in 5% CO2 under successive blue light illumination or in the dark for additional 48 h. Blue light illumination is performed using a 470 nm  20 nm LED light source. Blue light intensity is 1.2 W/m2.

Optical Control of Genome Editing by Photoactivatable Cas9

231

4. Remove the culture medium and then add 100μl of Phenol red free culture medium containing D-luciferin as a substrate. 5. After 30 min incubation at 37  C in 5% CO2, measure bioluminescence by Centro XS3 LB 960 microplate luminometer. 3.3 Optogenetic Genome Editing Experiments

1. To assay the NHEJ-mediated indel mutation, seed HEK293T cells at 1.0  104 cells/well in 24-well clear-walled cell culture plate, and culture the cells for 24 h at 37  C in 5% CO2. 2. Transfect the cells with the plasmids encoding N713-pMag, nMag-C714, and sgRNAs at 1:1:1 ratio using Lipofectamine 2000 according to the manufacturer’s protocols. As a positive control, transfect the plasmids encoding full-length Cas9 and sgRNAs at a 2:1 ratio. The total amount of DNA is 0.9μg/well. 3. After 24 h posttransfection, incubate the cells at 37  C in 5% CO2 under successive blue light illumination or in the dark as described above (see Subheading 3.2, step 3). 4. After incubation for 24 h, isolate genomic DNA using Genomic DNA extraction mini kit according to the manufacturer’s instructions. Isolated genomic DNA can be stored at 20  C. 5. To assay the HDR-mediated genome editing, nucleofect 6.0  105 HEK293T cells with plasmids encoding N713pMag, nMag-C714, sgRNA targeting EMX1 locus and single-stranded DNA donor using the SF cell line 4D-nucleofector X Kit S and the CA-189 program with 4D-nucleofector. Seed the nucleofected cells at 2.0  105 cells/well in 24-well plate. Twenty four hours after the nucleofection, incubate the cells at 37  C in 5% CO2 under successive blue light illumination or in the dark. After 48 h incubation, isolate the genomic DNA as described above (see Subheading 3.3, step 4).

3.4 T7 Endonuclease I (T7EI) Assay for Quantifying Indel Mutation of Endogenous Genes (Fig. 4)

1. Amplify the genomic region targeted by paCas9 using DNA polymerase. 2. Purify the genomic PCR amplicons using Gel/PCR Extraction Kits following the manufacturer’s protocol. 3. Combine the PCR amplicons with 2μl of 10  M buffer for restriction enzyme and Milli-Q water to make a final volume of 20μl, and denature and reanneal the PCR products to form heteroduplex DNA (95  C, 10 min; 90–15  C, 1  C/1 min). 4. Mix the heteroduplexed samples with 5 units of T7 endonuclease I incubate at 37  C for 30 min. 5. For the analysis of the DNA fragments, perform agarose gel electrophoresis. Stained the agarose gel with DNA dye and image it with E-shot II gel imaging system.

232

Takahiro Otabe et al.

Fig. 4 Optogenetic genome editing of mammalian endogenous genes by paCas9. Cells were transfected with paCas9 and sgRNAs separately targeting the EMX1 locus, the VEGFA locus and the two sites in AAVS1 locus. This result shows that paCas9 can induce indel mutations by NHEJ in a blue light dependent manner. (Reproduced from ref. 4 with permission from Springer Nature)

6. The indel mutation frequency can be quantified from relative band intensities. Use the following formula to calculate the percentage of indel mutation by paCas9: 100  (1  (1  (b + c)/(a + b + c))1/2), where a is the intensity of the uncleaved PCR product, and b and c are the intensities of each. 3.5 RFLP Assay for Detecting HDR-Mediated Modification in Endogenous Human Gene

1. Perform genomic PCR and purification as described above (see Subheading 3.4, steps 1 and 2). 2. Combine the PCR product with 30 units of Hind III, 2μl of 10 M buffer for restriction enzyme and Milli-Q water to make a final volume of 20μl, and incubate at 37  C for 30 min. 3. Analyze the digested products by agarose gel electrophoresis. Perform agarose gel staining and imaging as described above (see Subheading 3.4, step 5). 4. Quantify the HDR frequency from relative band intensities. Use the following formula to calculate the percentage of HDR by paCas9: 100  (b + c)/(a + b + c), where a is the intensity of the uncleaved PCR product, and b and c are the intensities of each Hind III enzyme digested product.

Optical Control of Genome Editing by Photoactivatable Cas9

4

233

Notes 1. Contact Moritoshi Sato ([email protected]) to request plasmids related paCas9. 2. Note that the PAM (NGG) sequence should not be included in the user-defined targeting sequence of sgRNA. Because the U6 promoter prefers “G” as the first transcribe nucleotide and the first “G” helps transcription more efficient, we recommend to add “G” at the 50 -end of the target sequence if the sequence does not include “G” at its 50 -end. 3. Useful web sites for sgRNA design are described in the Ref. 13.

Acknowledgments We would like to thank CREST grants (JPMJCR1653) from Japan Science and Technology Agency. This work was supported by a project grant from Kanagawa Institute of Industrial Science and Technology (KISTEC) to M.S. References 1. Ran FA, Hsu PD, Wright J, Agarwala V, Scott DA, Zhang F (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8 (11):2281–2308 2. Dominguez AA, Lim WA, Qi LS (2016) Beyond editing: repurposing CRISPR–Cas9 for precision genome regulation and interrogation. Nat Rev Mol Cell Biol 17(1):5–15 3. Mitsunobu H, Teramoto J, Nishida K, Kondo A (2017) A beyond native Cas9: manipulating genomic information and function. Trends Biotechnol 35(10):983–996 4. Nihongaki Y, Kawano F, Nakajima T, Sato M (2015) Photoactivatable CRISPR-Cas9 for optogenetic genome editing. Nat Biotechnol 33(7):755–760 5. Nihongaki Y, Furuhata Y, Otabe T, Hasegawa S, Yoshimoto K, Sato M (2017) CRISPR-Cas9-based photoactivatable transcription systems to induce neuronal differentiation. Nat Methods 14(10):963–966 6. Nihongaki Y, Otabe T, Ueda Y, Sato M (2019) A split CRISPR-Cpf1 platform for inducible genome editing and gene activation. Nat Chem Biol 15(9):882–888 7. Nihongaki Y, Yamamoto S, Kawano F, Suzuki H, Sato M (2015) CRISPR-Cas9based photoactivatable transcription system. Chem Biol 22(2):169–174

8. Bubeck F, Hoffmann MD, Harteveld Z, Aschenbrenner S, Bietz A, Waldhauer MC, Bo¨rner K, Fakhiri J, Schmelas C, Dietz L, Grimm D, Correia BE, Eils R, Niopek D (2018) Engineered anti-CRISPR proteins for optogenetic control of CRISPR-Cas9. Nat Methods 15(11):924–927 9. Mathony J, Hoffmann MD, Niopek D (2020) Optogenetics and CRISPR: a new relationship built to last. Methods Mol Biol 2173:261–281 10. Yu Y, Wu X, Guan N, Shao J, Li H, Chen Y, Ping Y, Li D, Ye H (2020) Engineering a far-red light–activated split-Cas9 system for remote-controlled genome editing of internal organs and tumors. Sci Adv 6(28):eabb1777 11. Zhou XX, Zou X, Chung HK, Gao Y, Liu Y, Qi LS, Lin MZ (2018) A single-chain Photoswitchable CRISPR-Cas9 architecture for light-inducible gene editing and transcription. ACS Chem Biol 13(2):443–448 12. Kawano F, Suzuki H, Furuya A, Sato M (2015) Engineered pairs of distinct photoswitches for optogenetic control of cellular proteins. Nat Commun 6(1):6256 13. Hanna RE, Doench JG (2020) Design and analysis of CRISPR-Cas experiments. Nat Biotechnol 38(7):813–823

Part IV Engineering Mammalian Cells in Combination with Chemical Compounds/Systems

Chapter 14 Chemogenetic Control of Protein Localization and Mammalian Cell Signaling by SLIPT Sachio Suzuki, Yuka Hatano, Tatsuyuki Yoshii , and Shinya Tsukiji Abstract Chemical control of protein localization is a powerful approach for manipulating mammalian cellular processes. Self-localizing ligand-induced protein translocation (SLIPT) is an emerging platform that enables control of protein localization in living mammalian cells using synthetic self-localizing ligands (SLs). We recently established a chemogenetic SLIPT system, in which any protein of interest fused to an engineered variant of Escherichia coli dihydrofolate reductase, DHFRiK6, can be rapidly and specifically translocated from the cytoplasm to the inner leaflet of the plasma membrane (PM) using a trimethoprim (TMP)-based PM-targeting SL, mDcTMP. The mDcTMP-mediated PM recruitment of DHFRiK6-fusion proteins can be efficiently returned to the cytoplasm by subsequent addition of free TMP, enabling temporal and reversible control over the protein localization. Here we describe the use of this mDcTMP/ DHFRiK6-based SLIPT system for inducing (1) reversible protein translocation and (2) synthetic activation of the Raf/ERK pathway. This system provides a simple and versatile tool in mammalian synthetic biology for temporally manipulating various signaling molecules and pathways at the PM. Key words Protein localization, Self-localizing ligand (SL), SL-induced protein translocation (SLIPT), Plasma membrane, Escherichia coli dihydrofolate reductase, Trimethoprim, Raf, ERK

1

Introduction Mammalian cells are highly compartmentalized systems composed of diverse organelles and membrane domains. In such cellular space, many biological and signaling processes are precisely regulated by protein localization, and mammalian cells elicit various functions by inducing dynamic protein translocation between cellular sites in a spatiotemporally controlled manner [1, 2]. Therefore, the ability to artificially control protein localization with small molecules would be a powerful chemical approach for the study and engineering of mammalian cellular processes. Most reported chemical biology tools for protein localization control are based on the chemically induced dimerization (CID) strategy [3]. The most well-characterized of these, the rapamycin CID system, which

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

237

238

Sachio Suzuki et al.

allows for rapid translocation of a protein of interest via rapamycinmediated heterodimerization of FK506-binding protein (FKBP) and FKBP-rapamycin-binding protein (FRB), has been extensively applied to various biological and bioengineering applications [3]. There has also been recent success in expanding the diversity of chemical dimerization agents that can be used in CID systems, and some of them enable reversible control over the complex formation [4–7]. However, the use of CID systems inevitably requires the coexpression of two constructs in a cell with appropriate stoichiometry to control a single target protein. For mammalian cell engineering applications, a method in which the localization of a protein of interest can be switched using only a small molecule as an extrinsic input (without relying on protein heterodimerization) is highly attractive. The self-localizing ligand-induced protein translocation (SLIPT) methodology is an emerging platform that we devised for this purpose. The SLIPT system enables control of protein localization using synthetic selflocalizing ligands (SLs) (Fig. 1) [8]. SLs are a new class of hybrid synthetic molecules consisting of three parts: (1) a specific smallmolecule ligand for a target protein, (2) a small-molecule localization motif that binds to the intended organelle or subcellular region, and (3) a flexible linker connecting them. Owing to this bifunctional design, following cell entry, the SL can bind to its target protein via the ligand moiety and relocate the protein to the targeting site spontaneously. Because the SLIPT system only uses a single protein component, it can reduce the need for genetic engineering and can be applied to diverse cell lines while obviating the stoichiometry issue. In addition, because of the “single ligandsingle protein” nature, expanding the repertoire of orthogonal SLIPT systems can be readily achieved by using various specific protein-ligand pairs, making the SLIPT approach suitable for multiplexing [9]. By changing the localization motif, we have previously developed SLIPT systems targeting various subcellular sites, including the plasma membrane (PM) [8–10], nucleus [8], microtubules [8], and ER/Golgi membranes (endomembranes) [11], thereby demonstrating the general applicability of the SLIPT approach. In this chapter, we focus on our leading-edge SLIPT system for targeting the PM. The inner leaflet of the PM is a central hub of intracellular signaling networks, where many important signaling molecules and pathways are activated to regulate cellular behavior. The PM-targeted SLIPT system described herein provides a powerful and useful tool for chemically manipulating various PM-localized signaling molecules and for engineering mammalian cell signaling. We constructed the system based on the use of the specific small-molecule trimethoprim (TMP) ligand and Escherichia coli dihydrofolate reductase (eDHFR) pair [12]. In this system, the TMP ligand linked via a flexible linker to a myristoyl-D-Cys (mDc)

Protein Localization and Cell Signaling Control by SLIPT

239

Fig. 1 Principle of the SLIPT methodology. SLs are synthetic small-molecule ligands modified with localization motifs that determine the site of localization. The SL molecule can relocate its target protein from the cytoplasm to the assigned subcellular site/organelle in a “single ligand-single protein” manner. POI: protein of interest

lipopeptide motif (termed mDcTMP [10]) is used as a PM-targeting SL (Fig. 2a, b). The mDc motif undergoes S-palmitoylation of the Cys residue by palmitoyl acyltransferases in cells to localize to the PM (and partially to the Golgi) [10]. Meanwhile, the engineered variant of eDHFR containing a hexalysine (K6) sequence between Asp69 and Asp70 in the internal loop region, termed DHFRiK6, is used as a SLIPT tag for fusion with a protein of interest (Fig. 2a) [13]. Nonengineered (wild-type) eDHFR is recruited not only to the PM but also undesirably to the Golgi by mDcTMP [10]. However, the internal K6 motif of DHFRiK6 stabilizes the PM localization of the mDcTMP-DHFRiK6 complex via electrostatic interactions with anionic lipids, such as phosphatidylserine, that are prevalent on the inner surface of the PM [9, 13]. Consequently, DHFRiK6-fusion proteins can be translocated from the cytoplasm to the PM specifically upon addition of mDcTMP (Fig. 2a) [13]. The protein translocation is rapid and is complete within minutes. Following induction of protein translocation by mDcTMP, the PM-recruited DHFRiK6-fusion proteins can be returned to the cytoplasm by subsequent addition of excess free TMP, enabling temporal control over the reversible protein

240

Sachio Suzuki et al.

Fig. 2 The mDcTMP/DHFRiK6-based PM-specific SLIPT system. (a) Schematic illustration of the reversible translocation of EGFP-DHFRiK6 by mDcTMP and TMP. (b) The chemical structure of mDcTMP. (c) Confocal fluorescence images of HeLa cells expressing EGFP-DHFRiK6 taken before (left), 30 min after incubation with 10μM mDcTMP (center), and 30 min after subsequent incubation with 100μM TMP (right). Scale bar, 20μm. (d) Time course of reversible EGFP-DHFRiK6 translocation by mDcTMP and TMP. To evaluate EGFP-DHFRiK6 translocation, normalized fluorescence intensities of the protein in the cytoplasm were plotted as a function of time. Data are represented as the mean
SD (n ¼ 7 cells)

localization (Fig. 2a). The DHFRiK6 tag can be fused to the N-terminus, C-terminus, or even between two domains of a multidomain protein without the loss of its PM-specificity, offering a high degree of flexibility in the construction of DHFRiK6 fusion proteins. Furthermore, the mDcTMP/DHFRiK6-based SLIPT system has been demonstrated to be applicable to artificially regulate/ activate various signaling molecules, including Raf, RasGEF, PI3K, Tiam1, Gαs, and Gαq at the PM [13]. This new chemogenetic protein recruitment system is, therefore, a versatile and valuable addition to the mammalian chemical and synthetic biology toolkit. As representative examples, here we show the use of the PM-targeted SLIPT system for inducing (1) reversible protein translocation (Fig. 2) and (2) synthetic activation of the Raf/ERK pathway (Fig. 3). The first experiment is intended to show the basic characteristics of the mDcTMP/DHFRiK6 SLIPT system and its reversibility. Here we used HeLa cells expressing enhanced green fluorescent protein (EGFP)-fused DHFRiK6 (EGFP-DHFRiK6) as a

Protein Localization and Cell Signaling Control by SLIPT

241

Fig. 3 Synthetic activation of the cRaf/ERK pathway by the SLIPT system. (a) Schematic illustration of the experimental setup. (b) Confocal fluorescence images of HeLa cells coexpressing EGFP-DHFRiK6-cRaf (above) and ERK-KTR-mKO (below) taken before (left) and 30 min after incubation with 10μM mDcTMP (right). Scale bar, 20μm. (c) To evaluate EGFP-DHFRiK6-cRaf translocation (green), normalized fluorescence intensities of the protein in the cytoplasm were plotted as a function of time. To evaluate ERK activity (orange), normalized ratios of the nuclear to cytosolic fluorescence intensity (C/N ratios) of ERK-KTR-mKO were plotted as a function of time. Data are presented as the mean and SD (n ¼ 5 cells)

model target protein (Fig. 2a). EGFP-DHFRiK6 was distributed throughout the cytoplasm at the initial state, but rapidly and specifically translocated to the PM upon addition of mDcTMP [with a half-maximal translocation time (t1/2) of 2.8 min] (Fig. 2c, d). Following the forward translocation, excess free TMP was added as a competitor ligand, which led to the reverse PM-tocytoplasm relocalization of EGFP-DHFRiK6 [with a half-maximal

242

Sachio Suzuki et al.

reverse translocation time (trev1/2) of 5.8 min] (Fig. 2c, d). Therefore, the present SLIPT system allows researchers to trigger reversible recruitment and release of DHFRiK6-fusion proteins to and from the PM at desired times using mDcTMP and TMP as extrinsic chemical inputs. In the second experiment (Fig. 3), we apply the SLIPT system to manipulate cell signaling. In particular, we focused on activating ERK signaling by controlling the localization of cRaf. cRaf is activated when recruited to the PM, which in turn activates its downstream MEK and ERK [14, 15]. To generate synthetic (mDcTMP-responsive) cRaf whose PM recruitment can be triggered by mDcTMP, we fused EGFP-DHFRiK6 to the N-terminus of a full-length cRaf (EGFP-DHFRiK6-cRaf) (Fig. 3a) and expressed the protein in HeLa cells. To monitor endogenous ERK activity in live-cells, we used a kinase translocation reporter (KTR) for ERK, which was fused to monomeric Kusabira Orange (mKO) (ERK-KTR-mKO) [9]. In this KTR system, the nuclear export of the reporter protein correlates with the kinase activity. Thus, the kinase activity can be quantified by the ratio of the cytosolic to the nuclear fluorescence intensity (C/N ratio) of the reporter protein [15, 16]. Following mDcTMP-induced translocation of EGFP-DHFRiK6-cRaf to the PM, the C/N ratios of ERKKTR-mKO increased significantly, indicating efficient activation of endogenous ERK (Fig. 3b, c). These results indicate that this PM-recruiting SLIPT system can be applied to generate synthetic mammalian cells in which the activity of a specific signaling molecule and pathway can be artificially switched by the mDcTMP input. In the following protocols, we describe the chemical synthesis of mDcTMP (Fig. 4) and its use for the aforementioned SLIPT experiments.

2

Materials

2.1 Solid-Phase Synthesis of mDcTMP

1. Empty polypropylene column with filter, for example, empty PD-10 column (GE Healthcare). 2. Rotary shaker equipped with a vacuum pump, for example, KMS-3 for solid-phase synthesis (Kokusan Chemical). 3. Rink Amide Resin. 4. Fmoc-Lys(ivDde)-OH. 5. Fmoc-8-amino-3,6-dioxaoctanoic acid (Fmoc-8-Adox-OH). 6. Fmoc-D-Cys(Trt)-OH. 7. Myristic acid. 8. Compound 1 (see Fig. 4): This compound is not commercially available and thus needs to be synthesized according to a previous report [17].

Protein Localization and Cell Signaling Control by SLIPT

243

Fig. 4 Synthetic route to mDcTMP

9. 2-(1H-benzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU). 10. 1-Hydroxybenzotriazole monohydrate (HOBt). 11. N,N-Dimethylformamide (DMF), peptide synthesis grade. 12. N,N-Diisopropylethylamine (DIPEA). 13. 20% Piperidine in DMF for Fmoc deprotection. 14. Kaiser test solution: (1) ninhydrin/ethanol (0.5 g/10 mL), (2) phenol/ethanol (40 g/10 mL), and (3) potassium cyanide (KCN)/pyridine (0.2 mL of 1 mM KCN aqueous solution + 9.8 mL pyridine). Solutions (1)–(3) are mixed at 1:1:1 ratio before use.

244

Sachio Suzuki et al.

15. Heating dry bath incubator (settable at 100  C). 16. Dichloromethane (CH2Cl2). 17. 5% Hydrazine monohydrate in DMF for ivDde deprotection. 18. Methanol (MeOH). 19. Trifluoroacetic acid (TFA). 20. 1,2-Ethanedithiol (EDT). 21. Water. 22. Diethyl ether (Et2O). 23. High-performance liquid chromatography (HPLC) machine, for example, Hitach LaChrome system equipped with a UV detector and a YMC-Pack ODS-A (C18) column (5μm, 10  250 nm). 24. Acetonitrile (MeCN), HPLC grade. 25. Lyophilizer (e.g., EYELA FDU-1200). 2.2

Plasmids

1. Mammalian expression plasmid for EGFP-DHFRiK6, pCMVEGFP-DHFRiK6 (see Note 1) [13]. 2. Mammalian expression plasmid for EGFP-DHFRiK6-cRaf, pPBpuro-EGFP-DHFRiK6-cRaf (see Notes 1 and 2) [13]. 3. Mammalian expression plasmid for ERK-KTR-mKO, pPBbsrERK-KTR-mKO (see Notes 1 and 2) [9].

2.3 Cell Culture and Transfection

1. HeLa cells (from Cell Resource Center for Biomedical Research, Institute of Development, Aging and Cancer Tohoku University). 2. Culture medium: Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), penicillin (100 U/mL), and streptomycin (100μg/mL). 3. Trypsin (0.05 w/v%)/EDTA (0.53 mM) solution. Aliquot and store at 20  C. 4. Transfection medium: Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% FBS (no penicillin/ streptomycin). 5. Opti-MEM I reduced serum medium (Gibco). 6. 293fectin (Invitrogen).

2.4 Live-Cell Fluorescence Imaging

1. 35-mm glass-bottomed dish (IWAKI). 2. Imaging medium: Serum- and phenol red-free DMEM supplemented with penicillin (100 U/mL) and streptomycin (100μg/mL). We routinely use a Gibco DMEM (# 21063029).

Protein Localization and Cell Signaling Control by SLIPT

245

3. Confocal laser scanning microscope. We use an IX83/FV3000 confocal laser-scanning microscope (Olympus) equipped with a PlanApo N 60/1.42 NA oil objective (Olympus), a Z drift compensator system (IX3-ZDC2, Olympus), and a stage top incubator (Tokai Hit). Lasers used for excitation are 488 nm for EGFP and 561 nm for mKO. 2.5 SLIPT Experiments

1. mDcTMP: For synthesis, see Subheading 3.1. 2. Trimethoprim. 3. Dimethyl sulfoxide (DMSO).

3

Methods

3.1 Synthesis of mDcTMP

The synthetic scheme is shown in Fig. 4. mDcTMP is synthesized by standard Fmoc-based solid-phase peptide synthesis protocols (see Note 3). Fmoc deprotection is performed with 20% piperidine in DMF at room temperature for 15 min. Amino acid coupling reactions are performed at room temperature with a mixture of Fmocprotected amino acid (3.1 eq.), HBTU (3.0 eq.), HOBt (3.0 eq.), and DIPEA (6.0 eq.) in DMF. All Fmoc deprotection and coupling steps are monitored by the Kaiser test [18]. Unless otherwise stated, all washing procedures are performed with DMF. In the following, we describe our standard protocol using Rink amide resin on a 50μmol scale. 1. Add Rink amide resin (loading: 0.58 mmol/g, 86.2 mg, 50μmol) into a polypropylene column and swell the resin in DMF by shaking the column. 2. By repeating Fmoc deprotection and amino acid coupling reactions, couple Fmoc-Lys(ivDde)-OH, Fmoc-Adox-OH (3), and Fmoc-D-Cys(Trt)-OH to the resin. 3. After Fmoc deprotection, myristoylate the N-terminus using a mixture of myristic acid (3.1 eq.), HBTU (3.0 eq.), HOBt (3.0 eq.), and DIPEA (6.0 eq.) in DMF/CH2Cl2 (1/1). 4. After washing the resin with DMF, deprotect the ivDde group by treatment with DMF containing 5% hydrazine monohydrate. 5. After washing, couple compound 1 (TMP-COOH) to the side chain of the lysine with a mixture of 1 (3.1 eq.), HBTU (3.0 eq.), HOBt (3.0 eq.), and DIPEA (6.0 eq.) in DMF. 6. After washing with DMF, MeOH, and CH2Cl2, dry the resin in vacuo. 7. Deprotect and cleavage the product from the resin by treating the resin with 5 mL of TFA containing 2.5% EDT and 2.5% H2O for 1 h.

246

Sachio Suzuki et al.

8. Collect the filtrate in a 50 mL round-bottomed flask and evaporate the solvent. 9. Precipitate the crude product by adding 10 mL of Et2O. 10. Collect the precipitate by filtration (or centrifugation) and dry in vacuo. 11. Purify the crude product by reversed-phase HPLC using a semipreparative C18 column (a linear gradient of MeCN containing 0.1% TFA and 0.1% aqueous TFA). 12. Lyophilize the collected solution to afford mDcTMP as a white powder [26 mg, 38% (as a mono-TFA salt)]. 1 H NMR (400 MHz, CD3OD): δ/ppm 7.22 (1H, s), 6.56 (2H, s), 4.44 (2H, m), 4.03 (2H, s), 4.00 (4H, s), 3.92 (2H, t, J ¼ 6.2 Hz), 3.80 (6H, s), 3.66 (14H, m), 3.59 (6H, m), 3.46 (6H, m), 3.18 (2H, t, J ¼ 7.0 Hz), 2.84 (2H, m), 2.25 (4H, m), 1.82 (2H, m), 1.71 (2H, m), 1.61 (2H, m), 1.53 (2H, m), 1.40 (2H, m), 1.28 (22H, m), 0.89 (3H, t, J ¼ 6.6 Hz). HRMS (ESI): calculated for [M + H]+, 1252.7221; found, 1252.7181. 3.2 Preparation of mDcTMP Stock Solution

mDcTMP is obtained as a mono-TFA salt after HPLC purification described in Subheading 3.1. We thus use the molecular weight of 1366.6 for calculating the concentration of mDcTMP stock solution (see Note 4). 1. Dissolve mDcTMP in DMSO to make a 10 mM stock solution (see Note 5). 2. Aliquot the stock solution into small volumes (e.g., 5 or 10μL). 3. Store the aliquots at 20  C (see Note 6).

3.3 Preparation of TMP Stock Solution

We calculate the concentration of the TMP solution using the molecular weight of 290.32. 1. Dissolve TMP in DMSO to make a 100 mM stock solution. 2. (Optional) Aliquot the stock solution into small volumes, for example, 5 or 10μL (see Note 7). 3. Store the solution/aliquots at 20  C.

3.4 Reversible SLIPT of EGFP-DHFRiK6

3.4.1 Cell Seeding and Transfection

This protocol describes the procedure for inducing reversible recruitment and release of DHFRiK6-fusion proteins, in this case, EGFP-DHFRiK6, to and from the PM using mDcTMP and TMP. In the following, we describe our standard protocol for transient expression experiments using HeLa cells (see Notes 8 and 9). 1. Seed 1  105 HeLa cells in a 35 mm glass-bottomed dish and culture for 24 h at 37  C in 5% CO2.

Protein Localization and Cell Signaling Control by SLIPT

247

2. On the day of the transfection, dilute 1μg of plasmid DNA (pCMV-EGFP-eDHFRiK6) and 1μL of 293fectin in 50μL Opti-MEM I medium separately. Incubate for 5 min at room temperature. Prepare DNA and 293fectin complex by mixing the solutions and incubating for 20 min at room temperature. 3. Prepare HeLa cells for transfection by replacing the culture medium with 1 mL of transfection medium. Add the DNA and the 293fectin complex prepared above and incubate for 6 h at 37  C in 5% CO2. 4. After 6 h, replace the medium with fresh culture medium and incubate for an additional 18–24 h at 37  C in 5% CO2. 3.4.2 Reversible SLIPT and Fluorescence Imaging

1. Prepare transfected cells for SLIPT and imaging experiments. Remove the culture medium, wash the cells twice with 1 mL of serum-free imaging medium, and add 1 mL of the same imaging medium to the dish (see Note 10). 2. Place and fix the dish on the stage top incubator of a confocal laser-scanning microscope (see Note 11). 3. Acquire fluorescence images of cells by time-lapse imaging. 4. Prepare 1 mL of 20μM mDcTMP-containing imaging medium by adding 2μL of 10 mM mDcTMP stock solution to 1 mL of fresh imaging medium just prior to addition to cells (see Note 12). 5. At the desired timepoint, gently add the 1 mL of 20μM mDcTMP-containing imaging medium to the culture dish (the final concentration of mDcTMP to be 10μM), which initiates the translocation of EGFP-DHFRiK6 from the cytoplasm to the PM. 6. For reverse protein translocation, carefully take 1 mL of the medium from the dish under observation. Add 2μL of 100 mM TMP stock solution to the collected medium to prepare a 1 mL of 200μM TMP-containing imaging medium. 7. Gently add back the TMP-containing medium to the culture dish (the final concentration of TMP to be 100μM) to release EGFP-DHFRiK6 from the PM to the cytoplasm.

3.5 SLIPT-Mediated cRaf Translocation and ERK Activation

3.5.1 Cell Seeding and Transfection

This protocol describes the procedure for artificially inducing ERK activation by recruiting synthetic cRaf (EGFP-eDHFRiK6-cRaf) to the plasma membrane using mDcTMP in HeLa cells (see Notes 9 and 13). 1. Seed 1  105 HeLa cells in a 35 mm glass-bottomed dish and culture for 24 h at 37  C in 5% CO2.

248

Sachio Suzuki et al.

2. On the day of transfection, dilute total 1μg of plasmid DNA (each 0.5μg of pPBpuro-EGFP-DHFRiK6-cRaf and pPBbsrERK-KTR-mKO) and 1μL of 293fectin in 50μL Opti-MEM I medium separately. Incubate for 5 min at room temperature. Prepare DNA and 293fectin complex by mixing the solutions and incubating for 20 min at room temperature. 3. Prepare HeLa cells for transfection by replacing the culture medium with 1 mL of transfection medium. Add the DNA and the 293fectin complex prepared above and incubate for 6 h at 37  C in 5% CO2. 4. After 6 h, replace the medium with fresh culture medium and incubate for an additional 18–24 h at 37  C in 5% CO2. 3.5.2 SLIPT-Mediated Activation of the cRaf/ERK Pathway

1. Prepare transfected cells for SLIPT and imaging experiments. Remove the culture medium, wash the cells with 1 mL of serum-free imaging medium, and add 1 mL of the same imaging medium to the dish (see Note 10). 2. Incubate the cells for at least 1 h for starvation (see Note 14). 3. Place and fix the dish on the stage top incubator of a confocal laser-scanning microscope (see Note 11). 4. Acquire dual-color fluorescence images of cells by time-lapse imaging. 5. Prepare 1 mL of 20μM mDcTMP-containing imaging medium by adding 2μL of 10 mM mDcTMP stock solution to a 1 mL of fresh imaging medium just prior to addition to cells (see Note 12). 6. At the desired timepoint, gently add the 1 mL of 20μM mDcTMP-containing imaging medium to the culture dish (the final concentration of mDcTMP to be 10μM), which initiates translocation of EGFP-DHFRiK6-cRaf from the cytoplasm to the PM. 7. Monitor endogenous ERK activity by observing the nuclear export of ERK-KTR-mKO (see Note 15).

4

Notes 1. All expression plasmids were generated using standard cloning techniques and the In-Fusion cloning system (Clontech). We used pEGFP-N1 (Clontech) as a vector backbone for pCMVEGFP-DHFRiK6. We also used pPBpuro [9] and pPBbsr [9, 19] as vector backbones for pPBpuro-EGFP-DHFRiK6cRaf and pPBbsr-ERK-KTR-mKO, respectively. We will provide these expression plasmids upon request. 2. This pPB-based plasmid can also be used to generate stable cell lines using a piggyBac transposon system [20].

Protein Localization and Cell Signaling Control by SLIPT

249

3. mDcTMP is not commercialized at the moment. We will provide samples of mDcTMP upon request. 4. The concentration of mDcTMP stock solution can be determined spectrophotometrically by measuring the absorbance at 294 nm (A294) in 1% triethylamine-containing DMSO using the molar extinction coefficient of TMP, 6.8  103 M1 cm1. 5. The mDcTMP powder is hygroscopic and sometimes becomes a transparent gel or a droplet. The powder also often tends to adhere to the wall of the tubes. 6. To avoid air-oxidation of the Cys side chain of mDcTMP, do not repeat freeze and thaw the aliquots. 7. Because TMP is stable in solution, aliquoting the TMP stock solution is optional. 8. This protocol is intended to describe our standard methods for SLIPT experiments using transiently transfected cells. However, we often establish stable cell lines expressing DHFRiK6tagged proteins and other reporter proteins using the piggyBac transposon system [20]. 9. We have experimentally confirmed that this mDcTMP-based SLIPT system is also operational in other cell lines, including NIH3T3, COS-7, and PC12 cells [10, 13], demonstrating the general applicability of the system to various mammalian cell lines. As additional information, we have verified that the present system also works in live nematodes, namely Caenorhabditis elegans (unpublished results). 10. Because mDcTMP binds to albumin in serum, we recommend using serum-free media for SLIPT experiments. When mDcTMP is used in serum-containing medium, protein translocation becomes significantly more inefficient and slower than in serum-free conditions. 11. We usually perform SLIPT experiments and imaging at 37  C. However, mDcTMP-mediated protein translocation may proceed even at room temperature. 12. We routinely dilute a DMSO stock solution of mDcTMP in serum-free medium first and then add the partially diluted medium to the culture dish to be a final concentration of 10μM mDcTMP. Because mDcTMP tends to be oxidized gradually in the medium, we recommend making a dilution of mDcTMP stock solution just before the addition to the culture dish. 13. We have so far experimentally verified that the present SLIPT system is generally applicable to control various signaling molecules, including RasGEF (leading to Ras activation), PI3K

250

Sachio Suzuki et al.

(phosphatidylinositol 3,4,5-trisphosphate [PI(3,4,5)P3] production), Tiam1 (Rac activation and lamellipodium induction), Gαs (cAMP production), and Gαq (Ca2+ signaling) [13]. We will provide expression plasmids used for these experiments upon request. 14. Cells are serum-starved for at least 1 h to suppress the basal activity of cRaf and ERK. 15. For quantification of the endogenous ERK activity in cells, we perform a region-of-interest (ROI) analysis and measure the ratio of the cytosolic to the nuclear fluorescence intensity (C/N ratio) of ERK-KTR-mKO using the Fiji distribution of ImageJ [21].

Acknowledgments We thank Dr. Akinobu Nakamura (ExCELLS, National Institutes of Natural Sciences) for his contribution to the development and application of the mDcTMP/DHFRiK6-based SLIPT system. This work was supported by JSPS Grants-in-Aid for Scientific Research (KAKENHI): grant nos. 15H03835, 15H05949 “Resonance Bio,” 18H02086, and 18H04546 and 20H04706 “Chemistry for Multimolecular Crowding Biosystems” (to S.T.). S.S. acknowledges scholarship support from the Hirota Scholarship Society and the SUNBOR Scholarship from the Suntory Foundation for Life Sciences. Conflicts of Interest: S.S., T.Y., and S.T. are coinventors on a patent application related to this work. Y.H. declares no competing interests. References 1. Teruel MN, Meyer T (2000) Translocation and reversible localization of signaling proteins: a dynamic future for signal transduction. Cell 103:181–184 2. Hurley JH, Meyer T (2001) Subcellular targeting by membrane lipids. Curr Opin Cell Biol 13:146–152 3. DeRose R, Miyamoto T, Inoue T (2013) Manipulating signaling at will: chemicallyinducible dimerization (CID) techniques resolve problems in cell biology. Pflu¨gers Arch 465:409–417 4. Feng S, Laketa V, Stein F, Rutkowska A, MacNamara A, Depner S, Klingmu¨ller U, Saez-Rodriguez J, Schultz C (2014) A rapidly reversible chemical dimerizer system to study lipid signaling in living cells. Angew Chem Int Ed 53:6720–6723

5. Liu P, Calderon A, Konstantinidis G, Hou J, Voss S, Chen X, Li F, Banerjee S, Hoffmann JE, Theiss C, Dehmelt L, Wu YW (2014) A bioorthogonal small-molecule-switch system for controlling protein function in live cells. Angew Chem Int Ed 53:10049–10055 6. Ballister ER, Aonbangkhen C, Mayo AM, Lampson MA, Chenoweth DM (2014) Localized light-induced protein dimerization in living cells using a photocaged dimerizer. Nat Commun 5:5475 7. Foight GW, Wang Z, Wei CT, Greisen PJ, Warner KM, Cunningham-Bryant D, Park K, Brunette TJ, Sheffler W, Baker D, Maly DJ (2019) Multi-input chemical control of protein dimerization for programming graded cellular responses. Nat Biotechnol 37:1209–1216

Protein Localization and Cell Signaling Control by SLIPT 8. Ishida M, Watanabe H, Takigawa K, Kurishita Y, Oki C, Nakamura A, Hamachi I, Tsukiji S (2013) Synthetic self-localizing ligands that control the spatial location of proteins in living cells. J Am Chem Soc 135:12684–12689 9. Nakamura A, Oki C, Kato K, Fujinuma S, Maryu G, Kuwata K, Yoshii T, Matsuda M, Aoki K, Tsukiji S (2020) Engineering orthogonal, plasma membrane-specific SLIPT systems for multiplexed chemical control of signaling pathways in living single cells. ACS Chem Biol 15:1004–1015 10. Nakamura A, Oki C, Sawada S, Yoshii T, Kuwata K, Rudd AK, Devaraj NK, Noma K, Tsukiji S (2020) Designer palmitoylation motif-based self-localizing ligand for sustained control of protein localization in living cells and Caenorhabditis elegans. ACS Chem Biol 15:837–843 11. Nakamura A, Katahira R, Sawada S, Shinoda E, Kuwata K, Yoshii T, Tsukiji S (2020) Chemogenetic control of protein anchoring to endomembranes in living cells with lipid-tethered small molecules. Biochemistry 59:205–211 12. Miller LW, Cai Y, Sheetz MP, Cornish VW (2005) In vivo protein labeling with trimethoprim conjugates: a flexible chemical tag. Nat Methods 2:255–257 13. Hatano Y, Suzuki S, Nakamura A, Yoshii T, Atsuta-Tsunoda K, Aoki K, Tsukiji S (2020) A chemogenetic platform for controlling plasma membrane signaling and synthetic signal oscillation. bioRxiv. https://doi.org/10.1101/ 2021.03.16.435568 14. Aoki K, Kumagai Y, Sakurai A, Komatsu N, Fujita Y, Shionyu C, Matsuda M (2013) Stochastic ERK activation induced by noise and

251

cell-to-cell propagation regulates cell densitydependent proliferation. Mol Cell 52:529–540 15. Maryu G, Matsuda M, Aoki K (2016) Multiplexed fluorescence imaging of ERK and Akt activities and cell-cycle progression. Cell Struct Funct 41:81–92 16. Regot S, Hughey JJ, Bajar BT, Carrasco S, Covert MW (2014) High-sensitivity measurements of multiple kinase activities in live single cells. Cell 157:1724–1734 17. Ando T, Tsukiji S, Tanaka T, Nagamune T (2007) Construction of a small-moleculeintegrated semisynthetic split intein for in vivo protein ligation. Chem Commun 47:4995–4997 18. Kaiser E, Colescott RL, Bossinger CD, Cook PI (1970) Color test for detection of free terminal amino groups in the solid-phase synthesis of peptides. Anal Biochem 34:595–598 19. Komatsu N, Aoki K, Yamada M, Yukinaga H, Fujita Y, Kamioka Y, Matsuda M (2011) Development of an optimized backbone of FRET biosensors for kinases and GTPases. Mol Biol Cell 22:4647–4656 20. Yusa K, Rad R, Takeda J, Bradley A (2009) Generation of transgene-free induced pluripotent mouse stem cells by the piggyBac transposon. Nat Methods 6:363–369 21. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A (2012) Fiji: an opensource platform for biological-image analysis. Nat Methods 9:676–682

Chapter 15 Engineering Hydrogel Production in Mammalian Cells to Synthetically Mimic RNA Granules Hideki Nakamura Abstract Recent studies revealed the biological significance of dynamic multicomponent assemblies of biomolecules inside living cells. Protein and nucleic acid assemblies are biomolecular condensates or non–membranebound organelles that have attracted increasing attention. Synthetic tools that manipulate the dynamic assembly/disassembly process of the structures are useful in elucidating both biophysical mechanisms of their assembly/disassembly and physiological roles of the condensates. In this report, general protocols to form and observe synthetic polymer-based condensates in living cells are described using the tool iPOLYMER. Taking advantage of the modular design of the tool, both chemical and light stimuli can induce formation of synthetic condensates inside living cells, which are observed by laser-scanning confocal microscopy. The experimental design described herein should help those who conduct experiments on synthetic manipulation of biomolecular condensates using iPOLYMER and other tools for synthetic manipulation of condensates. Technical notes for using iPOLYMER, including basic protocols of chemicalor light-inducible dimerization techniques (CID/LID), choice of proper control experiments, and advantages/disadvantages are also presented. Key words Synthetic biology, Inducible dimerization, iPOLYMER, Hydrogel, Biomolecular condensates, Stress granules

1

Introduction

1.1 A Synthetic Biological Approach to Intracellular Assembly of Multiple Biomolecules

A novel class of intracellular structures, biomolecular condensates, has recently attracted significant attention in the field of cell biology [1, 2]. Biomolecular condensates are intracellular structures without any surrounding membranes to define the boundary and typically consist of multiple biomolecule species such as proteins and nucleic acids. These condensates are found in a huge variety of biological contexts, ranging from essentially physiological processes such as gene expression to severely pathogenic situations including neurodegenerative diseases, underpinning the biological importance of these structures [3]. One remarkable feature of biomolecular condensates is that the mechanism by which they are assembled, often referred to as “phase separation,” involves

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

253

254

Hideki Nakamura

multivalent interactions between multiple constituents of the granules to enable the molecules to undergo phase transitions that distinguish their assemblies from ambient environments [4]. A basic understanding of biomolecular condensate assembly is thus fundamentally important for establishing the cell biology of this novel subcellular structure. In a previous study, we developed a synthetic biology tool, iPOLYMER (induced production of ligand(light)-yielded multivalent enhancers) that manipulates the sol–gel phase transition of two polypeptide species in living cells [5]. The tool was developed based on inducible dimerization paradigms, chemically inducible dimerization (CID) and light-inducible dimerization (LID). CID and LID are techniques that induce association between two specific proteins by chemical administration and light irradiation, respectively (Fig. 1a). Inspired by a seminal study demonstrating that two tandemly connected binding domains can form condensates in vitro and in cellulo [6], we designed a novel tool, iPOLYMER and iPOLYMER-LI by tandemly connecting the proteins used in CID/LID paradigms (Fig. 1a). Upon chemical or light stimulus, iPOLYMER can form peptide-based hydrogel condensates in living

a

SspB

rapamycin

FRB FKBP

Stress granule protein

b

TIA-1 N

C

RRM domains

iLID

dark

Prion-related domain (PRD)

Intrinsically disordered

YFP-SspB x 6

YFP-FKBP x 5

RNA binding

Phase Transition/Separation

mCherry-iLID x 6

CFP-FRB x 5

rapamycin

Blue light

Blue light

dark

cyto-YFP-FKBP x 5

TIA1 RRMYFP-SspB x 6

TIA1 RRMCFP-FRB x 5

TIA1 RRMmCherry-iLID x 6

rapamycin

Blue light Synthetic Stress Granule “Analogue”

Fig. 1 Design of iPOLYMER based on inducible dimerization techniques. (a) Chemically or light-inducible versions of iPOLYMER (lower panels) were designed based on inducible dimerization techniques, chemically inducible dimerization (CID) and light-inducible dimerization (LID) (upper panels), respectively. Note that LID is readily reversible, whereas CID is practically irreversible inside living cells. This difference is directly reflected in iPOLYMER tools. (b) Functionalizing iPOLYMER condensates to synthetically mimic stress granules. RNA-binding domains (RRM domains) from a stress granule marker protein, TIA-1 (upper panel), were fused to iPOLYMER polypeptides (lower panels)

Synthetic iPOLYMER Hydrogel Formation in Living Cells

255

cells in a spatiotemporally controlled manner. In the study, we also demonstrated that the tool can reconstitute or mimic the specific molecular constituents of a typical biomolecular condensate, stress granules (Fig. 1b) [5]. Fusing RNA-binding domains from a stress granule marker protein, TIA-1, to an iPOLYMER peptide (s) facilitated specific recruitment of RNA-binding proteins that are markers of stress granules to the artificial condensates, or stress granule analogs, as described in the following protocols. Stress granules are a widely studied example of biomolecular condensates. They are macroscopic condensates in the cytosol that consist of multiple proteins and mRNAs in cells under stress conditions [7]. Interestingly, a wide variety of stresses including UV irradiation, high temperature, osmotic pressure and virus infections lead to the formation of the granule, suggesting potentially important physiological and pathophysiological roles. However, the precise mechanism of stress granule formation under such diverse stress conditions remains unresolved. A major biological outcome of stress granule formation is inhibition of translation of the mRNAs sequestered in the granules, although the roles or importance of the translation inhibition in the context of stress survival are still under debate [8, 9]. Furthermore, recent reports indicate pathogenic roles of abnormally stabilized stress granules in the context of neurodegenerative diseases including amyotrophic lateral sclerosis (ALS) [10]. Synthetic reconstitution of “artificial” stress granules in living cells under stress-free conditions thus offers an attractive and unprecedented opportunity to gain insights into mechanisms and biological outcomes of stress granule formation. 1.2 Growing Palette of Synthetic Biology Tools in the Field

In accordance with growing biological interests into biomolecular condensates, tools that synthetically form “artificial” condensates in living cells have been developed recently [11]. These tools are distinct from each other in their design strategies, which eventually leads to differences in the physical and biological properties of the formed condensates. Known mechanisms of protein-dependent condensate formation can be classified into three groups: concentration-dependent condensation of intrinsically disordered domains (IDRs) (class I); oligomerization of folded proteins (class II); and tandem binding domains and multimerizing domains with readily defined stoichiometries (class III) [11]. Synthetic biology tools that manipulate condensate formation combine one or more of the three mechanisms. A comprehensive overview of the mechanisms exploited in various tools that have gained physiological insights have been reported elsewhere [11, 12]. Appropriate synthetic biology tools should be adopted, according to the scope of the experiment.

256

Hideki Nakamura

1.3 Advantages and Disadvantages of iPOLYMER

The major advantage of iPOLYMER over other synthetic biology tools described above is its simple and straightforward design. iPOLYMER is the only inducible tool that involves only the class III (tandem binding domains) mechanism. Physicochemical mechanisms underlying condensation of IDRs (class I) are actively investigated and have yet to be fully elucidated. Oligomerization of proteins (class II) are relatively well studied but lack well-defined stoichiometry. The sole dependency of iPOLYMER on the class III mechanism is thus ideal for experiments that require fine-tuning of stoichiometry or valence numbers underlying protein association. This simple design principle has enabled evaluation of valence number dependency of condensate formation in a previous study by simply changing the number of dimerizing domains in the peptide chains [5]. The second advantage of iPOLYMER is the in vitro characterization of the inducible sol-gel phase transition of the peptide chains, using exactly the same proteins as expressed in living cells. Although phase-separated liquid droplets and hydrogels formed by purified IDRs have been characterized extensively, in vitro characterization of the synthetic tools that are based on IDRs have not been reported. Additionally, characterization of condensates as hydrogels is often dependent on the slower turnover rate of material measured by FRAP, which is rather qualitative and not rigorous. In vitro characterization of an iPOLYMER hydrogel demonstrated its ability to hold water, a clearly defined pore size and elasticity [5]. Therefore, this tool should aid research where the physical character of the condensate as a hydrogel plays an important role. However, in vitro characterization was performed only for chemically inducible iPOLYMER and not for light-inducible iPOLYMER-LI because of technical difficulties. As a recently developed tool, iPOLYMER has a number of disadvantages. The biggest challenge in using iPOLYMER is its modest penetrance and large phenotypic deviation among cell populations. Rapid formation of iPOLYMER condensates in living cells (within tens of minutes) requires high-level expression of the two polypeptide chains, resulting in condensates in a relatively small fraction of cells. Therefore, it is challenging to combine iPOLYMER with bulk cell population assays such as western blotting. Even among cells with similar expression levels for both peptide chains, phenotypes vary considerably, in terms of numbers, sizes, and formation kinetics of the condensates. Thus, it is essential to prepare appropriate control experiments and make careful data interpretation to avoid drawing illusory conclusions from iPOLYMER experiments. Noteworthy, the modest penetrance of iPOLYMER condensate formation sharply contrasts with the robust performance of CID and LID, which iPOLYMER/iPOLYMER-LI are based on. CID and LID are among a few paradigms in the field of

Synthetic iPOLYMER Hydrogel Formation in Living Cells

257

synthetic biology that have been used widely and have been successful in many applications with high fidelity and robustness [13, 14]. Since CID and LID are the only intermolecular association mechanisms used in iPOLYMER and iPOLYMER-LI, respectively, it is intriguing that formation of iPOLYMER condensates show considerable variance between cells. One potential reason for this seeming discrepancy is the lack of heterotypic interactions between multiple components, which plays an important role in condensate formation by another synthetic biology tool, OptoDroplets [15]. The simple design principle of iPOLYMER, which is the major advantage, could also represent its major disadvantage by providing only modest robustness. Moreover, the stability of polypeptide chains may also be a limiting aspect of the tool. Polypeptide chains used in iPOLYMER inevitably contain repeat sequences, which may affect the stability of the proteins. Based on these considerations, further improvement of the tool especially the penetrance and robustness is being undertaken. 1.4 Requirement for Appropriate Control Experiments

2

In interpreting iPOLYMER experiment results, it is essential to design control experiments that are suitable for the scope of the research. Because of the modest penetrance of the tool, phenotypes are often below the detection limit in the majority of cells. Rational and quantitative comparison between experiment and control conditions is a vital discussion point. Stimuli used in iPOLYMER tools (rapamycin or blue light) may also affect biological processes. Typical control experiments for rapamycin-inducible iPOLYMER are listed in Table 1. Here, the functional domain (FD) is assumed to be fused to the N-termini of CFP-FRBx5 and YFP-FKBPx5. For the stress granule reconstitution experiment described below, FD corresponds to RNA-binding domains, RRM domains, from a stress granule protein, TIA-1 (Fig. 1b). Control conditions should be properly selected from the listed conditions based on the hypothesis tested in each experiment.

Materials Condensate formation by iPOLYMER has been confirmed in multiple cell lines, including COS-7, HEK293T, HeLa, and NIH3T3 cells. Cell lines COS-7 and HEK293T are more suitable because high-level overexpression is advantageous. COS-7 cells are especially feasible for several reasons: (1) COS-7 cells express transduced plasmids at high levels; (2) they are highly adhesive and robust; and (3) the morphology is thin and flat in the cytoplasm, which aids observation of the formed condensates in a single optical section by confocal microscopy. As a reference, three-dimensional observation of iPOLYMER condensates in a single COS-7 cell and in a HEK293T cell are shown in Fig. 2. The following protocols use

258

Hideki Nakamura

Table 1 Candidates for appropriate control experiments Rapamycin or DMSO Comments

Plasmids Experiment

l

Control

l

FD-CFP-FRBx5/ FD-YFP-FKBPx5

FD-CFP-FRBx5/ FD-YFP-FKBPx5 l FD-CFP-FRBx5/ cyto-YFP-FKBPx5 l Cyto-CFP-FRBx5/ FD-YFP-FKBPx5 l Cyto-CFP-FRBx5/ cyto-YFP-FKBPx5 l FD-CFP-FRBx5/ FD-YFP-FKBP l FD-CFP-FRB/ FD-YFP-FKBP

Rapamycin

N/A

DMSO

Requirement for condensate formation

Rapamycin

Requirement for larger amount of FD. Could be experiment conditions

Rapamycin

Requirement for FD

Rapamycin

Requirement for condensate formation. Effects of rapamycin and FD overexpression are considered

COS-7

HEK293T cyto-CFP-FRB x 5 / cyto-YFP-FKBP x 5

cyto-CFP-FRB x 5 / cyto-YFP-FKBP x 5

z

3D Side View

x

10 µm

10 µm 10 µm

10 µm

2D Images

Fig. 2 Three-dimensional distribution of iPOLYMER condensates in distinct cell lines. Administering rapamycin to a COS-7 cell (left panels) and to a HEK293T cell (right panels) induced the formation of iPOLYMER condensates. Blue and green signals are from cyto-CFP-FRBx5 and cyto-YFP-FKBPx5 polypeptides, respectively. Three-dimensional distributions of iPOLYMER condensates imaged by a confocal microscope are shown in upper panels as a 3D rendering presentation viewed from the sides of the cells (3D Side View). Lower panels are images at a single optical section in each cell (2D images). Scale bars: 10 μm

Synthetic iPOLYMER Hydrogel Formation in Living Cells

259

COS-7 cells as an experimental system, but other cell lines can be used with similar protocols by adjusting several parameters including cell densities and transfection details. Condensates formed by iPOLYMER are readily observable with a conventional epifluorescence microscope, which takes advantage of the fluorescent proteins, enhanced cyan fluorescent protein (ECFP) and enhanced yellow fluorescent protein (EYFP), which are part of the polypeptide chains. Typical filter combinations for these fluorescence proteins readily facilitate iPOLYMER experiments. If only green/red wavelengths are available, fluorescent proteins can be replaced by, for example, EGFP and mCherry. iPOLYMER-LI can also be used in combination with epifluorescence microscopes if an on-stage patterned blue-light illuminator is available or spatially uniform light irradiation is sufficient. In the latter case, illuminating the sample with blue light used for EGFP excitation can induce iPOLYMER-LI formation. Assuming that relatively high subcellular spatial resolution is required, the following method adopted for laser-scanning confocal microscopy can be employed. Apart from higher threedimensional spatial resolution of condensate observations, confocal microscopy provides subcellularly-patterned light stimulus to iPOLYMER-LI without any external illuminators, as well as feasible combinations with protein dynamics measurements such as FRAP experiments. Condensate formation by iPOLYMER takes tens of minutes with varied kinetics in different cells. Intensive imaging over the period typically leads to considerable photobleaching of fluorescent proteins. Therefore, to observe the phenotype or biological outcomes of iPOLYMER condensate formation, it is often feasible to observe cells that are fixed at a certain time after stimulus. In the following, protocols for inducing condensate formation in the incubator and those for immunostaining will be described, taking an example of immunostaining against stress granule markers. 2.1 Materials for Cell Culture

1. General disposables and instrumentation used for general mammalian cell cultures. 2. COS-7 Cercopithecus aethiops kidney cells. 3. 35 mm glass-bottom tissue culture dishes or chamber coverslips. 4. Poly-D-lysine for coverslip coating. 5. Dimethylsulfoxide (DMSO) for poly-D-lysine stock solution. Protocols for stock solution preparation will be described in the following. 6. Culture medium: Dulbecco’s modified Eagle medium (DMEM) with high glucose and supplemented with sodium pyruvate, L-glutamine, fetal bovine serum (FBS), and penicillin–streptomycin.

260

Hideki Nakamura

2.2 Materials for Transfection

1. Transfection reagent. 2. DNA plasmids used in iPOLYMER/iPOLYMER-LI. Constructs have been described in a previous report [5]. Name and plasmid number in Addgene of each construct, if available, are described in Table 2. Plasmids mentioned in the following sections are available either from Addgene, or upon request. 3. Serum-free medium such as Opti-MEM.

2.3 Materials for Live-Cell Fluorescence Imaging

1. Laser-scanning confocal microscope. In the following, a confocal microscope supplemented with a 405 nm diode laser and a 470–670 nm white laser is used. 2. Optional: Stage-top incubator for long-term observation of condensate formation in living cells. 3. Extracellular medium: for example, phenol red–free DMEM (DMEM( phenolred)) or Hank’s Balanced Salt Solution (HBSS). 4. Rapamycin stock solution in DMSO. Protocols for preparation are described in the following.

Table 2 DNA plasmids used in the current protocols

Name of plasmids

Comments

Addgene ID

Tom20-ECFP-FRB

Used for CID calibration (Subheading 3.1.2)

N/A

EYFP-FKBP

Used for CID calibration (Subheading 3.1.2)

N/A

cyto-CFP-FRBx5

Forms nonfunctionalized iPOLYMER condensates with cyto-YFPFKBPx5 (Subheadings 3.1.3 and 3.1.4)

103776

cyto-YFP-FKBPx5

Forms iPOLYMER condensates or stress granule analogs (Subheadings 3.1.3 and 3.1.4)

103777

pMT2 TIA1 RRM-CFPFRBx5

Forms functionalized iPOLYMER stress granule analogs with cytoYFP-FKBPx5 (Subheadings 3.1.3 and 3.1.4)

103783

Tom20-EYFP-iLID Used for LID calibration (Subheading 3.2)

N/A

mCherry-SspB

Used for LID calibration (Subheading 3.2)

N/A

YFP-SspBx6

Forms nonfunctionalized iPOLYMER-LI condensates with mCherry- 103778 iLIDx6 (Subheadings 3.2 and 3.3)

mCherry-iLIDx6

Forms nonfunctionalized iPOLYMER-LI condensates with YFP-SspBx6 (Subheadings 3.2 and 3.3)

103779

TIA1 RRM-YFPSspBx6

Forms functionalized iPOLYMER-LI stress granule analogs with TIA1 RRM-mCherry-iLIDx6 (Subheadings 3.2 and 3.3)

103781

TIA1 RRM-mCherryiLIDx6

Forms functionalized iPOLYMER-LI stress granule analogs with TIA1 RRM-YFP-SspBx6 (Subheadings 3.2 and 3.3)

103782

Synthetic iPOLYMER Hydrogel Formation in Living Cells

2.4 Materials for Immunostaining Against Stress Granule Markers and a Nonmarker

261

1. 16% paraformaldehyde (PFA). 2. Phosphate buffered saline (PBS). 3. Skim milk for blocking. 4. Sodium (meta)arsenite for inducing stress granule formation. 5. Antibodies listed in Table 3. 6. A custom-built illuminator for blue light stimulation in the incubator (design shown in Fig. 3). For details of the design, see Note 1.

3

Methods The following methods are used to optimize the formation of synthetic stress granule analogs and their nonfunctionalized control condensates in living cells by iPOLYMER/iPOLYMER-LI. Experimental notes containing further tips are also provided.

3.1 Induce iPOLYMER Condensate Formation in Living Cells by Chemical Stimulus 3.1.1 Preparing Rapamycin Stock

1. Weigh ~9.14 mg of rapamycin (MW 914.17) into a 10 mL conical tube. Add DMSO to make a 10 mM rapamycin stock solution. Scale the volume of added DMSO according to the amount of rapamycin weighed (see Note 2). 2. Mix thoroughly. Make sure no precipitant is left at the bottom of the conical tube. 3. Aliquot 10 μL of the 10 mM rapamycin stock into 1.5 mL tubes. 4. Add 990 μL DMSO to a 10 mM stock tube. Keep the rest of the 10 mM stock tubes at 20  C for later use. Mix thoroughly. This gives a 100 μM rapamycin stock solution in DMSO. 5. Aliquot the 100 μM rapamycin stock solution in 1.5 mL tubes by 1 μL. The volume can be adjusted according to the conditions of use. Store the stock at 20  C. Repetitive freeze–thaw cycles should be avoided.

3.1.2 Validating the Stock by CID-Dependent Protein Translocation

Although straightforward, preparation of the rapamycin stock occasionally fails. To avoid this, it is recommended to check the quality of the stock solution by a simple imaging experiment. Rapamycin-inducible translocation of proteins is an ideal benchmark because this is experimentally feasible [16]. The protocol for mitochondria-targeted translocation using two DNA plasmids, Tom20-ECFP-FRB and EYFP-FKBP, is described. Fluorescent proteins can be replaced by proteins with suitable wavelengths that are detected by the microscope of choice (e.g., EGFP and mCherry).

262

Hideki Nakamura

Table 3 Lists of antibodies used in the protocol Primary Antibodies

Stress granule markers

Nonmarker

Antigens

Manufacturer

Cat. #

Host

Clonal

Comments

PABP-1

Sigma-Aldrich

Mouse

Mono

Clone 10E10

G3BP1

Santa Cruz

Mouse

Mono

N/A

eIF4G

Santa Cruz

Rabbit

Poly

H-300

eIF3b

Santa Cruz

041467 sc81940 sc11373 sc16377

Goat

Poly

discontinued (replaced by sc-137214)

Ribosomal ImmunoVision HPOP-antigen 0100

Human Autoantibody N/A

Secondary Antibodies Comments Goat anti-mouse IgG, Alexa Fluor 594 conjugated

N/A

Goat anti-rabbit IgG, Alexa Fluor 594 conjugated

N/A

Rabbit anti-goat IgG, Alexa Fluor 594 conjugated

N/A

Goat anti-human IgG, Alexa Fluor 647 conjugated

Relatively unconventional, and could be hard to find In the current protocol, product from Jackson ImmunoResearch (AB_2337885) was used

Goat anti-mouse IgG, Alexa Fluor 647 conjugated

N/A

Goat anti-rabbit IgG, Alexa Fluor 647 conjugated

N/A

Rabbit anti-goat IgG, Alexa Fluor 647 conjugated

N/A

1. The day before transfection, seed COS-7 cells at 1–2  105 cells per dish in a 35 mm glass-bottom culture dish. Coverslip coating is not strictly required if the surface of the coverslip is treated for the cell culture. 2. Check the confluency of the cells under a microscope on the following day. A 70–80% confluency is optimal for transfection. 3. Add 200 μL serum-free medium in a sterile 1.5 mL tube.

Synthetic iPOLYMER Hydrogel Formation in Living Cells

263

Fig. 3 Design of a custom-made blue LED illuminator. Top-down view (upper panel) and side view (lower panel) are shown as schematic drawings. Electric wires are connected to LED drivers and power source, which are not depicted. The entire illuminator part shown in the figure was placed in an incubator, whereas the LED drivers are kept outside. The lid blocks the blue LED light to avoid potential effects on other cells in the incubator

4. Add 3 μg Tom20-ECFP-FRB plasmid and 1.5 μg EYFP-FKBP plasmid to the tube. The ratio between the two plasmids can be further adjusted according to the results (see Note 3). Mix well by pipetting. 5. Add 4.5 μL of ViaFect transfection reagent and mix thoroughly by pipetting. Let the DNA mixture incubate for 5–20 min at room temperature. 6. Add the whole volume of DNA mixture to a glass-bottom dish in a drop-wise manner. Gently shake the dish to mix the medium and incubate at 37  C, 5% CO2. 7. The next day, image the dish under an inverted fluorescence microscope. Before imaging, replace the culture medium by the extracellular medium of your choice (see Note 4). Use 900 μL of the extracellular medium for a 35 mm glass-bottom dish. The optical setup for ECFP/EYFP imaging is described in Note 5.

264

Hideki Nakamura

8. Before imaging, add 100 μL extracellular medium to a tube containing 1 μL of the rapamycin DMSO stock to dilute ~100 times. Mix well by pipetting. 9. The ECFP signal should be localized at mitochondria, showing a typical tubular morphology, whereas the EYFP signal should be diffusely distributed across the entire cytoplasm. Pick a cell/ cells with relatively high expression of ECFP and relatively modest expression of EYFP to observe sharp translocation upon rapamycin administration (see Note 3). Signals should not be saturated if quantitative analysis is required. In particular, the EYFP signal should be modest when considering the intensity increase at mitochondria upon stimulus. 10. Set parameters for time-lapse imaging in the application. Intervals of 10–30 s between frames are appropriate because the translocation typically takes place within 1–2 min. The entire imaging will be complete in 5–10 min if successful. It is usually favorable to use the autofocus function, as significant focal drift tends to take place. 11. Start time-lapse imaging. At a certain point, add 100 μL of the rapamycin solution prepared in step 8 to the dish by micropipetting (see Note 6). Do not touch the sample or the stage, as this could cause significant movement of the field of view, and thus disrupt cell tracking. 12. EYFP-FKBP should quickly translocate to mitochondria within a minute following rapamycin administration. If translocation is not observed, it is highly probable that the rapamycin stock was not prepared properly. Make fresh DMSO stocks from rapamycin powder and try the protocol again. Rapamycin stocks are usually stable over several months when stored at 20  C. 3.1.3 Live-Cell Imaging of iPOLYMER Condensate Formation in Living Cells

1. Prepare a poly-D-lysine solution for coating coverslip in a 35 mm glass-bottom culture dish (see Note 7). Add 50 mL sterile water to a bottle of 5 mg lyophilized poly-D-lysine hydrobromide to give a 0.1 mg/mL poly-D-lysine solution. Thoroughly dissolve poly-D-lysine by gently swirling the bottle or pipetting up and down. 2. Add 200 μL of the poly-D-lysine solution to a well of a glassbottom culture dish (see Note 8). This should cover the whole coverslip area with a thin layer of solution. Let the glass-bottom dish incubate for 30 min at room temperature in the culture hood. This process can be done in the incubator or overnight. 3. Aspirate the poly-D-lysine solution. Add 1 mL sterile water and aspirate the whole volume. Repeat the wash process three times. Try not to touch the coated surface with the tip of the aspirator during the procedure. Let the surface dry for longer than 5 min in the culture hood.

Synthetic iPOLYMER Hydrogel Formation in Living Cells

265

4. Seed 0.5–1  105 cells per dish and incubate overnight. Cell density is kept lower than Subheading 3.1.2 because imaging is performed 48 h after transfection in the current protocol. Cell density can be adjusted according to the results. 5. Transfect cells as described in steps 3–6 of Subheading 3.1.2 using 3 μg pMT2 TIA1 RRM-CFP-FRBx5, 3 μg cyto-YFPFKBPx5, and the transfection reagent of choice (see Note 9). The amount of the reagent should be optimized according to the manufacturer’s instructions. As a nonfunctionalized control condition, use cyto-CFP-FRBx5 and cyto-YFP-FKBPx5 instead, as suggested in Table 1. The amount of DNA should be adjusted according to the results. 6. Optional: The next day, replace DNA-containing culture medium with fresh culture medium. This procedure can be performed 8 h after transfection. Changing the medium may be dispensable if cells look healthy. 7. Approximately 48 h after transfection image cells under the microscope (see Note 10). Since iPOLYMER condensate formation takes place over tens of minutes, it is recommended to use a stage-top incubator for the experiment. Replace culture medium with 900 μL extracellular medium of choice. Optical parameters should be set in a similar manner to step 9 in Subheading 3.1.2. Adjust laser power and/or scan speed such that signals are not saturated in cells with the highest expression levels. 8. Pick multiple positions to be imaged. The number of positions could vary depending on the time required for scanning and frame interval. Intervals of 1–5 min are a good starting point when considering the kinetics of formation of the iPOLYMER condensate. Ensure that the positions are not too many for the intended interval. There are two options when picking positions. Option one: observe fluorescence through the eyepiece to quickly find cells with high expression levels. Option two: use confocal microscopy to find positions. Using the automatic focus stabilization functions is strongly recommended to compensate for differences in focal planes among positions and for focal drift over time. Another tip that helps compensate for focal drift is to use a pinhole size larger than the theoretically optimal value (Airy unit one) for imaging. Although sacrificing confocality, this also helps to grasp clearly the distribution of condensates over the entire cell, especially in morphologically flat cells such as COS-7 cells. 9. Dissolve 3–5 μL 100 μM rapamycin stock solution in 100 mL extracellular medium. Note that the concentration of rapamycin is several times higher than the protocol in Subheading 3.1.2 (see Note 11). This is to improve hydrogel formation

266

Hideki Nakamura

by increasing the chances of crosslinking between two iPOLYMER peptide chains. The conventional concentration of 100 nM also works. 10. Start time-lapse imaging of multiple positions selected in step 8. The total duration of imaging should be set to 30–90 min. 11. After several frames, add the rapamycin solution prepared in step 9 in a similar manner to that used in step 11 of Subheading 3.1.2. Use of the on-stage incubator demands additional care because direct access to the pipette is not available. In this case, pause the time-lapse imaging, carefully open the lid of the incubator and add rapamycin. The whole transmitted light illuminator part may have to be carefully moved away to allow for better access to the incubator. Alternatively, timelapse imaging before and after rapamycin administration can be performed separately, as independent sessions. 3.1.4 Immunostaining of Fixed Cells with iPOLYMER Condensates

Specific recruitment of stress granule markers to functionalized iPOLYMER condensates can be demonstrated by immunostaining of the condensates with antibodies against stress granule markers and a nonmarker RNA-binding protein, P-antigen (Table 3 in Subheading 2.4). The immunostaining method is described below. 1. Prepare the sample as described in steps 1–6 of Subheading 3.1.3. To confirm the functionalized iPOLYMER condensates as “stress granule analogs,” cyto-CFP-FRBx5/cyto-YFPFKBPx5 represent a good control, as this is the only straightforward control condition that gives condensate formation. As a positive control, cells should be treated with 0.5% w/v sodium arsenite for 30–60 min. This should lead to stress granule formation in most cells. Both rapamycin and sodium arsenite treatment should be done in normal culture medium in the incubator. 2. Break an ampoule of 16% PFA and move the content to a 15 mL conical tube. 3. Add 1 mL of 16% PFA to another 15 mL conical tube. Add 3 mL PBS and mix thoroughly to give 4% PFA for fixation. The amount of PFA solution can be scaled according to the number of samples. Seal the tube of 16% PFA with parafilm and store the tube at 4  C. 4. Remove culture medium from the sample glass-bottom culture dish. Add 1 mL of 4% PFA. 5. After 5 min, remove PFA and wash with PBS three times. 6. Prepare blocking solution. Make 0.4% w/v skim milk in PBS (see Note 12).

Synthetic iPOLYMER Hydrogel Formation in Living Cells

267

7. Replace culture medium with 1 mL 0.4% w/v skim milk or alternative blocking solution. Incubate the sample at room temperature for 1 h. Blocking at 4  C overnight is also acceptable. Avoid light by covering the sample with a lid to prevent unnecessary photobleaching of fluorescent proteins. 8. Wash three times with PBS. 9. Replace PBS with 0.4% w/v skim milk in PBS containing a primary antibody. A dilution of 1:500 is suitable for antibodies against PABP-1, G3BP1, eIF4G and eIF3b. The antibody against ribosomal P-antigen is used at 1:100 dilution. Let the sample sit for 1 h at room temperature. Avoid light by covering the sample with a lid. 10. Wash three times with PBS. 11. Replace PBS with a secondary antibody. As ECFP and EYFP are fused to iPOLYMER peptide chains, red or far-red dye-conjugated secondary antibodies are feasible, according to the availability of excitation/emission wavelengths on the microscope. A dilution of 1:1000 works for the current protocol. 12. Wash three times with PBS. Samples can be stored at 4  C for at least a week. For longer preservation, samples should be prepared on an independent coverslip, followed by mounting the coverslip on a slide glass. 13. Image samples with a fluorescence microscope. Signals of stress granule markers (PABP-1, G3BP1, eIF4G, eIF3b) should accumulate at “stress granule analogs” made by iPOLYMER under the experiment conditions, whereas the ribosomal P-antigen should not (see Note 13). The pattern is the same as endogenous stress granules induced by sodium arsenite. Conventional iPOLYMER condensates without RRM domains (formed by cyto-CFP-FRBx5/cyto-YFP-FKBPx5) should not accumulate any of the RNA-binding proteins. 3.2 Live-Cell Imaging of iPOLYMER-LI Condensate Formation

1. Prepare the sample as described in steps 1–6 in Subheading 3.1.3. Use TIA1 RRM-YFP-SspBx6 and TIA1 RRM-mCherry-iLIDx6 (see Note 14). After transfection, avoid light by covering the sample with a lid to prevent induction of condensate formation by environmental light. If necessary, a change in medium should be carried out under minimal light, ideally under red light. 2. Image the sample with a confocal microscope. Avoid exposure to light before imaging. Use a 514 nm laser at the lowest intensity to excite EYFP, as the wavelength can induce some association between SspB and iLID. If 514 nm or a similar wavelength is not available, avoiding the observation of EYFP fluorescence could be a good idea.

268

Hideki Nakamura

3. Set parameters for light stimulation. The FRAP function of the application usually provides a feasible measure to perform optical stimulation. In the current setup, 514 nm and 587 nm lasers were used to excite EYFP and mCherry, respectively. The 488 nm laser was used to induce SspB-iLID dimerization. In typical FRAP function of confocal microscopes, there are three parameters that need to be adjusted; laser intensity, frame rate and number of frames. Use a modest intensity that is similar to the level typically used to excite GFP (below 1%), as higher power lasers cause serious photobleaching of EYFP. Intervals of 5–20 s are suitable for the experiment because dissociation between SspB-iLID takes place approximately over a minute (see Note 15). The entire duration that cells are stimulated should be longer than 30 min. As a reference sample, cells expressing Tom20-EYFP-iLID and mCherry-SspB work well. This should result in an EYFP signal on the surface of mitochondria outer membranes, and a mCherry signal over the entire cell. The 488 nm stimulus induces translocation of mCherry toward mitochondria, which is readily recognized by the typical morphology of mitochondria. 4. Start imaging using the parameters described in step 3. Use of automatic focus stabilization functions is strongly recommended, as in Subheading 3.1.3. Representative images from a single cell are shown in Fig. 4. 3.3 Immunostaining of Fixed Cells with Light-Induced iPOLYMER-LI Condensates

1. Prepare samples as in Subheading 3.2. As a negative control condition, use nonfunctionalized iPOLYMER-LI constructs, YFP-SspBx6 and mCherry-iLIDx6 with the identical protocol. Plan experiments carefully in advance, so that the number of samples is sufficiently large for immunostaining conditions including various control conditions. 2. Forty-eight hours after transfection, sterilize the LED illuminator (see Subheading 2.4) by irradiating with UV light in the culture hood for 30 min. 3. Place the illuminator in the incubator. Use a lid to cover the entire illuminator to prevent any effects of blue light on other cells in the incubator. 4. Place sample on illuminator. Make sure the area of the culture is placed in the middle of the cylinder light guide. Place the lid prepared in step 3 on top. 5. Turn on the light. Emission of LED is sufficiently strong to be seen by light leaking through the gap between the lid and illuminator. 6. Prepare 4% PFA for fixation during the stimulus as in steps 2 and 3 of Subheading 3.1.4.

Synthetic iPOLYMER Hydrogel Formation in Living Cells

ROI1

ROI2

Overlaid images YFP-SspBx6/mCherry-iLIDx6

Before

ROI1 Light ON

269

ROI2 Light ON

OFF OFF

Fig. 4 Local and reversible condensate formation by iPOLYMER-LI. Overlaid images of fluorescence signals from iPOLYMER-LI polypeptides are shown at different time points. Local illumination of blue light was performed in ROI1 and then in ROI2 in a spatially restricted manner, as described in the protocol. Local and reversible formation of condensates are clearly observed in the stimulated regions. Scale bar: 10 μm. (The data is modified from Ref. 5)

7. After 60 min, take the entire illuminator out of the incubator. Caution: the LED is still on and extremely bright, so never look into the illuminator from the top during the fixation procedure. Take off the lid, and quickly replace medium with 4% PFA. Put the lid back on and let the sample incubate on the illuminator with the LED on for 5 min. 8. After 5 min, turn off the illuminator. Take off the lid and perform immunostaining as described in steps 8–13 of Subheading 3.1.4. Alexa Fluor 647-conjugated secondary antibodies should be used. Alexa Fluor 405 or 430-conjugated secondary antibodies could be options, according to the availability of the wavelengths. Representative results of immunostaining against a stress granule marker, G3BP1, and a nonmarker, ribosomal P-antigen, are shown in Fig. 5.

270

Hideki Nakamura

iPOLYMER-LI polypeptides EYFP mCherry

a

G3BP1 immunostaining

Overlaid

Non-functionalized iPOLYMER-LI (YFP-SspBx6/mCherryiLIDx6)

Functionalized iPOLYMER-LI B

(TIA1 RRM -YFP-SspBx6/ TIA RRM-mCherry-iLIDx6)

A

TIA1 RRMYFP-SspBx6

b

TIA1 RRM- Ribosomal P-antigen mCherry-iLIDx6 immunostaining

Overlaid

A B

Normalized signal along the line

c

G3BP1

Ribosomal P-antigen

(stress granule marker, (a))

(Non-marker, (b))

TIA1 RRM-YFP-SspBx6 TIA1 RRM-mCherry-iLIDx6 Immunostaining

Fig. 5 Specific recruitment of stress granule markers to functionalized iPOLYMER-LI condensates. (a) Representative results of immunostaining with an anti-G3BP1 antibody are shown for a cell expressing nonfunctionalized iPOLYMER-LI polypeptides (upper panels) and a cell expressing iPOLYMER-LI polypeptides fused to RNA-binding domains (RRM domains) from TIA-1 (lower panels). Images of EYFP fluorescence (green images), mCherry fluorescence (red images) and Alexa647 fluorescence (G3BP1 immunostaining, monochrome images) are shown, along with overlaid images of the three images with Alexa647 signals presented as blue. G3BP1 colocalization with iPOLYMER-LI condensates clearly requires RRM domains. In the overlaid image of functionalized iPOLYMER-LI condensates, the line used for the line-scan profile shown in (c) is shown as a white broken line (A to B). Scale bars: 10 μm. (b) Immunostaining of RRM domain-functionalized iPOLYMER-LI condensates with an anti-ribosomal P-antigen antibody. The nonstress granule marker protein is not accumulated at the synthetic condensates, unlike G3BP1. The line used for the line-scan profile shown in (c) is shown as a white broken line (A to B) in the overlaid image. Scale bar: 10 μm. (c) Line-scan fluorescence intensity profiles along broken lines in (a) and (b) are plotted as normalized signal intensities. Signals of immunostaining (blue lines) are colocalized to EYFP (green lines) and mCherry (red lines) signals from iPOLYMER-LI polypeptides in the G3BP1 immunostaining result (left panel), whereas colocalization is not clearly observed in the ribosomal P-antigen result (right panel). (Data is modified from Ref. 5)

Synthetic iPOLYMER Hydrogel Formation in Living Cells

4

271

Notes 1. In a previous report [5], the illuminator (depicted in Fig. 3) was kindly provided by Sergi Regot (The Johns Hopkins University). The illuminator used 3 W blue-light LED for stimulation. Briefly, six LEDs are located at the bottom of a cardboard box, connected to the electric wires. Surrounding each LED is a cardboard cylinder with aluminum foil glued to its inner surface with each LED centrally positioned. The top of the cardboard box was replaced by a transparent cellophane sheet to allow photostimulation of the sample (glass-bottom culture dish or chambered coverslips) placed on top. The height of the cardboard box that defines the distance between the light source and the sample was 6 cm. A single-output constant current LED driver connected by electric wires controlled each LED. The entire illuminator was placed in a CO2 incubator where the photostimulation process was performed. Note that other electronic circuits including LED drivers were placed outside the incubator. 2. Rapamycin stocks should be prepared at two concentrations (10 mM and 100 μM in the current protocol). Here, the final concentration that rapamycin is present in the sample is 100–500 nM. Given the Kd between FKBP and rapamycin (0.2 nM) and that between FKBP-rapamycin and FRB (12 nM) [17], the concentration used ensures a predominantly dimer population. A too dilute rapamycin DMSO stock in aqueous solution causes precipitation of the drug, which is readily visible as a white precipitant. Therefore, the final stock should be prepared as 1000 or higher-concentration in DMSO. Rapamycin is a potent inhibitor of the mechanistic target of rapamycin (mTOR) C1 pathway, with a subnanomolar IC50 in some cases. The final concentration and duration of rapamycin administration should thus be determined with care to ensure that the activity of rapamycin as an inhibitor does not interfere with appropriate data interpretation. 3. The ratio between the number of proteins overexpressed at the intended target locations (anchor unit) and those proteins that undergo translocation (translocating unit) should be carefully optimized. Using the same amount of plasmids, the translocating unit protein (EYFP-FKBP in the current case) is usually cytosolic and highly overexpressed when compared with that of the anchor unit protein (Tom20-ECFP-FRB), which is targeted to a subcellular location. Even with saturating concentrations of rapamycin, this should result in the translocation of a relatively small fraction of translocating unit proteins. Typically, using more plasmid encoding the anchor unit to that encoding the translocating unit improves the situation. Using

272

Hideki Nakamura

a 2:1 to 10:1 ratio (anchor unit –translocating unit) of the plasmids works in most experiments. Note that the same amount of the two DNA plasmids are used in other subheadings, because similar concentrations of both polypeptide chains are preferable. 4. The extracellular medium should be selected according to the experimental setup. For example, short-term experiments at room temperature are less susceptible to the lack of nutrients and metabolites such as glucose, essential amino acids, and sodium pyruvate. In long-term experiments greater than an hour, phenol red–free culture medium supplemented with constituents listed above is preferable in combination with CO2 incubation in a stage-top incubator at 37  C. COS-7 cells are relatively resistant against suboptimal temperatures, and many experiments can be performed without on-stage incubation, although potential effects of the ambient environment have to be carefully considered in data interpretation. 5. If available, 445/458 nm and 514 nm lasers are optimal to excite ECFP and EYFP, respectively. If a 514 nm laser is not available, it is challenging to image ECFP and EYFP in an independent manner. Fluorescence proteins should be replaced, for example, by mCherry in those cases. ECFP was excited by a 405 nm laser in the current method. The 413–508 nm and 518–670 nm emissions were collected as ECFP and EYFP fluorescence, respectively, with GaAsPavalanche hybrid detectors. 6. Access to the sample on the microscope stage is usually limited by the presence of the condenser of the transmitted light illuminator unit. However, the rapamycin solution can be added in a drop-wise manner by using a P200 micropipette through the gap between the condenser and the sample, as shown in Fig. 6 Note that the light is turned off during the actual experiment. A perfusion system should make an ideal alternative, which has not been tested because a large amount of rapamycin is required. 7. COS-7 cells are highly adhesive and coating is not strictly required in many experiments. However, coating is recommended in the protocol, as formation of iPOLYMER condensates in living cells seems to weaken cell-substrate adhesion in some cell lines in our preliminary results. Poly-L-lysine coating also works for the experiment. 8. The current protocol assumes 35 mm glass-bottom culture dishes with 14 mm-diameter wells. When the number of samples is large, a multiwell chambered coverslip (e.g., SCC-008, Matsunami) would be more favorable. Reagents and the number of cells should be scaled to the culture area for chambered coverslips.

Synthetic iPOLYMER Hydrogel Formation in Living Cells

273

Fig. 6 Manual addition of rapamycin to the imaging sample. Rapamycin was added through the gap between the condenser and the sample by using a micropipette, as shown in the image. Alternatively, the entire condenser part can be moved backward to improve access to the sample; however, this procedure often leads to the movement of the sample, thereby resulting in the loss of ability of trackingthe observed cells. Note that the addition of rapamycin is performed in the dark in actual experiments

9. In the current protocol, TIA1 RRM-CFP-FRBx5 is subcloned into the pMT2 plasmid, which suppresses endogenous stress granule formation caused by overexpression of exogenous proteins [18]. This should lower the risks of illusory results by inhibiting endogenous stress granules from recruiting iPOLYMER polypeptide chains. 10. In general, a two-day incubation after transfection tends to result in higher levels of expression than a one-day incubation. 11. In a previous publication, the final concentration used was typically 333 nM [5]. Higher rapamycin concentrations of 300–500 nM inevitably increase the DMSO concentration in the medium, which may affect the biological processes. One solution is to prepare higher concentrations of the rapamycin stock or prepare a vehicle control with the same plasmids and concentration of DMSO. 12. Alternatively, 1% w/v BSA in PBS or a 5% w/v serum solution can be used for blocking. In the latter case, serum from the same species as the host organism of the secondary antibody should be used. 13. The signal from ribosomal P-antigen immunostaining should not accumulate at stress granules or stress granule analogs. The lack of ribosomal protein accumulation suggests the lack of mRNA translation at the synthetic condensates.

274

Hideki Nakamura

14. In the current protocol, having RRM domains fused to only one of the two polypeptide chains did not result in significant recruitment of stress granule markers. Additionally, use of pMT2 plasmids (as in Note 10) is ideal in the protocol, although it is yet to be tested. 15. In light-inducible iPOLYMER-LI, the frame rate should directly affect apparent kinetics of condensate formation, as shown as a schematic in Fig. 7. This is because of the dissociation between iLID and SspB in the absence of a blue light stimulus.

Light stimulus ON

Pattern of Blue Light Irradiation

Extent of Condensate Formation

“True” kinetics

High frame rate

Low frame rate detection limit

time Timing of Blue light stimuli

Timing of Image acquisition

”Apparent” kinetics of condensate formation

Fig. 7 Effects of frame rates on apparent kinetics of iPOLYMER-LI condensate formation. In the current setup, cells are illuminated with blue light immediately before acquisition of each data time point. Consequently, association between iPOLYMER-LI polypeptides is reversed during each interframe period, as depicted by the blue and magenta solid lines for lower and higher frame rates, respectively. Compared with the “true” kinetics of condensate formation with a continuous stimulus (black solid line), the dissociation of the iPOLYMER-LI polypeptides results in a reduction in the “apparent” kinetics, which are shown as broken lines. The effect should be more prominent in the result for the lower frame rate (blue lines) when compared with that of the higher frame rate (magenta lines)

Synthetic iPOLYMER Hydrogel Formation in Living Cells

275

Acknowledgments The author thanks the following researchers for sharing essential resources: Itaru Hamachi for providing the environment for experiment and research; Takanari Inoue for related constructs and reagents; Nancy Kedersha for stress granule-related constructs and antibodies; and Sergi Regot for sharing his custom-made LED illuminator. Appreciation is extended to Albert A. Lee for his vast contribution to the research, Nancy Kedersha for insightful discussions on stress granules, all the lab members of the Takanari Inoue Lab for experimental assistance and fruitful discussions, and lab members of the Itaru Hamachi Lab for insightful discussions and technical assistance. The author thanks Edanz Group (https:// en-author-services.edanzgroup.com/ac) for editing a draft of the manuscript. References 1. Banani SF, Lee HO, Hyman AA, Rosen MK (2017) Biomolecular condensates: organizers of cellular biochemistry. Nat Rev Mol Cell Biol 18:285–298 2. Wheeler RJ, Hyman AA (2018) Controlling compartmentalization by non-membranebound organelles. Phil Trans R Soc B 373:20170193. https://doi.org/10.1098/ rstb.2017.0193 3. Shin Y, Brangwynne CP (2017) Liquid phase condensation in cell physiology and disease. Science 357:eaaf4382. https://doi.org/10. 1126/science.aaf4382 4. Weber SC, Brangwynne CP (2012) Getting RNA and protein in phase. Cell 149:1188–1191 5. Nakamura H, Lee AA, Afshar AS, Watanabe S, Rho E, Razavi S, Suarez A, Lin Y-C, Tanigawa M, Huang B, DeRose R, Bobb D, Hong W, Gabelli SB, Goutsias J, Inoue T (2018) Intracellular production of hydrogels and synthetic RNA granules by multivalent molecular interactions. Nat Mater 17:79–89 6. Li P, Banjade S, Cheng H-C, Kim S, Chen B, Guo L, Llaguno M, Hollingsworth JV, King DS, Banani SF, Russo PS, Jiang Q-X, Nixon BT, Rosen MK (2012) Phase transitions in the assembly of multivalent signalling proteins. Nature 483:336–340 7. Anderson P, Kedersha N (2002) Stressful initiations. J Cell Sci 115:3227–3234 8. Protter DSW, Parker R (2016) Principles and properties of stress granules. Trends Cell Biol 26:668–679

9. Ivanov P, Kedersha N, Anderson P (2019) Stress granules and processing bodies in translational control. Cold Spring Harb Perspect Biol 11:a032813. https://doi.org/10.1101/ cshperspect.a032813 10. Zhang K, Daigle JG, Cunningham KM, Coyne AN, Ruan K, Grima JC, Bowen KE, Wadhwa H, Yang P, Rigo F, Taylor JP, Gitler AD, Rothstein JD, Lloyd TE (2018) Stress granule assembly disrupts nucleocytoplasmic transport. Cell 173:958–971 11. Nakamura H, DeRose R, Inoue T (2019) Harnessing biomolecular condensates in living cells. J Biochem 166:13–27 12. Bracha D, Walls MT, Brangwynne CP (2019) Probing and engineering liquid-phase organelles. Nat Biotechnol 37:1435–1445 13. DeRose R, Miyamoto T, Inoue T (2013) Manipulating signaling at will: chemicallyinducible dimerization (CID) techniques resolve problems in cell biology. Pflugers Arch 465:409–417 14. Niu J, Ben Johny M, Dick IE, Inoue T (2016) Following optogenetic dimerizers and quantitative prospects. Biophys J 111:1132–1140 15. Riback JA, Zhu L, Ferrolino MC, Tolbert M, Mitrea DM, Sanders DW, Wei M-T, Kriwacki RW, Brangwynne CP (2020) Compositiondependent thermodynamics of intracellular phase separation. Nature 581:209–214 16. Komatsu T, Kukelyansky I, McCaffery JM, Ueno T, Varela LC, Inoue T (2010) Organelle-specific, rapid induction of

276

Hideki Nakamura

molecular activities and membrane tethering. Nat Methods 7:206–208 17. Banaszynski LA, Liu CW, Wandless TJ (2005) Characterization of the FKBP.rapamycin.FRB

ternary complex. J Am Chem Soc 127:4715–4721 18. Kedersha N, Anderson P (2007) Mammalian stress granules and processing bodies. Meth Enzymol 431:61–81

Chapter 16 AgDD System: A Chemical Controllable Protein Aggregates in Cells Yusuke Miyazaki Abstract There are increasing evidence and growing interest in the relationship between protein aggregates/phase separation and various human diseases, especially neurodegenerative diseases. However, we do not entirely comprehend how aggregates generate or the clearance network of chaperones, proteasomes, ubiquitin ligases, and other factors interact with aggregates. Here, we describe chemically controllable systems compose with a genetically engineered cell and a small drug that enables us to rapidly induce protein aggregates’ formation by withdrawing the small molecule. This trigger does not activate global stress responses induced by stimuli, such as proteasome inhibitors or heat shock. This method can produce aggregates in a specific compartment and diverse experimental systems, including live animals. Key words Chemical biology, Protein aggregates, Chaperones, Proteasome, Destabilizing domain, Phase separation

1

Introduction Proteins are involved in every aspect of cellular function and homeostasis, playing enzymatic, structural, or signaling roles that are typically made possible when the polypeptide folds into a particular three-dimensional structure and conformation. However, proteins do not always assume ideal conformations during protein synthesis, and are readily damaged by various environmental stresses such as heat and oxidative stress. These misfolded proteins often lose the ability to perform their intended function, and in some cases, damaged proteins can sometimes acquire new pathological functions. One significant consequence would be protein aggregates. We understand that protein aggregates are often associated with neurodegenerative diseases and are also strongly associated with aging [1–4]. However, we do not entirely understand the mechanisms of protein aggregation, including how protein aggregates are formed and cleared, which proteins and others

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

277

278

Yusuke Miyazaki

participated in the process, or why oligomers and aggregates are toxic to cells [5, 6]. There are currently three popular methods to generates cellular aggregates: heat shock, proteasome inhibitions, and genetic perturbations. Although many researchers generate protein aggregates by heat shock and proteasome inhibitions, we have to admit that these perturbants induce global cellular stress responses unrelated to protein aggregation and are also extreme stressors themselves [7–12]. Others have studied aggregates using genetic perturbations, including polyQ-fused mutant huntingtin (Htt) proteins; however, these genetic methods lack the control of the timing of protein aggregates and their size [13–15]. Here, to address these shortcomings and advance our understanding of how protein aggregates affect protein homeostasis in cells, we developed a novel dose-dependent system to rapidly create protein aggregates with precise spatial and temporal control. We previously developed a general technique allowing for conditional control of protein stability using cell-permeable ligands and engineered proteins [16–18]. These proteins are called destabilizing domains (DDs), and the ligand can be used as a switch to control the folding state of the DD. In other research publications, DDs have demonstrated that the procedures are not inducing global cellular stress [19]. Therefore, compared with previous methods used to investigate protein aggregates, the genetically engineered AgDD system has several advantages. It induces aggregates immediately and reversible, it can be precisely regulated both spatially and temporally, and it has fewer off-target effects and relatively low toxicity for cells. Thus, this system should enable detailed investigations of the mechanisms of the cellular response to protein aggregation. Deepening the understanding of protein aggregations should lead to a potential therapeutic strategy is to prevent the formation of protein aggregates by modulating protein homeostasis for treating neurodegenerative and other age-related diseases, therefore the research and developments for protein aggregates/ condensates are one of the hot current research area [2, 20, 21] (see Fig. 1 [22]).

Fig. 1 Schematic representation of inducing aggregated proteins inside cells using an AgDD

AgDD System: A Chemical Controllable Protein Aggregates in Cells

2

279

Materials Prepare all solutions using ultrapure water and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing waste materials.

2.1 Establishing AgDD Cell Lines

1. Culture medium: DMEM supplemented with 10% heatinactivated donor bovine serum, 2 mM glutamine, 100 U/ mL penicillin and 100 μg/mL streptomycin. Store at 4  C. Or depends on cell line used DMEM supplemented with 10% heat-inactivated fetal bovine serum, 2 mM glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin. Store at 4  C. 2. NIH 3T3 cell line (ATCC CRL-1658). Store at liquid nitrogen or 80  C. 3. HEK293 cell line. Store at liquid nitrogen or

80  C.

4. 1 μM Shield-1(S1) ligand in Ethanol. Store at Note 1).

20  C (see

5. 6-well cell plates. 6. 10 cm dish plates. 7. FKBP-derived AGDD-sfGFP cloned Piggybac Transposon System vector pB (System Biosciences). Store at 20  C. (AgDD construct is available on Addgene (Plasmid #78289). 8. Piggybac Transposase vector (System Biosciences). Store at 20  C. 9. TransIT-LT1 (Mirus). Store at 4  C (see Note 2). 10. Trypsin. Store at

20  C.

11. BD FACSAria. 2.2 Analyzing AgDD Cell Lines

1. FKBP(with F36V mutation) protein (350–500 μM stock in PBS buffer with 4% glycerol). Store at 80  C. 2. BD FACSCalibur. 3. FlowJo software. 4. Nikon Eclipse Ti fluorescence microscope. 5. μManager. 6. 8-well NuncLab-Tek Chambered Coverglass coated with polylysine. 7. Zeiss Axioskop 2 epifluorescence microscope equipped with a QICAM FAST 1394 digital CCD camera.

280

Yusuke Miyazaki

2.3 Recombinant Protein Purification: BL21 Cells Expressing His6-FKBP(F36V) Protein

1. BL21 cells. Store at

80  C.

2. LB media. 3. PBS. 4. Isopropyl β-D-1-thiogalactopyranoside. Store at 4  C. 5. lysozyme. Store at 4  C. 6. Triton X-100. 7. Ni-NTA resin. Store at 4  C. 8. Wash buffer: 50 mM NaH2PO4, 300 mM NaCl, 20 mM imidazole at pH 8. Store at 4  C. 9. Elution buffer: 50 mM NaH2PO4, 300 mM NaCl, 250 mM imidazole at pH 8. Store at 4  C. 10. Dialysis buffer: PBS with 4% glycerol. Store at 4  C. 11. Nanodrop (Thermo Fisher).

3

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

1. The NIH3T3 cell lines are cultured in culture medium. For the HEK 293 cell lines and U2OS cell lines are cultured with 10% heat-inactivated fetal bovine serum, 2 mM glutamine, 100 U/ mL penicillin, and 100 μg/mL streptomycin.

3.2 Preparing Purified Recombinant Protein for the Experiments

1. BL21 cells expressing His6-FKBP(F36V) protein [23] is cultured in LB media at 37  C until optical density at 600 nm reached between 0.5 and 0.7.

3.1

2. Protein expression is induced with 0.5 mM isopropyl β-D-1thiogalactopyranoside at 20  C for overnight, then cells are harvested by centrifugation at 3600  g for 20 min at 4  C. 3. Cells are lysed in PBS buffer supplemented with 1 mg/mL lysozyme and left on ice for 30 min. 4. After sonication, frozen at 80  C for overnight. Triton X-100 is added to 1%, and after 30 min incubation on ice the solutions are centrifuge at 44,000  g for 30 min at 4  C. 5. The soluble fraction is bound to Ni-NTA resin at 4  C for 1 h. The resin is washed several times with wash buffer until we do not observe protein level by nanodrop, then protein is eluted with elution buffer. 6. Eluted solutions is combined and dialyzed into dialysis buffer and stored at 80  C.

AgDD System: A Chemical Controllable Protein Aggregates in Cells

3.3 Establishing AgDD Cell Lines

281

1. Nascent cells are plated at 10  104 cells per well of a 6-well plate a day prior to transfection. 2. The cells are cotransfected with plasmids of a pB vector and PiggyBac Transposase vector following standard protocols using TransIT-LT1. 3. After the cells are cultured in growth media contains 1 μM S1 ligand for about a week to allow stable integration. 4. The transfected cells are trypsinized and resuspended in culture media before flow cytometry experiments (see Note 3). 5. Sort the cells by BD FACSAria sorter, and sort for high GFP level cells which is expressing high amounts of AgDD-GFP fusion protein (see Note 4).

3.4 Inducing Protein Aggregates

1. After the sorting, we maintain the stabilizing S1 ligand in culture media at all times to avoid cells expose to unfolded proteins. 2. To remove the S1 ligand from media containing S1 1μM, we directly add purified FKBP(F36V) protein (350–500 μM stock) in cultured media to achieve a final protein concentration of 3.5–5 μM (see Note 5).

3.5 Analyzing the Protein Aggregates in FACS

1. At the desired time points, cells are trypsinized and resuspended in culture media before undergoing FACS. 2. For sorting we use BD FACSAria. For analytical flow cytometry cells we use BD FACSCalibur with no less than 10,000 events represented. 3. Data are analyzed using FlowJo (see Note 6) (Fig. 2).

3.6 Analyzing the Protein Aggregates Under Fluorescence Microscopy

1. Time-lapse imaging is performed with a Nikon Eclipse Ti fluorescence microscope using μManager (https://www. micro-manager.org/). 2. Aggregates induced cells are prepared on an 8-well NuncLabTek Chambered Coverglass (Thermo Scientific) coated with polylysine (Invitrogen). 3. During time-lapse experiments, the same conditions are maintained including a temperature of 37  C and a CO2 concentration of 5%. 4. Live cell images are captured on a Zeiss Axioskop 2 epifluorescence microscope equipped with a QICAM FAST 1394 digital CCD camera. (see Note 7) (Fig. 3).

282

Yusuke Miyazaki

Fig. 2 Flow cytometry analysis showing GFP signal pulse width versus height of HEK cells stably expressing AgDD before and after drug removal. Cells with aggregates are defined as the fraction of the total population within the purple gate (reproduced from ref. 22)

4

Notes 1. Shield-1 could be purchased through several vendors. If organic chemistry setup is available in the lab, synthesize by following the paper [24]. 2. Although we use and described the reagent, other transfection reagents should work fine. 3. Use media including Shield-1 through all the preparation. Also, put cells on the ice immediately to keep cells healthy and not inducing aggregations. 4. Although it depends on transfection efficiency and other factors, try to sort “very-high” GFP signal first such as top 1% populations. We observed signal dependency in this techniques, thus more aggregated cells with high signal populations compared to low signal populations. 5. Without utilizing purified FKBP (F36V) protein, we could remove the ligand by washing cells twice with warmed culture media then incubated with the fresh media. However, we have

AgDD System: A Chemical Controllable Protein Aggregates in Cells

283

Fig. 3 (a) Time-lapse images of representative HEK cells stably expressing AgDD. Top: cells cultured in media with S1 throughout the experiment; middle: cells cultured in media with S1 withdrawn at 0 h and bottom: cells cultured in media with S1 withdrawn from 0 to 1 h then readministered from 1–8 h. Scale bars, 20 μm. (b) Early time-lapse images of representative HEK cells stably expressing AgDD following S1 withdrawal. Scale bar, 20 μm (reproduced from ref. 22)

observed quite small but still some RNA response during that washing method since changing media could be inducing physical stress response to cells. 6. Defined the aggregated cells populations from FACS data. Cells with aggregates displayed narrowly distributed fluorescent signals compared with cells that did not have these punctate structures [25]. 7. This technique is also available for immunofluorescence. For example, we have tested these fixations procedure: Following fixation in PBS with 4% paraformaldehyde (Ted Pella) for 20 min, cells were permeabilized using PBS with 0.2% Triton X (Sigma) for 15 min and incubated in 3% BSA (SigmaAldrich) in PBS for 60 min. Antibody incubation was carried out at 4  C overnight followed by Alexa Fluor–conjugated secondary antibodies for 1 h at room temperature (1:1000, Life Technologies) and Hoechst 33342 (1:10000, Life Technologies).

284

Yusuke Miyazaki

Acknowledgments This work was supported by the NIH (GM073046 and P50 GM107615), the Ellison Medical Foundation (AG-SS-2573-10), and the Stanford Discovery Innovation Fund. Y.M. was supported by the Nakajima Foundation. References 1. Powers ET, Morimoto RI, Dillin A, Kelly JW, Balch WE (2009) Biological and chemical approaches to diseases of proteostasis deficiency. Annu Rev Biochem 78:959–991 2. Douglas PM, Dillin A (2010) Protein homeostasis and aging in neurodegeneration. J Cell Biol 190(5):719–729 3. Morimoto RI (2011) The heat shock response: systems biology of proteotoxic stress in aging and disease. Cold Spring Harb Symp Quant Biol 76:91–99 4. Dubnikov T, Ben-Gedalya T, Cohen E (2017) Protein quality control in health and disease. Cold Spring Harb Perspect Biol 9(3):a023523 5. Tyedmers J, Mogk A, Bukau B (2010) Cellular strategies for controlling protein aggregation. Nat Rev Mol Cell Biol 11(11):777–788 6. Chen B, Retzlaff M, Roos T, Frydman J (2011) Cellular strategies of protein quality control. Cold Spring Harb Perspect Biol 3:a004374 7. Kisselev AF, Goldberg AL (2001) Proteasome inhibitors: from research tools to drug candidates. Chem Biol 8:739–758 8. Kawaguchi Y, Kovacs JJ, Mclaurin A, Vance JM, Ito A, Yao TP (2003) The deacetylase HDAC6 regulates aggresome formation and cell viability in response to misfolded protein stress. Cell 115:727–738 9. Szeto J, Kaniuk NA, Canadien V, Nisman R, Muzushima N, Yoshimori T, Bazett-Jones DP, Brumell JH (2006) ALIS are stress-induced protein storage compartments for substrates of the proteasome and autophagy. Autophagy 2:189–199 10. Miller SB, Ho CT, Winkler J, Khokhrina M, Neuner A, Mohamed MY, Guilbride DL, Richter K, Lisby M, Schiebel E, Mogk A, Bukau B (2015) Compartment-specific aggregases direct distinct nuclear and cytoplasmic aggregate deposition. EMBO J 34:778–797 11. Bush KT, Goldberg AL, Nigam SK (1997) Proteasome inhibition leads to a heat-shock response, induction of endoplasmic reticulum chaperones, and thermotolerance. J Biol Chem 272:9086–9092

12. Lee DH, Goldberg AL (1998) Proteasome inhibitors cause induction of heat shock proteins and trehalose, which together confer thermotolerance in Saccharomyces cerevisiae. Mol Cell Biol 18:30–38 13. Kaganovich D, Kopito R, Frydman J (2008) Misfolded proteins partition between two distinct quality control compartments. Nature 454:1088–1095 14. Bersuker K, Hipp MS, Calamini B, Morimoto RI, Kopito RR (2013) Heat shock response activation exacerbates inclusion body formation in a cellular model of huntington disease. J Biol Chem 288(33):23633–23638 15. Yu A, Shibata Y, Shah B, Calamini B, Lo DC, Morimoto RI (2014) Protein aggregation can inhibit clathrin-mediated endocytosis by chaperone competition. Proc Natl Acad Sci U S A 111:E1481–E1490 16. Banaszynski LA, Chen LC, Maynard-Smith LA, Ooi AG, Wandless TJ (2006) A rapid, reversible, and tunable method to regulate protein function in living cells using synthetic small molecules. Cell 126(5):995–1004 17. Iwamoto M, Bjo¨rklund T, Lundberg C, Kirik D, Wandless TJ (2010) A general chemical method to regulate protein stability in the mammalian central nervous system. Chem Biol 17(9):981–988 18. Miyazaki Y, Imoto H, Chen LC, Wandless TJ (2012) Destabilizing domains derived from the human estrogen receptor. J Am Chem Soc 134 (9):3942–3945 19. Maynard-Smith LA, Chen LC, Banaszynski LA, Ooi AG, Wandless TJ (2007) A directed approach for engineering conditional protein stability using biologically silent small molecules. J Biol Chem 282(34):24866–24872 20. Knowles TPJ, Vendruscolo M, Dobson CM (2014) The amyloid state and its association with protein misfolding diseases. Nat Rev Mol Cell Biol 15:384–396 21. Nakamura H, DeRose R, Inoue T (2019) Harnessing biomolecular condensates in living cells. J Biochem 166(1):13–27

AgDD System: A Chemical Controllable Protein Aggregates in Cells 22. Miyazaki Y, Mizumoto K, Dey G, Kudo T, Perrino J, Chen LC, Meyer T, Wandless TJ (2016) A method to rapidly create protein aggregates in living cells. Nat Commun 7:11689 23. Egeler EL, Urner LM, Rakhit R, Liu CW, Wandless TJ (2011) Ligand-switchable substrates for a ubiquitin-proteasome system. J Biol Chem 286:31328–31336

285

24. Grimley JS, Chen DA, Banaszynski LA, Wandless TJ (2008) Synthesis and analysis of stabilizing ligands for FKBP-derived destabilizing domains. Bioorg Med Chem Lett 18:759–761 25. Ramdzan YM, Polling S, Chia CPZ, Ng IHW, Ormsby AR, Croft NP, Purcell AW, Bogoyevitch MA, Ng DCH, Gleeson PA, Hatters DM (2012) Tracking protein aggregation and mislocalization in cells with flow cytometry. Nat Methods 9:467–470

Chapter 17 Intracellular Unnatural Catalysis Enabled by an Artificial Metalloenzyme Yasunori Okamoto and Ryosuke Kojima Abstract Artificial metalloenzymes, constructed by incorporating a synthetic catalyst into the internal spaces of a protein scaffold, can perform noncanonical chemical transformations that are not possible using natural enzymes. The addition of cell-permeable modules to artificial metalloenzymes allows for noncanonical catalysis to be implemented as a function of mammalian cells. In this chapter, we describe a protocol for controlling cellular function through a cascade consisting of an artificial metalloenzyme and a gene-circuit engineered via synthetic biology. Key words Intracellular catalysis, Artificial metalloenzyme, Gene switch, Cell-penetrating polydisulfides, Streptavidin, Thyroid hormone, Allylic dealkylation, Ruthenium complex

1

Introduction Rapid ongoing developments in synthetic biology have created the possibility of regulating cellular functions via redesigning biochemical reaction networks in a cell. From the viewpoint of chemical and pharmaceutical industries, cellular engineering holds great promise for generating microbial factories that synthesize high-value compounds [1] and cellular drugs that can release therapeutic and diagnostic biomolecules in response to the patient’s physiological environment [2–4]. With the typical synthetic biology approach, only genetically encodable modules are available for achieving biochemical regulation. To overcome this limitation, in recent years, synthetic chemists have attempted to implement noncanonical functions of synthetic molecules within cells. One of these approaches involves synthetic catalysts, which represent organic synthetic chemistry [5– 11]. However, introducing synthetic catalysts is challenging because of their inactivation in the sea of biomolecules present within a cell. This obstacle can be overcome by the use of artificial metalloenzymes (ArMs).

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

287

288

Yasunori Okamoto and Ryosuke Kojima

ArMs are obtained by incorporating a synthetic catalyst within a protein scaffold, which can improve the reaction rate and selectivity of the synthetic catalyst [12]. In addition to these features, previous studies have also focused on imparting biocompatibility to synthetic catalysts via ArM technology. It has been demonstrated that ArMs can be concurrently used with other natural enzymes to perform cascade reactions [13–16]. Recently, it was demonstrated that an ArM bearing a cell permeable module could perform an intracellular noncanonical chemical transformation [17]. It was shown that, relying on the strong supramolecular recognition between streptavidin (Sav) and biotin, the artificial deallylase, possessing a biotinylated ruthenium complex 1 in a Sav scaffold (Fig. 1), could catalyze an O-allyl carbamate cleavage reaction. The tetrameric nature of Sav allows further functionalization of this Sav-based artificial deallylase with a

Fig. 1 A biotinylated ruthenium complex 1 is an abiotic cofactor of the Sav-based artificial deallyase 1x
Sav. A fluorescent carboxytetramethylrhodamine (TAMRA) moiety in the biotinylated cellpenetrating poly(disulfide) (CPD) 2 allows the monitoring of cellular uptake of the cell-penetrating artificial deallylase. The cell-penetrating artificial deallylases 1x2y
Sav are prepared at various ratios (x and y) of biotinylated molecules 1 and 2 in tetrameric Sav. Adapted from [17]

Intracellular Catalysis by an Artificial Metalloenzyme

289

Fig. 2 (a) A cascade reaction comprising of endogenous esterases and the artificial deallylase sequentially converts the double-caged triiodothyronine (AM-AT3) 5 into triiodothyronine (T3) 3, via AT3 4. Upon binding of T3 3 to a thyroid-sensing receptor (TSR), encoded in pSP27, gene expression of secreted nanoluc-luciferase (sec-nluc) is induced. The activity of the constitutively expressed human placental secreted alkaline phosphatase (SEAP) can be used to assess transfection efficacy and cell viability. (b) The sec-nluc (luminescent conversion of 6 into 7) and SEAP (conversion of 8 into chromophore 9) activities are quantified using the supernatant of the cell culture medium

biotinylated cell-penetrating polydisulfide (CPD) 2, yielding the cell-penetrating artificial deallylase 1x2y
Sav (Fig. 1), where (x, y) indicates the ratio of biotinylated catalyst 1 and CPD 2 added to the tetrameric Sav. Catalytic activity of this cell-penetrating artificial deallylase can be tested via the following reaction cascade (Fig. 2): (1) a series of reactions involving endogenous esterases and the artificial deallylase to produce the thyroid hormone triiodothyronine (T3) 3 from a doubly caged T3 with an acetoxymethyl group (AM) and an allyloxycarbonyl (A) group (AM-AT3) 5, via AT3 4; (2) The produced

290

Yasunori Okamoto and Ryosuke Kojima

T3 3 binds to the thyroid-sensing receptor (TSR), constitutively expressed from the plasmid pSP27 (PhCMV-TSR-pA), [18] and subsequently the TSR-T3 complex induces the expression of a secreted nanoluc-luciferase (sec-nluc) [19] encoded in the plasmid pYO1 (PTSR5-secNluc-pA). The activity of sec-nluc converting furimazine 6 into product 7 accompanying emission of bioluminescence reflects the catalytic activity of the artificial deallylase. The third plasmid pSEAP2-control (PSV40-SEAP-pA) codes for a human placental secreted alkaline phosphatase (SEAP) [20], which is used as an internal standard for transfection efficacy and cell viability. This activity was evaluated by monitoring absorbance increase at 405 nm, that reflects conversion of p-nitrophenylphosphate 8 into p-nitrophenolate 9. The present chapter describes the detailed experimental protocol for inducing gene expression in designer mammalian cells via the aforementioned cell-penetrating artificial deallylase 1x2y
Sav.

2

Materials

2.1 Reagents Used for Preparation of an Artificial Deallylase and for Intracellular Catalysis

In this chapter, we aim at providing a detailed description of a procedure for intracellular catalysis, the reader is thus advised to refer to the listed references related to organic synthesis. 1. Lyophilized streptavidin (Sav) variants [21, 22]. 2. Cell-penetrating poly(disulfide) (CPD) 2 (Fig. 1) [23]. 3. Stock solution of a ruthenium complex 10 for the abiotic cofactor 1: 10 mM pentamethylcyclopentadienyl-tris(acetonitrile)ruthenium(II) hexafluorophosphate ([CpRu (MeCN)3]PF6) 10 (Fig. 3) in DMF. Weigh 4.34 mg [CpRu (MeCN)3]PF6 10 and dissolve it in 2 mL dry DMF (see Notes 1–4). 4. Stock solution of biotinylated ligand 11 for the abiotic cofactor 1: 10 mM biotinylated ligand 11 (Fig. 3) in DMF. Weigh 4.90 mg biotinylated ligand 11 and dissolve it in 2 mL dry DMF (see Notes 1–4) [17].

Fig. 3 The biotinylated ruthenium complex 1 is prepared by mixing [CpRu (NCCH3)3](PF6) 10 and the biotinylated ligand 11 in a 1:1 ratio

Intracellular Catalysis by an Artificial Metalloenzyme

291

5. Stock solution of substrate AM-AT3 5 for intracellular catalysis: 2 mM AM-AT3 5 (Fig. 2) in DMSO. Weigh 8.07 mg AM-AT3 5 and dissolve it in 5 mL dry DMSO (see Notes 1–4) [17]. 6. Stock solution of biotin-4-fluorescein (B4F) for the determination of free biotin-binding site (FBBS) of Sav: ca. 1 mM B4F in DMSO. Weigh 2.58 mg B4F and dissolve it in 4 mL DMSO (see Notes 3–5). 2.2 Plasmids and Transfection Reagents

1. pSEAP2-control (Clontech, 631717): Plasmid for constitutive SEAP expression. 2. pSP27 (PhCMV-TSR-pAbGH) [18]: Plasmid for constitutive TSR expression. 3. pYO1 (PUAS5-secNluc-pASV40). [17]: Plasmid for PUAS5driven sec-nanoluc expression. 4. Transfection reagent: 1 mg/mL polyethylenimine (PEI, 20000 MW). In a 50 mL conical tube, weigh 40 mg PEI and dissolve it in 40 mL ultrapure H2O. Sterilize the resulting solution by filtration using a 0.22 μm filter.

2.3 Reagents for Cell Culture

1. HEK-293T cells (DSMZ: ACC-635). 2. Medium A: Dulbecco’s modified eagle medium (DMEM) containing 10% (v/v) fetal bovine serum (FBS) and 1% (v/v) penicillin/streptomycin (PS) solution. To 445 mL DMEM, add 50 mL FBS and 5 mL 100 PS solution, then mix gently (see Notes 1, 6–8). 3. Medium B: DMEM containing 1%(v/v) PS. To 495 mL DMEM, add 5 mL 100 PS solution, then mix gently (see Notes 1, 6–8). 4. Medium C: DMEM containing 5% (v/v) charcoal-stripped FBS and 1%(v/v) PS. To 470 mL DMEM, add 25 mL charcoal-stripped FBS and 5 mL 100 PS solution, then mix gently (see Notes 1, 6–8). 5. Medium D: DMEM containing 1% (v/v) PS and 0.1 mg/mL heparin sodium salt. In a sterile 50 mL conical tube, weigh 4 mg heparin sodium salt and dissolve it in 40 mL Medium B (see Notes 1–3, 6–8). 6. 0.05% trypsin-EDTA.

2.4 Reagents for the Enzymatic Activity Assays

1. 2 SEAP buffer: 20 mM homoarginine, 1 mM MgCl2, and 21% (v/v) diethanolamine–HCl, pH ¼ 9.8. Store at 4  C in dark (up to 6 months). 2. p-Nitrophenyl phosphate 8 stock solution: 120 mM p-nitrophenyl phosphate 8 in water. Store at 20  C (preferably in single-use aliquots to thaw only once). 3. Nanoglo assay solution (Promega).

292

Yasunori Okamoto and Ryosuke Kojima

2.5 Instruments and Consumables

1. Autoclave. 2. Sterilizing filter. 3. Electronic precision balance placed on an antivibration table. 4. Micropipette, electronic micropipette. 5. Gastight syringe. 6. Quartz cuvette. 7. Multiwell plate (96-well black bottom, 96-well transparent bottom, 384-well black bottom). 8. Spectrophotometers (for absorbance, fluorescence). 9. Laminar flow cabinet (Clean bench). 10. Cell culture dish (diameter ¼ 10 cm). 11. CO2 incubator for mammalian cell culture. 12. Constant temperature water-bath. 13. Microscope. 14. Aspirator. 15. Centrifuge (for 96-well plate, 1.5 mL tube, 15 mL conical tube, and 50 mL conical tube). 16. Cell counter. 17. Collagen-coated 24-well cell culture plate. 18. Heat block (for 96-well plate, 1.5 mL tube). 19. Microplate reader luminescence).

3

(for

absorbance,

fluorescence,

Method

3.1 Preparation of Sav Stock Solution with 800 μM FBBS

1. Weigh 16.4 mg lyophilized Sav and dissolve it in 1 mL sterile water (see Notes 1–3, 9) (¼ca. 1 mM). 2. Add 4 μL ca. 1 mM Sav aqueous solution to 996 μL sterile PBS. (¼ca. 4 μM) (see Notes 1, 2, and 9). 3. Add 10 μL ca. 1 mM B4F stock solution (prepared in Subheading 2.1, item 6) to 990 μL PBS, then measure the UV-vis absorption spectrum using a spectrophotometer. Calculate the accurate B4F stock solution concentration by using the absorption coefficient at 495 nm ¼ 68,000 M1 cm1 [24] (see Note 9). 4. Based on the above-determined B4F stock solution concentration (prepared in Subheading 2.1, item 6), dilute it with PBS to 1.0, 1.5, 2.0, 2.5, 3.0, 3.5, 4.0, 4.5, 5.0, 5.5, and 6.0 μM (see Note 9).

Intracellular Catalysis by an Artificial Metalloenzyme

293

5. In a 96-well plate (flat black bottom), mix 50 μL ca. 4 μM Sav aqueous solution (prepared in step 2) with 50 μL B4F solution (prepared in step 4) (see Note 9). 6. Measure fluorescence at 520 nm (excitation at 485 nm) using a microplate reader. Plot fluorescent intensities in the function of B4F concentration, listed in step 4. Draw two regression lines both in ranges of lower and higher B4F concentrations, respectively. The x coordinate of the crossing point of these two regression lines corresponds to the concentration of the Sav solution FBBS (prepared in step 2). 7. Calculate the concentration of the Sav solution FBBS (prepared in step 1) and dilute it to 800 μM with sterile water (see Notes 1, 2, and 9). 3.2 Preparation of Artificial Deallylase 1x
Sav

1. Mix 50 μL 10 mM [CpRu(MeCN)3]PF6 10 in DMF (prepared in Subheading 2.1, item 3) with 50 μL 10 mM biotinylated ligand 11 in DMF (prepared in Subheading 2.1, item 4) (see Notes 2 and 4). 2. Incubate the mixture at an ambient temperature for 10 min to afford 5 mM biotinylated ruthenium complex 1. 3. Prepare the artificial deallylase 1x
Sav by mixing 5 mM biotinylated ruthenium complex 1 and 800 μM Sav (as monomer) under the conditions listed in Tables 1, 2, 3, respectively. Incubate the ruthenium complex 1 with water for 10 min before the addition of the Sav solution (see Notes 1, 2, 7, and 9).

3.3 Preparation of Cell-Penetrating Artificial Deallylase 1x2y
Sav

1. Prepare the cell-penetrating artificial deallylases 1x2y
Sav by mixing artificial deallylase 1x
Sav (prepared in Subheading 3.2) and 110 μM CPD 2 under the conditions listed in Tables 4, 5, 6, 7, 8, 9. After mixing artificial deallylases 1x
Sav and water, add CPD 2 and incubate the mixture at an ambient temperature for 2 h (see Notes 1, 2, and 9).

3.4 Cell Culture and Transfection (Fig. 4)

1. In a cell culture dish (diameter ¼ 10 cm), culture HEK-293T cells in 10 mL medium A (prepared in Subheading 2.3, item 2) at 37  C in a humidified atmosphere containing 5% CO2. 2. At approximately 80% confluency observed by a microscope, discard the medium A by using a sterile glass pipet connected to an aspirator. 3. Add 800 μL 0.05% trypsin-EDTA (Gibco) to the cells and incubate them in a humidified atmosphere containing 5% CO2 for 5 min. 4. Detach the cells by gentle pipetting.

294

Yasunori Okamoto and Ryosuke Kojima

Table 1 Stock solution of artificial deallylase 13
Sav Stock solution

Initial conc. (μM)

Final conc. (μM)

Vol. (μL)

Order of addition

Sav in water

800 (FBBS)

133.3

83.3

3

Ruthenium complex 1

5000

100.0

10.0

1

406.7

2

Sterile water

Total 500 μL

Table 2 Stock solution of artificial deallylase 12
Sav Stock solution

Initial conc. (μM)

Final conc. (μM)

Vol. (μL)

Order of addition

Sav in water

800 (FBBS)

200.0

125

3

Ruthenium complex 1

5000

100.0

10.0

1

365

2

Sterile water

Total 500 μL

Table 3 Stock solution of artificial deallylase 11
Sav Stock solution

Initial conc. (μM)

Final conc. (μM)

Vol (μL)

Order of addition

Sav in water

800 (FBBS)

400.0

250.0

3

Ruthenium complex 1

5000

100.0

10.0

1

240

2

Sterile water

Total 500 μL

Table 4 Stock solution of cell-penetrating artificial deallylase 1321
Sav

Stock solution

Initial conc. Content (μM)

Final conc. (μM)

Artificial deallylase 13
Sav

Sav Ru 1

133.3 (FBBS) 100.0

CPD 2

CPD 2

110.0

Sterile water

Vol. (μL)

Order of addition

100.0 75.0

15.0

2

25.0

4.5

3

0.5

1

Total 20 μL

Intracellular Catalysis by an Artificial Metalloenzyme

295

Table 5 Stock solution of cell-penetrating artificial deallylase 1222
Sav

Stock solution

Initial conc. Content (μM)

Final conc. (μM)

Vol. (μL)

Order of addition

Artificial deallylase 12
Sav

Sav Ru 1

200.0 (FBBS) 100.0

50.0 25.0

5.0

2

CPD 2

CPD 2

110.0

25.0

4.5

3

10.5

1

Sterile water

Total 20 μL

Table 6 Stock solution of cell-penetrating artificial deallylase 1221
Sav

Stock solution

Initial conc. Content (μM)

Final conc. (μM)

Vol. (μL)

Order of addition

Artificial deallylase 12
Sav

Sav Ru 1

200.0 (FBBS) 100.0

100.0 50.0

10.0

2

CPD 2

CPD 2

110.0

25.0

4.5

3

5.5

1

Sterile water

Total 20 μL

Table 7 Stock solution of cell-penetrating artificial deallylase 1121
Sav

Stock solution

Initial conc. Content (μM)

Final conc. (μM)

Artificial deallylase 11
Sav

Sav Ru 1

400.0 (FBBS) 100.0

CPD 2

CPD 2

110.0

Sterile water

Vol. (μL)

Order of addition

100.0 25.0

5.0

2

25.0

4.5

3

10.5

1

Total 20 μL

5. Add 2 mL medium A and transfer 350 μL of the cells into 10 mL of medium A in a cell culture dish (diameter ¼ 10 cm) for subculture. 6. Transfer the rest of the cells into a 15 mL sterile conical tube.

296

Yasunori Okamoto and Ryosuke Kojima

Table 8 Stock solution of cell-penetrating artificial deallylase 1123
Sav

Stock solution

Initial conc. Content (μM)

Final conc. (μM)

Vol. (μL)

Order of addition

Artificial deallylase 11
Sav

Sav Ru 1

400.0 (FBBS) 100.0

33.3 8.3

1.7

2

CPD 2

CPD 2

110.0

25.0

4.5

3

13.8

1

Sterile water

Total 20 μL

Table 9 Stock solution of cell-penetrating artificial deallylase 1122
Sav

Stock solution

Initial conc. Content (μM)

Final conc. (μM)

Artificial deallylase 11
Sav

Sav Ru 1

400.0 (FBBS) 100.0

CPD 2

CPD 2

110.0

Sterile water

Vol. (μL)

Order of addition

50.0 12.5

2.5

2

25.0

4.5

3

13.0

1

Total 20 μL

7. Centrifuge the conical tube at an ambient temperature and 200  g for 3 min. 8. Discard the supernatant by using a sterile glass pipette connected to an aspirator. 9. Tap the conical tube and gently resuspend the cells in 1 mL medium A. 10. Determine the number of the cells in 1 mL of the resuspension using a cell counter. 11. Seed 1.25  105 of HEK-293T cells into 10 mL medium A in a cell culture dish (diameter ¼ 10 cm). 12. Incubate the cells at 37  C in a humidified atmosphere containing 5% CO2 for 43 h. 13. Discard the cell culture medium A using a sterile glass pipette connected to an aspirator and add 10 mL medium C (prepared in Subheading 2.3, item 4) at least 30 min before the addition of the transfection mixture to the cells.

Intracellular Catalysis by an Artificial Metalloenzyme

297

Fig. 4 HEK-293T cells transfected with pSP27, pYO1, and pSEAP2-control plasmids, then dispensed into a collagen-coated 24-well cell culture plate and incubated overnight. After 1 h of incubation with the cellpenetrating artificial deallylase 1x2y
Sav, the cells are washed with medium D. Subsequently, the cell culture medium D is exchanged in the medium C containing AM-AT3 5 (4 μM) and the cells are incubated for 24 h. Activities of sec-nluc and SEAP activities are quantified using the supernatant of the cell culture medium

14. To prepare the transfection mix, add 1300 ng pSP27, 7800 ng pYO1, and 900 ng pSEAP2-control to 1 mL medium B (prepared in Subheading 2.3, item 3). Mix 50 μL 1 mg/mL PEI (prepared in Subheading 2.2, item 4) to the plasmid mixture solution and incubate it at an ambient temperature for 15 min. 15. Add 1 mL DNA–PEI mix to the cells (prepared in step 13) and incubate them in a humidified atmosphere containing 5% CO2 for 8 h. 16. Repeat above-described steps 3–8. 17. Resuspend the cells in 20 mL medium C (prepared in Subheading 2.3, item 4) in a 50 mL sterile conical tube. 18. Transfer 500 μL resuspension into a collagen-coated 24-well cell culture plate and culture it in a humidified atmosphere containing 5% CO2 for 18 h. 3.5 Intracellular Catalysis of Artificial Deallylase 1x2y
Sav (Fig. 4)

1. Dilute cell-penetrating artificial deallylases 1x2y
Sav (prepared in Subheading 3.3, step 1 and Tables 4, 5, 6, 7, 8, 9) by a factor fifty with medium B (prepared in Subheading 2.3, item 3) (see Notes 2 and 9).

298

Yasunori Okamoto and Ryosuke Kojima

2. Discard the transfected HEK-293T cell culture medium C (prepared in Subheading 3.4, step 18) in the 24-well plate and add 500 μL medium B containing artificial deallylase 1x2y
Sav (prepared in step 1) (see Notes 2, 9, and 10). 3. After 1 h of incubation in a humidified atmosphere containing 5% CO2, wash the cells three times with 500 μL medium D (prepared in Subheading 2.3, item 5) to remove cell-surface adsorbed artificial deallylases 1x2y
Sav [25] (see Notes 2, 9, and 10). 4. Prepare the medium C containing 4 μM AM-AT3 5 by mixing 20 μL 2 mM AM-AT3 5 in DMSO (prepared in Subheading 2.1, item 5) and 9.980 mL of the medium C. 5. Discard the cell culture medium D and add 500 μL medium C containing 4 μM AM-AT3 5 (see Notes 2, 9, and 10). 6. Incubate the cells in a humidified atmosphere containing 5% CO2 for 24 h. 3.6 Sec-nluc Activity Assay (Fig. 4)

1. Transfer 7.5 μL cell culture medium supernatant into a 384-well plate (flat black bottom). 2. Supplement with 7.5 μL nanoglo assay solution (Promega). 3. After 5 min of incubation, measure the luminescence using a microplate reader (see Note 11).

3.7 SEAP Activity Assay (Fig. 4)

1. Transfer 100 μL cell culture medium supernatant into a 96-well plate (flat transparent bottom). 2. Seal the 96-well plate and incubate it at 65  C for 30 min to heat-inactivate endogenous alkaline phosphatases. 3. Incubate at 4  C for 5 min, then centrifuge the 96-well plate at 4000  g for 5 min. 4. Transfer 80 μL heated supernatant into a new 96-well plate (flat transparent bottom) and mix with 100 μL 2 SEAP buffer and 20 μL 120 mM p-nitrophenyl phosphate 8. 5. Immediately after mixing, record the time-course of the absorption increase at 405 nm and 37  C using a microplate reader. Set the measurement conditions as follows: plate shaking for 5 s at 900 rpm before measurement, recording over 30 min with 30 s intervals. 6. From the linear part of the time-course plot, calculate the enzymatic activity of SEAP (in U/L). The correlation between the slope and the corresponding activity could be determined using a standard.

Intracellular Catalysis by an Artificial Metalloenzyme

4

299

Notes 1. All procedures should be performed on a clean bench dedicated to mammalian cell cultures. 2. Prepare fresh solutions on the day of experiments. 3. Use an electronic precision balance placed on an anti-vibration table to weigh materials. 4. Use gas-tight glass syringe for organic solvents. 5. Store in -20  C 6. Use sterile serological pipettes for only one time. 7. Store the materials at 4  C and use them within one month. 8. Do not put a micropipette tip to the stock media. First, transfer 40 mL of medium into a sterile conical tube (50 mL) and use it after warming up to 37  C. 9. Use sterile micropipette tips and discard them after every single dispensation. 10. Use a multichannel electronic pipette and dispense the solution at the slowest speeds (over 5 s) to avoid cell detachment. 11. The expression level of sec-nanoluc could be enhanced by increasing the biotinylated ruthenium complex 1-to-CPD 2 ratio.

Acknowledgments YO thanks JST ACT-X (JPMJAX1913), JSPS KAKENHI Grant (20K15393), and generous support from FRIS, Tohoku University. RK thanks financial support by JST-PRESTO grant (JPMJPR17H5), HFSP CDA (CDA-00008/2019-C), and JSPS KAKENHI grant (20H02874). References 1. Keasling JD (2010) Manufacturing molecules through metabolic engineering. Science 330:1355–1358. https://doi.org/10.1126/ science.1193990 2. Kojima R, Aubel D, Fussenegger M (2015) Novel theranostic agents for next-generation personalized medicine: small molecules, nanoparticles, and engineered mammalian cells. Curr Opin Chem Biol 28:29–38. https://doi. org/10.1016/j.cbpa.2015.05.021 3. Kojima R, Aubel D, Fussenegger M (2016) Toward a world of theranostic medication: programming biological sentinel systems for

therapeutic intervention. Adv Drug Deliv Rev 105:66–76. https://doi.org/10.1016/j.addr. 2016.05.006 4. Ausl€ander S, Ausl€ander D, Fussenegger M (2017) Synthetic biology—the synthesis of biology. Angew Chem Int Ed 56:6396–6419. https://doi.org/10.1002/anie.201609229 5. Rebelein JG, Ward TR (2018) In vivo catalyzed new-to-nature reactions. Curr Opin Biotechnol 53:106–114. https://doi.org/10.1016/j. copbio.2017.12.008 6. Tonga GY, Jeong Y, Duncan B et al (2015) Supramolecular regulation of bioorthogonal

300

Yasunori Okamoto and Ryosuke Kojima

catalysis in cells using nanoparticle-embedded transition metal catalysts. Nat Chem 7:597–603. https://doi.org/10.1038/ nchem.2284 7. Li J, Yu J, Zhao J et al (2014) Palladiumtriggered deprotection chemistry for protein activation in living cells. Nat Chem 6:352–361. https://doi.org/10.1038/ nchem.1887 8. Yusop RM, Unciti-Broceta A, Johansson EMV et al (2011) Palladium-mediated intracellular chemistry. Nat Chem 3:239–243. https://doi. org/10.1038/nchem.981 9. Vo¨lker T, Dempwolff F, Graumann PL, Meggers E (2014) Progress towards bioorthogonal catalysis with organometallic compounds. Angew Chem Int Ed 53:10536–10540. https://doi.org/10.1002/anie.201404547 10. Toma´s-Gamasa M, Martı´nez-Calvo M, Cou˜ as JL (2016) Transition ceiro JR, Mascaren metal catalysis in the mitochondria of living cells. Nat Commun 7:12538. https://doi. org/10.1038/ncomms12538 11. Coverdale JPC, Romero-Canelo´n I, SanchezCano C et al (2018) Asymmetric transfer hydrogenation by synthetic catalysts in cancer cells. Nat Chem 10:347–354. https://doi. org/10.1038/nchem.2918 12. Schwizer F, Okamoto Y, Heinisch T et al (2018) Artificial Metalloenzymes: reaction scope and optimization strategies. Chem Rev 118:142–231. https://doi.org/10.1021/acs. chemrev.7b00014 13. Ko¨hler V, Wilson YM, Du¨rrenberger M et al (2013) Synthetic cascades are enabled by combining biocatalysts with artificial metalloenzymes. Nat Chem 5:93–99. https://doi.org/ 10.1038/nchem.1498 14. Okamoto Y, Ko¨hler V, Paul CE et al (2016) Efficient In SituRegeneration of NADH mimics by an artificial metalloenzyme. ACS Catal 6:3553–3557. https://doi.org/10. 1021/acscatal.6b00258 15. Okamoto Y, Ko¨hler V, Ward TR (2016) An NAD(P)H-dependent artificial transfer hydrogenase for multienzymatic cascades. J Am Chem Soc 138:5781–5784. https://doi. org/10.1021/jacs.6b02470 16. Okamoto Y, Ward TR (2017) Cross-regulation of an artificial Metalloenzyme. Angew Chem Int Ed 56:10156–10160. https://doi.org/10. 1002/anie.201702181

17. Okamoto Y, Kojima R, Schwizer F et al (2018) A cell-penetrating artificial metalloenzyme regulates a gene switch in a designer mammalian cell. Nat Commun 9:1943. https://doi.org/ 10.1038/s41467-018-04440-0 18. Saxena P, Hamri CG, Folcher M et al (2016) Synthetic gene network restoring endogenous pituitary-thyroid feedback control in experimental graves’ disease. Proc Natl Acad Sci U S A 113:1244–1249. https://doi.org/10. 1073/pnas.1514383113 19. Hall MP, Unch J, Binkowski BF et al (2012) Engineered luciferase reporter from a deep sea shrimp utilizing a novel imidazopyrazinone substrate. ACS Chem Biol 7:1848–1857. https://doi.org/10.1021/cb3002478 20. Berger J, Hauber J, Hauber R et al (1988) Secreted placental alkaline phosphatase: a powerful new quantitative indicator of gene expression in eukaryotic cells. Gene 66:1–10. https://doi.org/10.1016/0378-1119(88) 90219-3 21. Klehr J, Zhao J, Kron AS, Ward TR, Ko¨hler V (2020) Streptavidin (Sav)-based artificial Metalloenzymes: cofactor design considerations and large-scale expression of host protein variants. In: Iranzo O, Roque A (eds) Peptide and protein engineering. Springer Protocols Handbooks. Humana, New York, NY 22. Mallin H, Hestericova´ M, Reuter R, Ward TR (2016) Library design and screening protocol for artificial metalloenzymes based on the biotin-streptavidin technology. Nat Protoc 11:835–852. https://doi.org/10.1038/ nprot.2016.019 23. Derivery E, Bartolami E, Matile S, GonzalezGaitan M (2017) Efficient delivery of quantum dots into the cytosol of cells using cellpenetrating poly(disulfide)s. J Am Chem Soc 139:10172–10175. https://doi.org/10. 1021/jacs.7b02952 24. Mittal R, Bruchez MP (2011) Biotin-4-fluorescein based fluorescence quenching assay for determination of biotin binding capacity of streptavidin conjugated quantum dots. Bioconjug Chem 22:362–368. https://doi.org/ 10.1021/bc100321c 25. Gasparini G, Matile S (2015) Protein delivery with cell-penetrating poly(disulfide)s. Chem Commun 51:17160–17162. https://doi.org/ 10.1039/c5cc07460f

Chapter 18 Feeder-Free Human Induced Pluripotent Stem Cell Culture Using a DNA Aptamer-Based Mimic of Basic Fibroblast Growth Factor Yuri Hayata, Ryosuke Ueki, and Shinsuke Sando Abstract Cell culture media are often supplemented with recombinant growth factors and cytokines to reproduce biological conditions in vitro. Basic fibroblast growth factor (bFGF) has been widely used to support the pluripotency and self-renewal activity of human induced pluripotent stem cells (hiPSCs). We had previously developed a synthetic surrogate for bFGF on the basis of a DNA aptamer that binds to one of the FGF receptors. Since DNA aptamers have advantages over recombinant proteins in terms of thermal stability and production cost, replacing recombinant growth factors in cell culture media with DNA aptamers would be of great interest. Herein, we describe our protocol for feeder-free hiPSC culture using a DNA aptamerbased mimic of bFGF. Key words DNA aptamers, Agonistic aptamers, Receptor agonists, Human induced pluripotent stem cells, Fibroblast growth factors

1

Introduction DNA aptamers, which are single-stranded DNA molecules that adopt a unique folding structure and recognize target molecules with high specificity, have attracted much attention as synthetic alternatives to recombinant proteins [1, 2]. DNA aptamers have high thermal stability and can be chemically synthesized at a lower cost than that of recombinant proteins produced by bacterial or animal cells [1, 2]. Therefore, DNA aptamers are promising synthetic surrogates for recombinant growth factors. Several aptamerbased functional mimics of growth factors have been reported to date, such as hepatocyte growth factor [3, 4], fibroblast growth factor (FGF) [5, 6], and vesicular endothelial growth factor [7, 8]. We had previously reported our development of a 38-mer DNA aptamer that binds to FGF receptor 1 (FGFR1) and demonstrated that it could be converted into an FGFR1 agonist when synthesized in a dimeric form [5]. We also revealed that this

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

301

302

Yuri Hayata et al.

agonistic aptamer, designated TD0, behaved like a functional mimic of basic fibroblast growth factor (bFGF), which is as a fundamental growth factor for the maintenance of human induced pluripotent stem cells (hiPSCs) because its signaling activity supports their pluripotency and self-renewal activity [5]. Herein, we describe our protocol in which this DNA aptamer-based mimic of bFGF is used instead of recombinant bFGF for feeder-free hiPSC culture. As mentioned in our original report [5], this aptamer does not perfectly recapitulate the cell signaling induced by bFGF. For instance, the current protocol could not sustain the expression level of pluripotent markers in long-term culture [5]. Thus, it is necessary to further confirm whether the cells maintained according to this protocol can be used for their intended downstream applications. We also note that this protocol is not suitable for diagnostic and therapeutic uses.

2

Materials

2.1 Cell Culture Medium

1. StemFit Basic02: Mix solution A and solution B according to the manufacturer’s protocol. Store at 4
C. For long-term storage, store at 20
C. 2. StemFit AK02N: Mix solution A, solution B, and solution C according to the manufacturer’s protocol. Store at 4
C. For long-term storage, store at 20
C. 3. iMatrix-511 0.5 μg/μL solution: Store at 4
C. 4. Y-27632: 10 mM in autoclaved water. Store at 20
C. 5. 10 Dulbecco’s phosphate-buffered saline (DPBS) solution without calcium (Ca2+) and magnesium (Mg2+), 0.2-μm sterile filtered. 6. Autoclaved water or distilled water (0.2-μm sterile filtered). 7. bFGF: 100 nM in autoclaved DPBS. For long-term storage, 0.1% (w/v) bovine serum albumin (BSA) can be added as a carrier protein. Store at 20
C. 8. DNA aptamer: Order the following TD0 sequence (76 mer) from a standard oligo supplier: 50 GCC GCG TCT TTA TGG CTG GGG ATG GTG TGG GTT GCG CCG CGT CTT TAT GGC TGG GGA TGG TGT GGG TTG CGG C 30 . The oligo should have been purified with an OPC column or by HPLC and shipped in a lyophilized form. Store at 20
C in the dark before reconstitution. 9. Cell culture dish (35 mm). 10. Cell dissociation buffer: 0.5 mM ethylenediaminetetraacetic acid (EDTA) in autoclaved PBS.

hiPSC Culture Using a DNA-Based Growth Factor Mimic

303

11. Humidified CO2 incubator: Keep at 37
C, with 5% CO2 atmosphere. 12. hiPSCs: 409B2 cells (#HPS0076, RIKEN BRC Cell Bank) are used in this protocol. The cells are maintained in StemFit AK02N complete medium. 13. Microvolume spectrometer. 14. Thermal cycler.

3

Methods Basically, this protocol was designed according to a previously reported hiPSC culture method that used laminin fragments in an uncoated manner [9]. All the procedures must be performed aseptically, unless otherwise specified. Penicillin–streptomycin can be supplemented to the culture medium.

3.1 Preparation of the Aptamer Stock Solution

1. Dissolve enough lyophilized DNA powder with autoclaved water to make up an approximately 100 μM solution (see Note 1). 2. After sterilization through a 0.2-μm filter, aliquot a small portion for concentration measurement (see Note 2). 3. Determine the DNA concentration with a microvolume spectrometer. This step does not have to be performed aseptically. Discard the DNA sample used for the measurement. 4. On the basis of the DNA concentration determined, prepare a 20 aptamer solution in 1 DPBS (aptamer stock solution) using 10 DPBS and autoclaved water. Typically, we set the final TD0 concentration to 500 nM. 5. Refold the aptamer by heating to 95
C for 5 min, and then cool the solution slowly (e.g., at 0.1
C/s) to 25
C using a thermal cycler. Store the aptamer stock solution at 4
C.

3.2 Maintenance of Human iPSCs in Culture Medium Containing the bFGF-Mimicking Aptamer

The cell culture can be performed in cell culture dishes or microwell plates of any type or size. The volume of the medium is determined according to the typical cell culture conditions. In this protocol, a method using a 35-mm dish is described. 1. Prepare the seeding medium (StemFit Basic02 containing 10 μM Y-27632 and 0.3 μg/cm2 iMatrix-511) in a conical tube. Dispense 1900 μL of seeding medium to the dish and then add 100 μL of the 20 aptamer stock solution. Place the dish in a CO2 incubator until the cells are seeded. 2. Wash the hiPSCs that have been maintained in StemFit AK02N complete medium twice with 1 mL of DPBS. 3. Add 1 mL of cell dissociation buffer to the dish.

304

Yuri Hayata et al.

Fig. 1 Phase-contrast images of hiPSCs after 4 days of culture in the presence of (a) vehicle control, (b) aptamer TD0 (500 nM), or (c) bFGF (3 nM). Scale bars: 250 μm

4. Place the dish in the CO2 incubator. 5. After 2–4 min of incubation, observe the cells with a microscope. If the cells are still tightly packed in the colonies, place the dish in the CO2 incubator again and incubate for a further 1–2 min. This step is continued until gaps can be observed between the cells in the colonies. 6. Aspirate the cell dissociation buffer carefully and then add 1 mL of seeding medium. Immediately detach the cells from the dish with a cell scraper and dissociate the cells into a single-cell suspension through gentle pipetting. 7. Count the cells and then seed 2.5  103 cells/cm2 into another dish (see Note 3). 8. Place the dish in the CO2 incubator. 9. After 24 h from cell seeding, change the medium to StemFit Basic02 supplemented with the DNA aptamer (see Note 4). 10. Place the dish in the CO2 incubator. 11. Change the medium every 1 or 2 days (Fig. 1). Passage the cells according to procedures Subheading 3.2, steps 1–8 until they reach approximately 70–80% confluency (see Note 5).

4

Notes 1. If insoluble materials remain during reconstitution of the lyophilized DNA powder, centrifuge the solution and recover the supernatant. 2. Measure the DNA concentration after the filtration because the concentration of DNA may be reduced as a result of adsorption to the filter.

hiPSC Culture Using a DNA-Based Growth Factor Mimic

305

3. The volume of cell suspension added to the dish or well should be no more than 1/100 of the total medium volume in each dish or well, so that the addition of cell suspension does not affect the agonist concentration. If the cell suspension is too concentrated, dilute it by adding the seeding medium. 4. Y-27632 and iMatrix-511 are added to the medium for cell seeding only. Y-27632 should be removed from the medium at 24 h after the seeding. Avoid shortened or extended incubations with Y-27632. 5. The optimal timing for cell passaging may vary depending on the conditions. If differentiated cells are observed at the edge of the colonies, passage the cells even if they do not reach high confluency.

Acknowledgments This work was supported by a Leave a Nest Scientific Research grant from Ikedarika Scientific Co., Ltd. and Leave a Nest Co., Ltd., and a Noguchi-Shitagau Research Grant from the Noguchi Institute, and partly by a research grant from Asahi Glass Foundation. References 1. Keefe AD, Pai S, Ellington A (2010) Aptamers as therapeutics. Nat Rev Drug Discov 9:537–550 2. Zhou J, Rossi J (2017) Aptamers as targeted therapeutics: current potential and challenges. Nat Rev Drug Discov 16:181–202 3. Ueki R, Ueki A, Kanda N, Sando S (2016) Oligonucleotide-based mimetics of hepatocyte growth factor. Angew Chem Int Ed 55:579–582 4. Ueki R, Atsuta S, Ueki A, Sando S (2017) Nongenetic reprogramming of the ligand specificity of growth factor receptors by bispecific DNA aptamers. J Am Chem Soc 139:6554–6557 5. Ueki R, Atsuta S, Ueki A, Hoshiyama J, Li J, Hayashi Y, Sando S (2019) DNA aptamer assemblies as fibroblast growth factor mimics and their application in stem cell culture. Chem Commun 55:2672–2675

6. Kamatkar N, Levy M, He´bert JM (2019) Development of a monomeric inhibitory RNA aptamer specific for FGFR3 that acts as an activator when dimerized. Mol Ther Nucleic Acids 17:530–539 7. Ramaswamy V, Monsalve A, Sautina L, Segal MS, Dobson J, Allen JB (2015) DNA aptamer assembly as a vascular endothelial growth factor receptor agonist. Nucleic Acid Ther 25:227–234 8. Yoshitomi T, Hayashi M, Oguro T, Kimura K, Wayama F, Furusho H, Yoshimoto K (2020) Binding and structural properties of DNA aptamers with VEGF-A-mimic activity. Mol Ther Nucleic Acids 19:1145–1152 9. Miyazaki T, Isobe T, Nakatsuji N, Suemori H (2017) Efficient adhesion culture of human pluripotent stem cells using laminin fragments in an uncoated manner. Sci Rep 7:41165

Part V New Techniques to Engineer Specific Mammalian Cells in a Targeted Manner

Chapter 19 Protocol for De Novo Gene Targeting Via In Utero Electroporation Yuji Tsunekawa, Raymond Kunikane Terhune, and Fumio Matsuzaki Abstract Developments in genome-editing technology, especially CRISPR-Cas9, have revolutionized the way in which genetically engineered animals are generated. However, the process of generation includes microinjection to the one-cell stage embryo and the transfer of the microinjected embryo to the surrogate animals, which requires trained personnel. We recently reported the method includes introduction of CRISPR-Cas9 systems to the developing cerebral cortex via in utero electroporation thus generating gene-targeted neural stem cells in vivo. This technique is widely applicable for gene knockout, monitoring gene expression, and lineage analysis in developmental biology. In this chapter, the detailed protocol of EGFP (enhanced green fluorescent protein) knock-in method via in utero electroporation is described. Key words De novo gene targeting, In utero electroporation, CRISPR-Cas9, Developmental biology, Visualization of protein subcellular localization

1

Introduction Genetic manipulation of animals is an indispensable approach for understanding the molecular mechanism underlying the development of an organism as well as of human diseases. Generating these animals, especially knockout (KO) and knock-in (KI) mice, have been conventionally done by embryonic stem (ES) cell-based genetargeting method [1, 2]. Mutations or reporter genes are inserted into genomes of mouse ES cells through homologous recombination (HR), and targeted ES cells are injected into blastocysts to obtain gene-targeted mice. Even though this is established method, it is a laborious and time-consuming method; therefore, an alternative method has been developed to accelerate the process of KO-animal generation. The mRNA/DNA/protein injection of site-specific nucleases ZFN, TALEN, and CRISPR-Cas9 to the

Supplementary Information: The online version of this chapter (https://doi.org/10.1007/978-1-0716-14419_19) contains supplementary material, which is available to authorized users. Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

309

310

Yuji Tsunekawa et al.

CRISPR cut site

Genome GOI

0.5kbp-1kbp

2-3kbp

Linker or 2A sequence 5’HR arm

EGFP

3’HR arm

pLeakless III Backbone vector

Fig. 1 Schematic depiction of an example of de novo targeted-KI donor vector [12]. In this example, the donor vector is constructed with the EGFP reporter gene flanked by short homology arms (0.5–1 kbp, 2–3 kbp each) of the gene of interest (GOI), which was inserted between the last amino acid codon and the stop codon to be fused in frame with the coding sequence of the GOI. The targeting sequence was cloned into pLeaklessIII backbone vector to avoid the unexpected leakage. To express EGFP as a fusion protein with the GOI product, a short linker sequence should be inserted between the 50 homology arm and EGFP. To express EGFP separately from the GOI product under its promoter activity, the self-cleaving P2A peptides should be inserted between the homology arm and EGFP

one-cell stage embryo was successfully applied to generate KO animals via nonhomologous end joining, which results in insertions or deletions at the targeted sites [3–10]. Generation of KI animals via HR by injecting site-specific nucleases with ectopic donor DNA to the one-cell embryo has also been reported [11]. However, to obtain KO and KI animals, the following steps need to be followed: (1) microinjection of site-specific nucleases to the one-cell stage embryo, and (2) transplantation to surrogate animals, which is still a difficult technique to perform by individual researchers. Hence an alternative technique called in utero electroporation is widely used in many fields such as brain development [12–14]; we recently demonstrated high-efficiency somatic gene targeting via in utero electroporation (de novo gene targeting) [15]. Here, a detailed protocol of the brain-specific gene-targeting method via in utero electroporation is reported with the EGFP gene as an example for the inserted gene (Fig. 1).

2

Materials Prepare all solutions using ultrapure water and analytical grade reagents. 1. Mouse genome: Prepare the mouse genome from the soft tissue of mice by DNeasy Blood & Tissue Kit (QIAGEN, Hilden, Germany).

Protocol for de novo Gene Targeting

311

2. LB medium: Dissolve 2.5 g of LB broth to 100 ml of distilled water (DW), in a trypsinizing flask for microbiology, autoclave at 120  C for 20 min, and keep it room temperature until use. 3. Ampicillin stock: Dissolve 1 g of ampicillin sodium for injection in 20 ml of DW to make 50 mg/ml stock (500 stock) solution and keep it at 20  C until use. 4. End toxin free midi prep kit: Macherey-Nagel, Du¨ren Germany (Include TE buffer). 5. Betaine solution: Sigma-Aldrich, St. Louis, USA. 6. In-Fusion HD Cloning Kit: Takara Bio Inc., Shiga, Japan. 7. PrimeSTAR GXL DNA Polymerase: Takara Bio Inc., Shiga, Japan (including PrimeSTAR GXL, 5 PrimeSTAR GXL Buffer, dNTP Mixture). 8. PCR primer for genome amplification: Design PCR primers via website (http://bioinfo.ut.ee/primer3/) with the following criteria: primer size: minimum 25 bp/optimal 30 bp/maximum 35 bp, primer Tm: minimum 65  C/optimal 68  C/ maximum 72  C. 9. PCR primer for gRNA construction: Design the guide RNAs (gRNA) against the gene of your interest by CRISPRdirect website (https://crispr.dbcls.jp/) and order oligo DNAs as follows (see Note 3): Forward gRNA 50 -TTATATATCTTGTG GAAAGGACGAAACACC gX XXXXXXXXXXXXXXXXX –30 (x is the 20-bp target sequence. Change the first x to g), Reverse gRNA 5’- ATTTTAACTTGCTATTTCTAGCTC TAAAACYYYYYYYYYYYYYYYYYYYc-30 (change the last Y to c, YYYY is the reverse complementary of XXXX). 10. Wizard SV Gel and PCR Clean-UP System: Promega, Madison, USA. 11. Request all plasmids and backbone vectors to [email protected] (pCAG-hCas9, pLeaklessIII backbone vector, gRNA backbone vector, pCAX-EGFP and pCAG-mCherry). 12. Pregnant mice: Embryonic stages are calculated considering noon on the day of the vaginal plug as embryonic day 0.5 (E0.5). All animal experiments must be performed in compliance with the guidelines for animal experiments at the researcher’s institution. In this protocol, E13.5 ICR mice purchase from company will be used. 13. 0.1% Fast Green solution stock: Dissolve 10 mg of Fast Green FCF in 10 ml of TE buffer. Keep an aliquot of the stock solution (1 ml) at 20  C until use. 14. Glass capillary: Pull a glass capillary tube using a micropipette puller (Sutter Instrument Company, Novato, USA,

312

Yuji Tsunekawa et al.

Model#P-97/IVF) with the program: Time ¼ 1, P ¼ 500, Hear ¼ 800, Pull ¼ 30, Vel ¼ 40. Pinch off the tip with forceps to make the tip with a 30 μm diameter.

3

Methods

3.1 Plasmid Construction

3.2 Donor Vector Construction

All plasmids used in this protocol are purified via end toxin free midi/maxi prep kit. PCR primers to amplify homology arm sequences from the genomic DNA should avoid repeat sequences residing in the genome [16] (see Note 1). Donor vectors are constructed into pLeaklessIII backbone vector (full sequence available in Supplemental data 1) [15] to avoid unexpected expression of donor sequences from the donor vector (leakage). Lengths of each homology arms are set as 0.5–1 kbp (50 -side) and 2–3 kbp (30 -side). 1. Amplify the genomic region containing the homology arm sequence (see Note 2 and Fig. 2). PCR reagent for genome amplification 5 PrimeSTAR GXL Buffer

10 μl

dNTP mixture

4 μl

Betaine solution

5 μl

Primers forward and reverse (10 μM) see Note 2

0.25 μl each

Mouse genomic DNA (100 ng/μl)

1 μl

PrimeSTAR GXL DNA polymerase

1 μl

DW

28.5 μl

Total volume

50 μl

PCR program for genome amplification Preincubation

96  C

3 min

# 30 cycles

96  C 

68 C

30 s (DNA length/1000) min

# 68  C

5 min

# Hold

12  C

2. Amplify homology arm sequences from the PCR amplicon of step 1.

Protocol for de novo Gene Targeting

Homology arm1 Backbone vector

Homology arm2 Backbone vector

EGFP

EcoRI cut

313

KpnI cut

Fig. 2 Schematic depiction of donor vector construction by In-Fusion HD Cloning Kit. PCR primers (blue arrow) are designed to have 20 bp overhang sequences (blue dashed line)

PCR reagent for homology arm amplification 5 PrimeSTAR GXL Buffer

10 μl

dNTP mixture

4 μl

Primer forward and reverse (10 μM) for in fusion kit construction (see Note 2 and Fig. 2)

0.25 μl each

PCR amplicon from step 1

1 μl (100 pg/μl)

PrimeSTAR GXL DNA polymerase

1 μl

DW

33.5 μl

Total volume

50 μl

PCR program for genome amplification Preincubation

96  C

3 min

#

25 cycles

96  C

20 s

55  C

20 s

68  C

Gene length/1000  min

# 68  C

5 min

# Hold

12  C

3. Amplify the EGFP donor sequence by PCR (details are described below). PCR reagent for EGFP amplification 5 PrimeSTAR GXL Buffer

10 μl

dNTP mixture

4 μl

Primer forward and reverse (10 μM)

0.25 μl each

EGFP cDNA (1 ng/μl)

1 μl

PrimeSTAR GXL DNA polymerase

1 μl (continued)

314

Yuji Tsunekawa et al. DW

33.5 μl

Total volume

50 μl

PCR program for EGFP amplification Preincubation

96  C

3 min

# 96  C

30 s



DNA length/1000  min

55 C 25 cycles

30 s



68 C # 68  C

5 min

# Hold

12  C

4. Digest pLeaklessIII vector with EcoRI and KpnI for 1.5–16 h (Fig. 2). 5. Gel purify the amplified EGFP DNA fragment and the digested pLeaklessIII by Wizard SV Gel and PCR Clean-UP System. Gel purification Electrophoresis PCR-amplified DNA # Cut the expected size of DNA band # Add equal gel volume of membrane binding solution # Incubate at 50  C for 10 min # Load sample onto a minicolumn # Spin down at 10,000 rpm for 15 s # Repeat this step 2 times Wash minicolumn by adding 750 μl membrane wash solution #

(continued)

Protocol for de novo Gene Targeting

315

Spin down at 10,000 rpm for 15 s # Dry minicolumn by spin down at 10,000 rpm for 2 min # Elute DNA fragment by adding 30 μl of DW # Spin down at 10,000 rpm for 2 min # Proceed to the DNA fragment assembly

6. Assemble EcoRI/KpnI cut pLeaklessIII vector, 50 homology arm, 30 homology arm, and EGFP DNA fragment using In-Fusion HD Cloning Kit and transform the assembled DNA to E. coli competent cells. In vitro DNA fragment assembly and E. coli transformation reaction mixture EcoRI/KpnI cut pLeaklessIII vector (50 ng/μl)

0.5 μl

50 homology arm DNA fragment (50 ng/μl)

0.3 μl

0

3 homology arm DNA fragment (50 ng/μl)

0.7 μl

EGFP DNA fragment (50 ng/μl)

0.5 μl

5 in-fusion HD enzyme mix

0.5 μl

Total volume

50 μl 

Incubate at 50 C for 15 min # Mix with 50 μl of E. coli competent cells # Incubate on ice for 15 min # Incubate at 42  C for 40 s # Incubate on ice for 2 min # (continued)

316

Yuji Tsunekawa et al. Plate all E. coli competent cells to the ampicillin plate # Incubate at 37  C overnight

7. Pick up several colonies of E. coli and inoculate them in 2 ml LB medium containing ampicillin and incubate for 10 h. Extract plasmids by miniprep and check the sequence of the plasmids. Reserve 200 μl of cultured E. coli for the next step. 8. Choose the right colonies, add 100 ml of ampicillin added LB (final 100 μg/ml ampicillin) to the reserved E. coli from step 7 and incubate in shaking culture at 37  C for 14–16 h. 9. Purify plasmid DNA for in utero electroporation by end toxin free midi prep kit. Elute DNA with TE buffer and stock it at 20  C until use. 3.3 Guide RNA Vector Construction

1. Mix forward and reverse gRNA DNA primers and amplify gRNA DNA fragments by PCR self-amplification. PCR reagent for gRNA amplification 5 PrimeSTAR GXL Buffer

10 μl

dNTP mixture

4 μl

gRNA Primer Forward & Reverse (10 μM)

0.25 μl each

PrimeSTAR GXL DNA polymerase

1 μl

DW

34.5 μl

Total volume

50 μl

PCR program for gRNA amplification Preincubation

96  C

30 s

#

33 cycles

96  C

10 s

65  C

20 s

68  C

5s

# 68  C

2 min

# Hold

12  C

2. Digest gRNA backbone vector (full sequence available in Supplemental data 2) by AflII for 1.5–16 h.

Protocol for de novo Gene Targeting

317

3. Gel purify the amplified gRNA DNA fragment and restriction enzyme digested gRNA backbone plasmid by Wizard SV Gel and PCR Clean-Up System. 4. Assemble gRNA backbone and gRNA DNA fragment using In-Fusion HD Cloning Kit and transform the assembled DNA to E. coli competent cells. DNA fragment assembly AflII cut gRNA backbone plasmid (50 ng/μl)

0.5 μl

gRNA DNA fragment (5 ng/μl)

1.5 μl

5 in-fusion HD enzyme mix

0.5 μl

Total volume

2.5 μl

Incubate at 50  C for 15 min # Mix with 50 μl of E. coli competent cells # Incubate on ice for 15 min # Incubate at 42  C for 40 s # Incubate on ice for 2 min # Plate all E. coli to the ampicillin plate # Incubate at 37  C overnight

5. Pick up several colonies of E. coli and culture them in 2 ml LB buffer with ampicillin for 10 h or overnight under shaking. Extract plasmids using miniprep and check the sequence of the plasmids. Reserve 200 μl of cultured E. coli for the next step. 6. Choose the right colonies, add 100 ml of LB buffer with ampicillin to reserved E. coli from step 5, and incubate at 37  C for 16 h or overnight under shaking. 7. Purify plasmid DNA for in utero electroporation using the end toxin free midi prep kit (Macherey-Nagel). Elute DNA with TE buffer and stock it at 20  C until use. 3.4 In Utero Electroporation

1. Prepare DNA solution for in utero electroporation. DNA solution will contain pCAG-hCas9 (humanized cas9 expression vector), pLeaklessIII-Donor (homology donor vector for EGFP KI), pgRNA (gRNA expression vector), pCAG-

318

Yuji Tsunekawa et al.

mCherry (mCherry expression vector for an electroporation maker), and Fast Green (dye for visualizing DNA solution). DNA solution for in utero electroporation pCAG-hCas9 (5 μg/μl stock)

1 μl

pLeaklessIII-donor (5 μg/μl stock)

1 μl

pgRNA (2 μg/μl stock)

1 μl

pCAG-mCherry (1 μg/μl stock)

1 μl

0.1% FastGreen

0.5 μl

TE buffer

5.5 μl

Total volume

10 μl

Settle the glass capillary in the aspirator tube assembly (Drummond SC, Broomall, USA) and fill it with the DNA solution. 2. Gas anesthetize pregnant ICR mice (E13.5) with 2.5% isophlorin using NARCOBIT-E (Natsume Seisakusho Co., Ltd., Tokyo, Japan). 3. Sterilize the abdomen with 70% alcohol. Make an incision at the abdominal midline with fine scissors, and place sterilize gauze around the wound. Wet the gauze with saline. Pull out all uterine horns carefully onto the wet gauze. 4. Hold the embryo with two fingers, insert the glass capillary carefully into the lateral ventricle of the embryo, which can be seen through by illuminating the uterus. Inject approximately 1 μl of the DNA solution. 5. Pinch the embryo with platinum electrodes for electroporation (Nepa Gene Co., Ltd., Chiba, Japan), and apply square-wave current pulses (Volt ¼ 35, pON ¼ 50, pOFF ¼ 950, pulse number ¼ 5) using Cuy21 electroporator (BEX Co., Ltd., Tokyo, Japan). 6. Place back the uterine horn into the original location and add 1 ml of warm saline. 7. Suture the abdominal muscle with a surgical suture and close the outer skin layer with the 9 mm autoclip (BD Japan Co., Ltd., Fukushima, Japan). 8. Place animals on a heating pad for 10 min until they recovered from anesthesia. Put them back into their cages and wait for 3 days. 9. Sample the E17.5 embryo (see Note 4), and check for EGFP de novo KI by genotyping and immunohistochemistry.

Protocol for de novo Gene Targeting

4

319

Notes 1. Genomic repeat sequence can be visualized by [Mask repeat] in sequence formatting options at the UCSC genome browser (https://genome.ucsc.edu/index.html>go to the link of the genome browser >choose the species and the gene>go to the link of [genomic sequence] of the Sequence and Links to Tools and Databases). 2. PCR primers to amplify the homology arm usually have 20–40 bp overhang sequence for DNA assembly via In-Fusion HD Cloning Kit (Fig. 2). Because of these overhang sequences, direct amplification of the homology arm sequences from the genome sometimes fails using the PCR primers designed for In-Fusion HD Cloning Kit. Genomic DNAs including the homology arm sequences therefore should be once amplified with ordinary PCR primers from the genomic DNA, and then, the these amplified genomic DNAs should be amplified using PCR primers for In-Fusion Kit construction of the donor vector. 3. gRNA should be designed as near to the stop codon as possible. According to the authors experience, de novo KI can be occurred with the gRNA 30 bp away from stop codon with a lower efficiency. 4. Wait for at least 2 days for de novo knock-in.

References 1. Capecchi MR (2001) Generating mice with targeted mutations. Nat Med 7:1086–1090 2. Zijlstra M, Li E, Sajjadi F, Subramani S, Jaenisch R (1989) Germ-line transmission of a disrupted beta 2-microglobulin gene produced by homologous recombination in embryonic stem cells. Nature 342:435–438 3. Carbery ID et al (2010) Targeted genome modification in mice using zinc-finger nucleases. Genetics 186:451–459 4. Geurts AM et al (2009) Knockout rats via embryo microinjection of zinc-finger nucleases. Science 325:433 5. Hai T, Teng F, Guo R, Li W, Zhou Q (2014) One-step generation of knockout pigs by zygote injection of CRISPR/Cas system. Cell Res 24:372–375 6. Kou Z et al (2015) CRISPR/Cas9-mediated genome engineering of the ferret. Cell Res 25:1372–1375 7. Ni W et al (2014) Efficient gene knockout in goats using CRISPR/Cas9 system. PLoS One 9:e106718

8. Niu Y et al (2014) Generation of genemodified cynomolgus monkey via Cas9/ RNA-mediated gene targeting in one-cell embryos. Cell 156:836–843 9. Sung YH et al (2013) Knockout mice created by TALEN-mediated gene targeting. Nat Biotechnol 31:23–24 10. Tesson L et al (2011) Knockout rats generated by embryo microinjection of TALENs. Nat Biotechnol 29:695–696 11. Aida T et al (2015) Cloning-free CRISPR/Cas system facilitates functional cassette knock-in in mice. Genome Biol 16:87 12. Tabata H, Nakajima K (2001) Efficient in utero gene transfer system to the developing mouse brain using electroporation: Visualization of neuronal migration in the developing cortex. Neuroscience. 103:865–872

320

Yuji Tsunekawa et al.

13. Saito T, Nakatsuji N (2001) Efficient gene transfer into the embryonic mouse brain using in vivo electroporation. Develop Biol 240:237–246 14. Fukuchi-Shimogori T, Grove EA (2001) Neocortex patterning by the secreted signaling molecule FGF8. Science 294:1071–1074 15. Tsunekawa Y et al (2016) Developing a de novo targeted knock-in method based on in

utero electroporation into the mammalian brain. Development 143:3216–3222 16. de Koning APJ, Gu W, Castoe TA, Batzer MA, Pollock DD (2011) Repetitive elements may comprise over two-thirds of the human genome. Plos Genet 7:e1002384

Chapter 20 Magnetically Single-Cell Virus Stamping Rajib Schubert Abstract Single-cell engineering via virus based genetic manipulation allows the possibility of understanding of complex tissues. However, current delivery methods for the genetic engineering of single cells via viral transduction suffer from limitations that restrict their application. Here I present a protocol describing a precise technique which can be used for the targeted virus infection of single cells in a monolayer of cells that is optically accessible. The protocol, demonstrated here by stamping cultured Hela cells with lentiviruses (LVs), completes in a few minutes and allows stable transgene expression within a few days, at success rates approaching 80%. Key words Lentivirus, Single cell transduction, Virus stamping, Virus transduction, Magnetic guidance, Optical guidance, Enveloped virus, Animal cell, Cultured neuron, Magnetic nanoparticle, Electromagnet

1

Introduction The heterogeneity in single-cell responses and intracellular interactions has been well established by a number of landmark studies in the last decade [1]. Single-cell analysis allows not only the detection of individual cellular characteristics but allows the correlation of genetic content with phenotypic traits in the same cells [2]. Conventional bulk cell-based assays reflect average responses which drown out important information from underrepresented cells and have vital consequences for therapy [3]. To address these issues, it is crucial to be able to access and manipulate single cells by regulating their gene expression [4]. Over the past several years, viruses have emerged as one of the preeminent methods for the delivery of genetic material in cell biology and gene therapy [5, 6]. The vast array of engineered viruses offers researchers cell-type-specific tropism and a variety of different expression time-courses and stabilities [7–9]. However, precise control of targeted viral transduction remains challenging [9–11]. To overcome such limitations, we have developed a

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9_20, © Springer Science+Business Media, LLC, part of Springer Nature 2021

321

322

Rajib Schubert

Fig. 1 Virus stamping of a single cell by magnetically guiding virus-bound magnetic nanoparticles (a and b). Cartoon of the key steps and experimental design. (a) Viruses encoding GFP are electrostatically bound to magnetic nanoparticles. (b) After this, a pulled glass pipette is filled with a solution of virus-bound magnetic nanoparticles in PBS. Using a micromanipulator, the capillary is then approached to the target cell using optical guidance. Once the target cell is reached, the electromagnet is turned on and the pulsed electromagnetic field guides the virus-bound magnetic nanoparticles to the membrane of a target cell for infection. After 1 min, the magnet is turned off, the pipette is retracted, and the stamped area monitored for gene expression of eGFP as an example (Take and modified from ref. 12 with permission)

technique termed virus stamping (Fig. 1) [12, 13]. Here we describe magnetically guided virus stamping, a method which provides a targeted gene delivery method for studying and controlling single cells. Virus stamping delivers viruses to single cells through forced physical contact. Specifically, viruses are bound to magnetic nanoparticles using charge based interactions where the strength of the attachment between virus and magnetic nanoparticles is sufficiently strong to keep them attached in solution, and yet is weak enough to allow the virus to get released and infect its target cell when the virus-bound magnetic nanoparticle is brought into contact with its target cell [12]. In this protocol we use optical microscopy to visualize and select the targeted cell. The virus-bound magnetic nanoparticles are then directed into physical contact with the selected cell using magnetic forces (Fig. 1). Although the protocol detailed here describe procedures applied to Hela cell culture, it can be extended for targeting single cells embedded in tissue or live animals by integrating it with deep tissue imaging methods such as shadow imaging [14, 15].

2

Materials Prepare and store all reagents at 4  C (unless otherwise indicated). Diligently follow all waste disposal and safety regulations.

2.1 Biological Materials

1. Wild type Hela cell line. 2. eGFP expressing Hela cell line, 3. Lenti virus encoding tdTomato, titer 9  109 TU/ml.

Magnetically Single-Cell Virus Stamping

2.2

Reagents

323

1. Culture medium for Hela cells: Dulbecco’s high glucose eagle medium (DMEM) supplemented with 10% fetal bovine serum. Store at 20  C until used. 2. Silica coated magnetite core nanoparticles 50–200 nm in diameter. 3. Borosilicate glass capillaries for pulling patch pipettes 5 cm in length. 4. Distilled water.

2.3

Equipment

1. Micropipette puller (see Note 1). 2. Upright microscope fluorescent microscope. 3. Z-deck scanning stage. 4. Software module for microscopy sample holder. 5. Magnetic stand to hold the magnet. 6. 20 water-immersion objective. 7. Image acquisition software. 8. Optical breadboards with sealed holes. 9. ScienceDesk workstations for optical. 10. Cell culture chamber (see Note 2). 11. Micromanipulator. 12. Post and platform for micromanipulator. 13. Dovetail probe holder to fit a wide range of bars and probes. Electromagnet. 14. Digital Gaussmeter (see Note 3). 15. 35 mm diameter petri dish with a glass bottom. 16. 35 mm diameter petri dish with polymer coverslip bottom, low walls and an imprinted 500 μm cell location grid. 17. Mechanical pipettors: 0.1–2 μl, 1–10 μl, 2–20 μl, 10–100 μl, 20–200 μl and 100–1000 μl. Microloader pipette tip 0.5–20 μl, Pipette tips, 0.1–10 μl, 1–200 μl, 50–1250 μl. 18. Water bath at 37  C for cell-culture media. 19. Vortexer. 20. Incubator at 37  C with 5% CO2 and humidity >95%. 21. Reaction tubes safe-lock 0.5 ml, and 2 ml. 22. Borosilicate cover glass of 13 mm diameter. 23. Transparent double-faced adhesive tape 10 m  15 mm. 24. Science wipe tissues. 25. Neodymium boron permanent magnet (see Note 4). 26. Sonicator.

324

Rajib Schubert

27. 0.22 μm syringe filter. 28. Syringe U-40. 29. NAP-5 20ST. 30. Biomixer.

3

Method

3.1 Preparing Virus-Bound Magnetic Nanoparticles

1. Vortex and re-suspend 100 μl of diluted stock solution (0.1 mg/ml) of magnetic nanoparticles in 1 ml fresh sterile filtered PBS in a 1.5 ml microcentrifuge tube. 2. Sonicate resuspended magnetic nanoparticles from step 1 for 30 min at 35 kHz and 37  C. 3. Add 1 μl of Lenti virus suspension of titers ranging from 1  107 to 1  109 TU ml 1 to resuspended magnetic nanoparticles (see Notes 5 and 6). 4. Mix virus-bound magnetic nanoparticles solution by pipetting up and down five times using a 1 ml pipette. 5. Leave the mixture on the Biomixer at 24 rotations per min for 1 h at 25  C. 6. Take the tube off the Biomixer and attach a neodymium boron permanent magnet to the side of the tube using a double-sided tape and leave in this configuration for 1 min. This procedure will trap the virus-bound magnetic nanoparticles to the side of the microcentrifuge tube. 7. Keep magnet held on to 1.5 ml tube, aspirate out supernatant using a 1 ml pipette and resuspend in 1 ml PBS. Repeat this cycle three times to ensure that the unbound virus is washed off. 8. Take magnet off and resuspend nanoparticles in 1 ml of fresh sterile filtered PBS. 9. Remove aggregated virus-bound nanoparticles. Run virusbound magnetic nanoparticles through an NAP-5 desalting column to remove large aggregates.

3.2 Calibrating the Cell Culture Chamber

1. Turn on the temperature control of the chamber to 37  C. Switch on microscope controlling software.

3.3 Calibrating the Magnet for Nanoparticle Pullout and Assessing Gene Expression

1. Cultured Helas for 1 day before experiment in a 35 mm diameter petri dish. Place the dish on the microscope dish holder.

2. Set up a 95%) [14]. However, any other cell culture chamber providing cell culture-like conditions for optical imaging and manipulating cells may be used. 3. Take a Gaussmeter and measure the magnetic field of electromagnet at the bottom of the petri dish. The magnetic field should be adjustable and reach at least 40 mT at the pipette tip. Ensure that the magnet is aligned parallel to the pipette tip and that the maximum strength of the magnetic field is reached at the pipette tip. Note that the strength and gradient of the magnetic field depend on the geometry of the magnet and should be measured with the Gaussmeter. 4. We recommend using Supermagnet, Cat. No. S-05-05-N. 5. Thaw virus shortly before usage and keep at 4  C. The maximum storage time for a virus is 1 month at 4  C. Verify your viruses can infect cells robustly. In this protocol, we use Lentivirus encoding tdTomato. Verify infectivity of Lentis by injecting 1 μl of virus in a 35 mm diameter petri dish with a confluent layer of ~1  106 Hela cells. Wait >24 h postinfection to monitor tdTomato expression and the number of cells infected. Suitable virus preparations should infect >80% of the cells (Fig. 3). 6. Note, local gene regulatory and biosafety laws must be followed for each virus type used. For our lab, the engineered Lentis have been classified as biosafety level 2 based on regulations enforced in Switzerland. However, when handling

Magnetically Single-Cell Virus Stamping

327

Fig. 3 Quality control of virus stamping. Pullout of virus-bound magnetic nanoparticles in the presence of a magnetic field of 120 mT. The virus was encoding tdTomato(in red). Scale bar, 200 μm

biohazardous material, following higher biosafety levels if necessary. When switching to other viruses please follow your biosafety rules and regulations regarding experimental handling. 7. We recommend a seeding density of ~1  105 cells per dish. Maximum culture time for 5 days for stamping experiments. Change media every 2 days. 8. Visually check your pipette tip with the 20 objective with wide field imaging to approximate the bore diameter opening and to ensure the tip is not clogged. Maximum storage time for freshly pulled pipettes is 6 h at room temperature. Pull fresh pipettes if time stored exceeds 6 h. 9. In parallel to doing your experiment, you can assess the quality of the virus-bound nanoparticle preparation by injecting 1 μl of virus-bound magnetic nanoparticles into a 35 mm diameter petri dish plated with a confluent layer of ~1  106 cells. Wait for 24 h and assess infectivity by monitoring gene expression of tdTomato (in case of using tdTomato encoding viruses). A good virus preparation should have >80% of the cells infected (Fig. 3).

328

Rajib Schubert

Acknowledgments This study was supported by a Burroughs Wellcome Fund grant (ID #1018793 to R.S.) Competing financial interests: The author declares competing financial interests. The author applied for a patent related to the virus stamping approach. References 1. Hodzic E (2016) Single-cell analysis: advances and future perspectives. Bosn J Basic Med Sci 16:313–314 2. Di Carlo D, Tse HT, Gossett DR (2012) Introduction: why analyze single cells? Methods Mol Biol 853:1–10 3. Stuart T, Satija R (2019) Integrative single-cell analysis. Nat Rev Genet 20(5):257–272 4. Fugger L, Friese MA, Bell JI (2009) From genes to function: the next challenge to understanding multiple sclerosis. Nat Rev Immunol 9(6):408–417 5. Akhtar A et al (2011) A decade of molecular cell biology: achievements and challenges. Nat Rev Mol Cell Biol 12(10):669–674 6. Chari R, Church GM (2017) Beyond editing to writing large genomes. Nat Rev Genet 18 (12):749–760 7. Lotze MT, Kost TA (2002) Viruses as gene delivery vectors: application to gene function, target validation, and assay development. Cancer Gene Ther 9(8):692–699 8. Wang D et al (2019) Adeno-associated virus vector as a platform for gene therapy delivery. Nat Rev Drug Discov 18(5):358–378 9. Belousova N et al (2002) Modulation of adenovirus vector tropism via incorporation of

polypeptide ligands into the fiber protein. J Virol 76(17):8621–8631 10. Uchida E et al (2007) Optimization of the virus concentration method using polyethyleneimine-conjugated magnetic beads and its application to the detection of human hepatitis A, B and C viruses. J Virol Methods 143(1):95–103 11. Walther W, Stein U (1996) Cell type specific and inducible promoters for vectors in gene therapy as an approach for cell targeting. J Mol Med (Berl) 74(7):379–392 12. Schubert R et al (2019) Magnetically guided virus stamping for the targeted infection of single cells or groups of cells. Nat Protoc 14 (11):3205–3219 13. Schubert R et al (2018) Virus stamping for targeted single-cell infection in vitro and in vivo. Nat Biotechnol 36(1):81–88 14. Judkewitz B et al (2009) Targeted single-cell electroporation of mammalian neurons in vivo. Nat Protoc 4(6):862–869 15. Kitamura K et al (2008) Targeted patch-clamp recordings and single-cell electroporation of unlabeled neurons in vivo. Nat Methods 5 (1):61–67

INDEX A AdoB12....................................... 90, 91, 94, 98, 102–104 Agonistic aptamer ......................................................... 301 Allylic dealkylation ........................................................ 287 Amplicon sequencing.......................................... 196, 197, 213–215, 219, 221 Animal cells.................................................................... 301 Antibodies ..............................................3, 5, 7, 8, 13, 16, 24, 59–72, 113, 121, 160, 261, 262, 265, 267, 269, 270, 273, 275, 283 Artificial metalloenzyme (ArMs) ........................ 287–299, 310, 312

B Base editing ................................................. 194, 195, 198 Bioengineering .............................................................. 238 Biomolecular condensates ................................... 253–255 Biotechnology ................................................................. 15

C Caffeine..............................................16, 18, 38, 160–167 CarH ................................................................... 90, 91, 98 CAR-T cell................................................................... 3–12 CAR-T cell activation assay ........................................ 8–10 Cas12a .................................................................. 171–191 Cell death signaling...................................................60, 61 Cell-penetrating polydisulfides (CPD) .......................289, 290, 293–296 Cell sorting ................................................................74, 75 Cell therapies ..................................................................... 3 Chaperones.................................................................... 277 Chemical biology .......................................................... 237 CRISPR ........................................ 53, 171, 172, 193–221 CRISPR array synthesis ................................................ 171 CRISPR-Cas9....................................................... 225, 309 CRISPR-Cas12a............................................................ 189 Cultured neuron ........................................................... 321

D De novo gene targeting ....................................... 309–319 Designer cells ..............................................37–39, 42–47, 51, 52, 54, 126, 127 Destabilizing domains .................................................. 278 Developmental biology................................................. 309

Diabetes ...................................................... 35–38, 42, 44, 54, 141, 142, 160, 161 Diabetes mellitus ..............................................35, 54, 141 Differentiation........................................... 36, 52, 73, 142 DNA aptamers............................................. 301, 302, 304 Dynabeads ......................................................................... 5

E Electromagnet ............................................. 322, 323, 325 Engineered cells ........................................ 37, 39, 60, 143 Enveloped virus ............................................................. 321 Epitopes ........................................................16, 22, 59–61 ERK ............................................ 240–242, 247–248, 250 Escherichia coli dihydrofolate reductase (eDHFR)...... 238

F Far-red light (FRL) .............................. 37, 38, 42–44, 49, 50, 53, 126, 127, 130, 135, 137, 138, 142, 145, 147, 156 Fibroblast growth factors (FGF).................................. 301 Flow cytometry ...............8, 11, 13, 31, 68, 70, 281, 282

G Gene editing ........................................126, 172, 174, 190 Gene switching................................42, 89–105, 159–167 Genetics ............................................. 3, 51, 52, 110, 126, 128, 132, 135, 138, 157, 159, 278, 309 Genome editing ........................................... 53, 173, 174, 177, 186–190, 193–221, 225–233 Genome engineering ........................................... 171–191 Genomic PCR ...................................................... 231, 232 Glucose homeostasis ............................... 35–55, 141–157 Green light ........................................................90, 91, 98, 99, 102, 103, 105 Guide RNA (gRNA) .................................. 172, 193–199, 202–213, 217, 220, 225, 311, 316–317, 319

H High-throughput sequencing ...................................... 193 Homology-directed repair (HDR) .............................225, 226, 230–232 Human induced pluripotent stem cells (hiPSCs) .................................................... 302–304 Hydrogels ..................................... 90, 143, 156, 253–274

Ryosuke Kojima (ed.), Mammalian Cell Engineering: Methods and Protocols, Methods in Molecular Biology, vol. 2312, https://doi.org/10.1007/978-1-0716-1441-9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

329

MAMMALIAN CELL ENGINEERING: METHODS

330 Index

AND

PROTOCOLS

I

R

Induced production of ligand(light)-yielded multivalent enhancers (iPOLYMER) ..... 254–267, 272, 273 Inducible dimerization ................................................. 254 Inducible expression .............................................. 42, 159 Intracellular catalysis ............................................ 290, 291 In utero electroporation ..............................310, 316–318

Raf .................................................................................. 240 Receptor agonists .......................................................... 301 Receptor engineering.................................................... 160 Receptors ............................................................. 3, 15–20, 22–24, 28, 29, 32, 60, 90, 126, 160–162, 164, 167, 290, 301 Regenerative medicine ........................................... 73, 142 Retronectin ................................................ 5, 7, 61, 64, 66 Ruthenium complex ........................................... 288, 290, 293, 294, 299

L Lentivirus vector ............................................................... 3 Library screening............................................................. 60 Light-controlled designer cells..................................... 125 Light-switchable gene expression .................................. 89

M Magnetic guidance ........................................................ 322 Magnetic nanoparticles .......................322, 324, 325, 327 Mammalian cells.......................................................15–32, 35–55, 59–72, 89–105, 109–122, 125–138, 145, 159–167, 172, 174, 177, 179, 186–188, 190, 193–221, 226, 237–250, 253–275, 290, 292, 299 Mammalian synthetic biology ................ 15, 16, 159–161 Membrane proteins...................................................59, 62 Microbubbles ................................................................ 110 MicroRNAs (miRNAs) ............................... 74, 78, 84, 85 Multiplexed ......................................... 171–191, 193–221

N Nonhomologous end joining (NHEJ) .............. 190, 225, 226, 232 Noninvasive therapy...................................................... 109

O Optical guidance ........................................................... 322 Optogenetics ............................................ 50, 90, 91, 104, 125–137, 142, 226, 231, 232 Orthogonal genome engineering ...................... 172, 173, 177, 179

P PEG precipitation ............................................................. 3 Phase separation ................................................... 105, 253 Plant cells .................................................................89–106 Plasma membrane ................................................ 238, 247 Prestin ...........................................................110–113, 122 Proteasome .................................................................... 278 Protein aggregates................................................ 277–284 Protein engineering ............................161, 194, 195, 226 Protein localization .............................................. 237–250

S Self-localizing ligand (SL) ................................... 238, 239 Signal transduction ................................................ 16, 161 Single cell transduction................................................. 321 Single-chain Fv (scFv).................. 3, 7, 12, 59–62, 68–72 SL-induced protein translocation (SLIPT)..................................................... 237–250 Smartphone-controlled cells........................125–138, 143 Sonogenetics......................................................... 109–122 Stem cells .................................................... 36, 52, 73–86, 142, 167, 196, 301–305 Streptavidin........................................................... 288, 290 Stress granules ............................................ 254, 255, 257, 259–262, 265, 267, 269, 270, 273–275 Synthetic biology .......................................................3, 90, 138, 157, 159, 160, 240, 254, 256, 257, 287 Synthetic designer cells ................................................. 141 Synthetic mRNA .......................................................73–85 Synthetic receptors....................................................15–31

T T7E1 assay..................................................................... 225 Telemedicine ................................................................. 142 Thyroid hormone.......................................................... 289 Transcriptional regulation .................................. 174, 177, 186, 188–190 Transcription factors .......................................50, 98, 126, 135, 145, 160–162, 166, 171 Trimethoprim ....................................................... 238, 245

U Ultrasound..................................................................... 109

V Virus stamping ..................................................... 321–328 Virus transduction......................................................... 321 Visualization of protein sub-cellular localization......................................................... 309