Stem Cells and Lineage Commitment: Methods and Protocols (Methods in Molecular Biology, 2736) 1071635360, 9781071635360

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Methods in Molecular Biology 2736

Kursad Turksen  Editor

Stem Cells and Lineage Commitment 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.

Stem Cells and Lineage Commitment Methods and Protocols

Edited by

Kursad Turksen Ottawa, ON, Canada

Editor Kursad Turksen Ottawa, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-0716-3536-0 ISBN 978-1-0716-3537-7 (eBook) https://doi.org/10.1007/978-1-0716-3537-7 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Illustration Caption: Artwork created by Dr. Kursad Turksen. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface Studies in the area of lineage commitment are one of the major research areas. Over the years, many protocols have been developed, improved, and extended to become more directed and consequently more informative. In this volume, I have collected a series of protocols that cover diverse aspects of the topic and represent some of the developments and improvements in the stem cell field. Once again, the protocols gathered here are faithful to the mission statement of the Methods in Molecular Biology series: They are well-established and described in an easy-tofollow, step-by-step fashion so as to be valuable for not only experts but also for novices in the stem cell field. That goal is achieved because of the generosity of the contributors who have carefully described their protocols in this volume, and I am very grateful for their efforts. My thanks as well go to Dr. John Walker, the Editor-in-Chief of the Methods in Molecular Biology series, for giving me the opportunity to create this volume and for supporting me along the way. I am also grateful to Patrick Marton, the Executive Editor of Methods in Molecular Biology and the Springer Protocols collection, for his continuous support from idea to completion of this volume. A special thank you goes to Anna Rakovsky, Assistant Editor for Methods in Molecular Biology, for her support from the beginning to the end of this project. I would also like to thank David C. Casey, the Senior Editor of Methods in Molecular Biology, for his outstanding editorial work during the production of this volume. Finally, I would like to thank the production crew for their work in putting together an outstanding volume. Ottawa, ON, Canada

Kursad Turksen

v

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

v ix

Lineage Tracing by Single-Cell Transcriptomics Decoding Dynamics of Lineage Commitment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Ping Yu and Lin Cheng Cleavage Under Targets & Release Using Nuclease (CUT&RUN) of Histone Modifications in Epidermal Stem Cells of Adult Murine Skin . . . . . . . . . . . 9 Pooja Flora and Elena Ezhkova Targeted Gene Silencing by Using GapmeRs in Differentiating Human-Induced Pluripotent Stem Cells (hiPSC) Toward Pancreatic Progenitors . . . 23 Lucas Unger, Luiza Ghila, and Simona Chera Retinoic Acid-Mediated Differentiation of Mouse Embryonic Stem Cells to Neuronal Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 Sangeeta Dutta, Debosree Pal, and M. R. S. Rao Isolation and Functional Analysis of Myoepithelial Cells from Adult Mouse Submandibular Glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 Rika Yasuhara, Seya Kang, Rino Tokumasu, and Kenji Mishima Chromosomal Analysis in Lineage-Specific Mouse Hematopoietic Stem Cells and Progenitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65 Nur Afizah Yusoff, Zariyantey Abd Hamid, Paik Wah Chow, Salwati Shuib, Izatus Shima Taib, and Siti Balkis Budin Easy and Rapid Methods for Human Umbilical Cord Blood–Derived Mesenchymal Stem Cells and Human Umbilical Wharton’s Jelly–Derived Mesenchymal Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Figen Abatay Sel, Ayse Erol, Mediha Suleymanoglu, Durdane Serap Kuruca, and Fatma Savran Oguz Encapsulation of MSCs in PRP-Derived Fibrin Microbeads . . . . . . . . . . . . . . . . . . . . . . 85 ¨ zge Lalegu € lker, S¸u ¨ l-U ¨ kran S¸eker, Ays¸e Eser Elc¸in, and Yas¸ar Murat Elc¸in O Assessing Neuronogenic Versus Astrogenic Bias of Neural Stem Cells Via In Vitro Clonal Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Laura Rigoldi and Antonello Mallamaci Stem Cell-Based Modeling Protocol for Parkinson’s Disease. . . . . . . . . . . . . . . . . . . . . . 105 Babak Arjmand, Shayesteh Kokabi-Hamidpour, Hamid Reza Aghayan, Sepideh Alavi-Moghadam, Rasta Arjmand, Mostafa Rezaei-Tavirani, Parisa Goodarzi, Ensieh Nasli-Esfahani, and Mohsen Nikandish Standard Operating Procedure for Production of Mouse Brown Adipose Tissue-Derived Mesenchymal Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Babak Arjmand, Mostafa Rezaei-Tavirani, Sepideh Alavi-Moghadam, Akram Tayanloo-Beik, Mahdi Gholami, Shayesteh Kokabi-Hamidpour, Rasta Arjmand, Ahmad Rezazadeh-Mafi, Fereshteh Mohamadi-jahani, and Bagher Larijani

vii

viii

Contents

Development and Validation of Type 2 Diabetic Zebrafish Model for Cell-Based Treatments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Babak Arjmand, Sepideh Alavi-Moghadam, Shayesteh Kokabi-Hamidpour, Rasta Arjmand, Mostafa Rezaei-Tavirani, Bagher Larijani, Parisa Goodarzi, Neda Mehrdad, and Mohsen Rajaeinejad Neuromuscular Junction-on-a-Chip for Amyotrophic Lateral Sclerosis Modeling . . . Sepideh Alavi-Moghadam, Shayesteh Kokabi-Hamidpour, Mostafa Rezaei-Tavirani, Bagher Larijani, Rasta Arjmand, Fakher Rahim, Ahmad Rezazadeh-Mafi, Hossein Adibi, and Babak Arjmand Primary Human Leukemia Stem Cell (LSC) Isolation and Characterization . . . . . . . . Neslihan Meric¸ and Fatih Kocabas¸ GMP-Compliant Mesenchymal Stem Cell-Derived Exosomes for Cell-Free Therapy in Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Babak Arjmand, Sepideh Alavi-Moghadam, Mostafa Rezaei-Tavirani, Shayesteh Kokabi-Hamidpour, Rasta Arjmand, Kambiz Gilany, Mohsen Rajaeinejad, Fakher Rahim, Nazli Namazi, and Bagher Larijani Establishing Brain Tumor Stem Cell Culture from Patient Brain Tumors and Imaging Analysis of Patient-Derived Xenografts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elham Mahmoudian and Arezu Jahani-Asl An Optimized Protocol for piggyBac-Induced iPSC Generation from hPBMCs by Automatic Electroporation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pelin Kilic and Begum Cosar Signaling Pathways in Trans-differentiation of Mesenchymal Stem Cells: Recent Advances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vaishak Kaviarasan, Dikshita Deka, Darshini Balaji, Surajit Pathak, and Antara Banerjee Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

127

139

151

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193

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225

Contributors FIGEN ABATAY SEL • Istanbul University, Istanbul Faculty of Medicine, Department of Medical Biology, Istanbul, Turkey; Department of Medical Biology, Istanbul University, Institute of Graduate Studies in Health Science, Istanbul, Turkey ZARIYANTEY ABD HAMID • Centre for Diagnostic, Therapeutic and Investigative Studies, Faculty of Health Sciences, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia HOSSEIN ADIBI • Diabetes Research Center, Endocrinology and Metabolism Clinical Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran HAMID REZA AGHAYAN • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran SEPIDEH ALAVI-MOGHADAM • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran BABAK ARJMAND • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran; Iranian Cancer Control Center (MACSA), Tehran, Iran RASTA ARJMAND • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran DARSHINI BALAJI • Department of Medical Biotechnology, Faculty of Allied Health Sciences, Chettinad Academy of Research and Education (CARE), Chettinad Hospital and Research Institute (CHRI), Chennai, India ANTARA BANERJEE • Department of Medical Biotechnology, Faculty of Allied Health Sciences, Chettinad Academy of Research and Education (CARE), Chettinad Hospital and Research Institute (CHRI), Chennai, India SITI BALKIS BUDIN • Centre for Diagnostic, Therapeutic and Investigative Studies, Faculty of Health Sciences, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia LIN CHENG • Shanghai Institute of Hematology, State Key Laboratory of Medical Genomics, National Research Center for Translational Medicine at Shanghai, Ruijin Hospital Affiliated to Shanghai Jiao Tong University School of Medicine, Shanghai, China SIMONA CHERA • Mohn Research Center for Diabetes Precision Medicine, Department of Clinical Science, Faculty of Medicine, University of Bergen, Bergen, Norway PAIK WAH CHOW • Centre for Diagnostic, Therapeutic and Investigative Studies, Faculty of Health Sciences, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia; All Life Advance Immunology Sdn. Bhd., Petaling Jaya, Selangor, Malaysia BEGUM COSAR • Hu¨creCELL® Biotechnology Development and Commerce, Inc., Ankara, Turkey; Department of Molecular Biology and Genetics, Institute of Science, Bas¸kent University, Ankara, Turkey DIKSHITA DEKA • Department of Medical Biotechnology, Faculty of Allied Health Sciences, Chettinad Academy of Research and Education (CARE), Chettinad Hospital and Research Institute (CHRI), Chennai, India SANGEETA DUTTA • Molecular Biology and Genetics Unit, Jawaharlal Nehru Centre for Advanced Scientific Research, Bangalore, India

ix

x

Contributors

AYS¸E ESER ELC¸IN • Ankara University Faculty of Science, and Ankara University Stem Cell Institute, Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara, Turkey AYSE EROL • Istanbul University, Istanbul Faculty of Medicine, Department of Medical Biology, Istanbul, Turkey ELENA EZHKOVA • Black Family Stem Cell Institute, Department of Cell Development and Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA POOJA FLORA • Black Family Stem Cell Institute, Department of Cell Development and Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA LUIZA GHILA • Mohn Research Center for Diabetes Precision Medicine, Department of Clinical Science, Faculty of Medicine, University of Bergen, Bergen, Norway MAHDI GHOLAMI • Department of Toxicology & Pharmacology, Faculty of Pharmacy; Toxicology and Poisoning Research Center, Tehran University of Medical Sciences, Tehran, Iran KAMBIZ GILANY • Integrative Oncology Department, Breast Cancer Research Center, Motamed Cancer Institute, ACECR, Tehran, Iran; Reproductive Immunology Research Center, Avicenna Research Institute, ACECR, Tehran, Iran PARISA GOODARZI • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran AREZU JAHANI-ASL • Department of Cellular and Molecular Medicine, University of Ottawa, Ottawa, ON, Canada; Brain and Mind Research Institute, University of Ottawa, Ottawa, ON, Canada; Ottawa Hospital Research Institute, Ottawa, ON, Canada SEYA KANG • Division of Pathology, Department of Oral Diagnostic Sciences, School of Dentistry, Showa University, Tokyo, Japan VAISHAK KAVIARASAN • Department of Medical Biotechnology, Faculty of Allied Health Sciences, Chettinad Academy of Research and Education (CARE), Chettinad Hospital and Research Institute (CHRI), Chennai, India PELIN KILIC • Department of Stem Cells and Regenerative Medicine, Stem Cell Institute, Ankara University, Ankara, Turkey; Hu¨creCELL® Biotechnology Development and Commerce, Inc., Ankara, Turkey FATIH KOCABAS¸ • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Tu¨rkiye SHAYESTEH KOKABI-HAMIDPOUR • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran DURDANE SERAP KURUCA • Istanbul University, Istanbul Faculty of Medicine, Department of Physiology, Istanbul, Turkey; Istanbul Atlas University, Faculty of Medicine, Department of Physiology, Istanbul, Turkey ¨ ZGE LALEGU¨L-U € LKER • Ankara University Faculty of Science, and Ankara University Stem O Cell Institute, Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara, Turkey BAGHER LARIJANI • Endocrinology and Metabolism Research Center, Endocrinology and Metabolism Clinical Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran ELHAM MAHMOUDIAN • Department of Cellular and Molecular Medicine, University of Ottawa, Ottawa, ON, Canada; Brain and Mind Research Institute, University of Ottawa, Ottawa, ON, Canada

Contributors

xi

ANTONELLO MALLAMACI • Laboratory of Cerebral Cortex Development, Department of Neuroscience, SISSA, Trieste, Italy NEDA MEHRDAD • Elderly Health Research Center, Endocrinology and Metabolism Population Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran NESLIHAN MERIC¸ • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Tu¨rkiye; Faculty of Engineering and Life Sciences, Ku¨tahya Health Sciences University, Ku¨tahya, Tu¨rkiye KENJI MISHIMA • Division of Pathology, Department of Oral Diagnostic Sciences, School of Dentistry, Showa University, Tokyo, Japan FERESHTEH MOHAMADI-JAHANI • Brain and Spinal Cord Injury Research Center, Neuroscience Institute Tehran University of Medical Sciences, Tehran, Iran YAS¸AR MURAT ELC¸IN • Ankara University Faculty of Science, and Ankara University Stem Cell Institute, Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara, Turkey; Biovalda Health Technologies, Inc., Ankara, Turkey NAZLI NAMAZI • Diabetes Research Center, Endocrinology and Metabolism Clinical Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran ENSIEH NASLI-ESFAHANI • Diabetes Research Center, Endocrinology and Metabolism Clinical Sciences Institute, Tehran University of Medical Sciences, Tehran, Islamic Republic of Iran MOHSEN NIKANDISH • AJA Cancer Epidemiology Research and Treatment Center (AJACERTC), AJA University of Medical Sciences, Tehran, Iran DEBOSREE PAL • Molecular Biology and Genetics Unit, Jawaharlal Nehru Centre for Advanced Scientific Research, Bangalore, India; UCL Cancer Institute, University College London, London, UK SURAJIT PATHAK • Department of Medical Biotechnology, Faculty of Allied Health Sciences, Chettinad Academy of Research and Education (CARE), Chettinad Hospital and Research Institute (CHRI), Chennai, India FAKHER RAHIM • Health Research Institute, Thalassemia and Hemoglobinopathies Research Center, Ahvaz Jundishapur University of Medical Sciences, Ahvaz, Iran; Department of Anesthesia, Cihan University- Sulaimaniya, Kurdistan Region, Iraq MOHSEN RAJAEINEJAD • AJA Cancer Epidemiology Research and Treatment Center (AJACERTC), AJA University of Medical Sciences, Tehran, Iran M. R. S. RAO • Molecular Biology and Genetics Unit, Jawaharlal Nehru Centre for Advanced Scientific Research, Bangalore, India MOSTAFA REZAEI-TAVIRANI • Proteomics Research Center, Shahid Beheshti University of Medical Sciences, Tehran, Iran AHMAD REZAZADEH-MAFI • Clinical Oncologist, Imam Hossein Hospital, Shahid Beheshti University of Medical Sciences, Tehran, Iran; Department of Radiation Oncology, Imam Hossein Hospital, Shaheed Beheshti Medical University, Tehran, Iran LAURA RIGOLDI • Laboratory of Cerebral Cortex Development, Department of Neuroscience, SISSA, Trieste, Italy FATMA SAVRAN OGUZ • Istanbul University, Istanbul Faculty of Medicine, Department of Medical Biology, Istanbul, Turkey S¸U¨KRAN S¸EKER • Ankara University Faculty of Science, and Ankara University Stem Cell Institute, Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara, Turkey SALWATI SHUIB • Department of Pathology, Faculty of Medicine, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia

xii

Contributors

MEDIHA SULEYMANOGLU • Istanbul University, Istanbul Faculty of Medicine, Department of Medical Biology, Istanbul, Turkey IZATUS SHIMA TAIB • Centre for Diagnostic, Therapeutic and Investigative Studies, Faculty of Health Sciences, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia AKRAM TAYANLOO-BEIK • Cell Therapy and Regenerative Medicine Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of Medical Sciences, Tehran, Iran RINO TOKUMASU • Division of Pathology, Department of Oral Diagnostic Sciences, School of Dentistry, Showa University, Tokyo, Japan LUCAS UNGER • Mohn Research Center for Diabetes Precision Medicine, Department of Clinical Science, Faculty of Medicine, University of Bergen, Bergen, Norway RIKA YASUHARA • Division of Pathology, Department of Oral Diagnostic Sciences, School of Dentistry, Showa University, Tokyo, Japan PING YU • Shanghai Institute of Hematology, State Key Laboratory of Medical Genomics, National Research Center for Translational Medicine at Shanghai, Ruijin Hospital Affiliated to Shanghai Jiao Tong University School of Medicine, Shanghai, China NUR AFIZAH YUSOFF • Centre for Diagnostic, Therapeutic and Investigative Studies, Faculty of Health Sciences, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia

Methods in Molecular Biology (2023) 2736: 1–7 DOI 10.1007/7651_2022_476 © Springer Science+Business Media, LLC 2023 Published online: 08 February 2023

Lineage Tracing by Single-Cell Transcriptomics Decoding Dynamics of Lineage Commitment Ping Yu and Lin Cheng Abstract Tracing the fate of individual cells and their progeny is necessary and significant for stem cell research and cancer research. Changes in lineage-specific transcription factor levels during lineage commitment are gradual and continuous. Development of single-cell sequencing technology allows many different states of cells to be sequenced at an unprecedented resolution, and it has been proved that single-cell transcriptomics meets lineage tracing. Here, we introduce a detailed protocol for the lineage tracing by single-cell transcriptomics to clarify the dynamics of lineage commitment. Key words Cell transcriptomics

1

fate,

Development,

Lineage

commitment,

Lineage

tracing,

Single-cell

Introduction Multipotent stem cells can differentiate into diverse cell types through cell fate choices during the development of cells. Cells gradually switch along an ordered series of states during a continuous process of differentiation. Reconstructing the lineage relationship among these different cell types not only provides significant insights into the fundamental processes of normal tissue development but also contributes to explain the pathogenesis of cancers. Lineage tracing is an important method of genetic labeling that allows us to follow the fate of individual cells and their progeny, which has been applied to stem cell research and cellular heterogeneity in cancers [1]. Traditional methods of lineage tracing are mainly imaging based, such as direct observation, labeling cells with dyes and radioactive tracers, and introduction of genetic markers by transfection or viral transduction, but they are limited by their scalability and the lack of molecular information underlying fate transitions [1, 2]. Currently, lineage tracing is primarily divided into two strategies naming prospective tracing and retrospective tracing [3, 4]. The former requires the introduction of exogenous markers into cells, including integration barcode, polylox barcode, CRISPR

1

2

Ping Yu and Lin Cheng

barcode [5–8], and their use is restricted in vitro studies or model organisms because they are impossible to implement in a completely wild-type setting like in vivo human studies. The latter needs endogenous markers containing nuclear genome changes including single-nucleotide variants (SNVs), copy number variations (CNVs), retrotransposon elements, and repeat regions like microsatellites repeat and mitochondrial genome changes [9– 12]. Retrospective tracing can be applied to trace cell change in the context of human development and cancers due to the universality and naturalness of mutation occurring. However, the method has a major limitation that some mutation may be rare, which may be hard to be applied for the construction of the lineage relationship. An alternative solution to this limitation is to combine several methods to increase the accuracy and reliability of lineage tracing. With the development of sequencing, single-cell RNA sequencing (scRNA-seq) can perform the profiling of thousands of individual cells and the identification of cell types at an unprecedented resolution simultaneously. Many cells at different development stages can be sequenced by scRNA-seq and cells with similar transcriptomes will cluster together along the developmental trajectory, which provide a possibility for trajectory reconstruction at the single cell level [4]. Recently, two main trajectory reconstruction algorithms (monocle and RNA velocity) are widely used [13–16]. In this chapter, we will describe a simple step-by-step pipeline about tracking the lineage through scRNA-seq to reveal the dynamics of lineage commitment (Fig. 1).

Fig. 1 Schematic representation of the lineage tracing using scRNA-seq

Lineage Tracing by Single-Cell Transcriptomics Decoding Dynamics of Lineage. . .

2

3

Materials

2.1 Reagents and Equipment

1. Cellometer K2 Fluorescent Viability Cell Counter (Nexcelom). 2. Hemocytometer. 3. 10× Chromium Controller (10× Genomics). 4. Chromium Single Cell 3′ v2 reagent kit (10× Genomics). 5. The Illumina NovaSeq 6000 (Illumina).

2.2

Software

1. Cell Ranger v6.02 (10× Genomics, https://support.10 xgenomics.com/single-cell-gene-expression/software/ downloads/6.0/). 2. R v4.1.3 (The R Foundation, https://www.r-project.org). 3. Python v3.7.9 (https://www.python.org/). 4. Seurat v4.0.1 (Satija Lab, https://satijalab.org/seurat/) [17]. 5. DoubletFinder v2.0.3 (https://github.com/chris-mcginnisucsf/DoubletFinder) [18]. 6. clustree v0.5.0 (https://github.com/lazappi/clustree) [19]. 7. SingleR v1.4.1 (https://github.com/dviraran/SingleR) [20]. 8. Monocle v2.12.0 (http://cole-trapnell-lab.github.io/mono cle-release/) [16]. 9. Velocyto v0.17.17 (http://velocyto.org/velocyto.py/index. html) [14]. 10. scVelo v0.2.3 (https://scvelo.readthedocs.io/) [21].

3

Methods

3.1 Sample Collection and SingleCell Sequencing

Collected samples are carefully digested, washed, and resuspended to obtain single-cell suspensions (see Note 1). Subsequently, the single cell suspensions are counted using both the Cellometer K2 Fluorescent Viability Cell Counter (Nexcelom) and hemocytometer, and cell number is adjusted to 1000 cells/μL with cell viability greater than 80%. The adjusted cell suspension is loaded into Chromium microfluidic chips with 3′ v2 chemistry and barcoded with a 10× Chromium Controller (10× Genomics). Then, RNA from the barcoded cells is reverse-transcribed. After libraries are constructed with reagents from a Chromium Single Cell 3′ v2 reagent kit (10× Genomics) according to the manufacturer’s instructions, sequencing is performed using the Illumina NovaSeq 6000 sequencing platforms (see Note 2).

4

Ping Yu and Lin Cheng

3.2 Data Preprocessing

The gene-barcode matrices are generated after the raw sequencing reads are mapped to the human reference genome GRCh38 using the “cellranger count” command with default parameters in the Cell Ranger v6.02. The matrices are loaded using the Read10× function from the Seurat v4.0.1 package [17], which will return a unique molecular identified (UMI) count matrix. Then, the CreateSeuratObject function is applied to obtain a Seurat object. The percentage of reads that align to the mitochondrial genome is calculated using the PercentageFeatureSet function with the pattern parameter set “^MT-”. The potential doublets are removed by the DoubletFinder v2.0.3 [18]. Subsequently, genes expressed in more than three cells and cells expressing >500 genes with 50 bp, see Note 14. 16. Use 1 μL of CUT&RUN-enriched DNA using the Qubit™ fluorometer per manufacturer’s instructions, see Note 15. 17. CUT&RUN-enriched DNA can be stored at -20 °C at this point. 3.4 Library Preparation and Quality Assessment

1. We recommend using ~5–10 ng of purified CUT&RUNenriched DNA to prepare Illumina NGS libraries using the NEBNext® Ultra™ II DNA Library Prep Kit for Illumina®. Please follow all manufacturer’s instructions except for the following modifications for each step. 2. Add 1× TE to CUT&RUN-enriched DNA to achieve a final volume of 50 μL before proceeding with End Prep step. 3. For adapter ligation step, prepare a 1:25 dilution of adapter in Tris/NaCl, i.e., add 4 μL of adapter to 96 μL of Tris/NaCl, before adding it to each sample reaction. 4. Use the following cycling parameters for primer indexing: (a) 45 s @ 98 °C. (b) 15 s @ 98 °C. (c) 10 s @ 60 °C. (d) Repeat steps (b)–(c) for a total of 15×. (e) 1 min @ 72 °C. 5. Perform PCR product cleanup using a ratio of 1× AMpure XP beads to sample volume. 6. Elute DNA in 17 μL of 10 mM of 10 mM Tris–HCl or 0.1× TE. DNA libraries can be stored at -20 °C. 7. It is recommended to assess library size, prior to sequencing, on an Agilent Bioanalyzer. The trace should show enrichment of mononucleosome (~150 bp + 125 bp sequence adapters) (Fig. 2).

3.5 Sequencing and Data Analysis

To obtain 10–20 million paired-end reads/sample, constructed libraries can be sequenced on Illumina NextSeq 500/550 platform with 150-cycles, as recommended [19]. Paired-end reads are subjected to quality control and aligned to UCSC mouse genome (GRC38/mm10) using bowtie2 [20]. Experimental normalization should be done using the E. coli spike-in DNA [21, 22]. TDF files can be generated from normalized reads to visualize peaks on genomic regions using Integrative Genomics Viewer (Fig. 3) [23].

Pooja Flora and Elena Ezhkova

a

1455

2000 1500 1000 500 10,000

1500

Size [bp]

100

0 15

Sample Intensity [Normalized FU]

96

CUT&RUN-enriched DNA: IgG

106 2000 1500 1000 500

10,000

Size [bp]

100

0 15

Sample Intensity [Normalized FU]

1432

CUT&RUN-enriched DNA: H3K27me3

b

1500

18

Fig. 2 Quality assessment of CUT&RUN libraries. Agilent Bioanalyzer traces of PCR amplified libraries prepared from 10 ng of CUT&RUN-enriched DNA using (a) IgG (negative control) and (b) H3K27me3 antibody. H3K27me3 libraries are predominantly mononucleosome at ~275 bp (~150 bp mononucleosome +125 bp adapters) Input ChIP-seq

CUT&RUN

H3K27me3 IgG H3K27me3

[0-2] [0-2] [0-2] [0-2]

Hox cluster of genes

Fig. 3 Integrated Genome Viewer (IGV) tracks of H3K27me3. Representative genome browser tracks of two independent runs of ChIP-seq and CUT&RUN with H3K27me3 in EpSCs, respectively. ChIP-seq was carried out with 500,000 FACS-purified EpSCs, while CUT&RUN was conducted using 100,000 FACS-purified EpSCs. Input and IgG serve as negative controls for the ChIP-seq and CUT&RUN method, respectively. Peaks are shown on Hox cluster of genes that are demarcated and repressed by H3K27me3. CUT&RUN shows sharper peaks over the repressed genomic regions when compared to ChIP-seq tracks

Cleavage Under Targets & Release Using Nuclease (CUT&RUN) of Histone. . .

4

19

Notes 1. Preparation of CUT&RUN buffers can be time consuming and, therefore, we recommend preparing stock solutions ahead of the day of the experiment. 2. While most CUT&RUN protocols suggest using 0.01% digitonin, 0.025% digitonin is optimal for keratinocytes. 3. New scalpels are usually very sharp. Therefore, there is a possibility of tearing the skin during scraping of adipose layer. Therefore, care should be taken as to not rip the skin at this stage, as scraping after trypsin–EDTA incubation will be challenging. 4. In the event that the animal was in anagen, the skin will be much thicker. Therefore, 1 h of trypsin–EDTA incubation will not be sufficient. Therefore, after scraping of adipose layer, tissue should be incubated in 0.1% collagenase + DNase for 1 h at 37 °C, followed by trypsin–EDTA incubation for 30 min at 37 °C as described originally. 5. We recommend pre-wetting the walls of 50 mL, 15 mL, and 5 mL tubes used during this part of the procedure with the solution that will be added to the respective tubes prior to using them for cells. For example, wet 50 mL conical tubes with ice-cold E-media and 15 mL conical tubes with ice-cold DPBS. Discard the solution after this step. Also, wet one 100 μm and one 40 μm strainer with E-Media prior to straining cells. This helps in buffering and protecting cell membranes and increases viability. 6. Up to 500,000 EpSCs can be collected in a 1.5 mL centrifuge tube to conduct CUT&RUN with different histone modification antibodies. We recommend using 100,000 cells for IgG and H3K27me, respectively. However, CUT&RUN for other histone modifications may require more cells. Therefore, we suggest collecting in one tube and then dividing cells into desired amount in Subheading 3.3.2. 7. We do not recommend freezing cells at this point. Our protocol is optimized for usage on live epidermal cells following FACS isolation. 8. If more than 500,000 cells are collected, it is not required to add more E-media before centrifugation at this step. 9. If 200,000 cells are collected during FACS isolation, every 100 μL of cell suspension will have 100,000 cells which is sufficient to conduct CUT&RUN with H3K27me3. If more cells are needed for particular histone modifications, please adjust dilution volume accordingly, so that at the end ConA beads + cells volume is no more than 110 μL.

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Pooja Flora and Elena Ezhkova

10. The suggested volume of 1.5 μL of antibody is from our experiments with rabbit mAb tri-methyl-histone H3 (Lys27) (C36B11) (Cell Signaling, 9733S). Therefore, the amount of antibody used needs to be optimized for intended histone modification CUT&RUN. 11. Make sure the cap sides of the tubes are elevated to ensure that the solution stays in the bottom of the tubes. Do not use a rotator. 12. This buffer stops the enzymatic activity of MNase by chelating Ca2+ ions. 13. 0.1 ng of E. coli DNA is used for 100,000 cells for sequencing normalization. This can be increased linearly with increasing cells, but the amount needs to be adjusted to achieve read counts in the optimal range. After sequencing, in addition to alignment with reference genome, reads also need to be aligned to the E. coli K12, MG1655 reference genome. Normalization factor needs to be calculated such that the E. coli spike-in signal is set to be equal across all samples [22]. 14. We recommend using the EpiCypher DNA purification kit for this step. Also, DNA should be eluted with 12 μL of elution buffer provided in the kit. 15. Yields are influenced by a variety of factors including the viability and quality of cells, antibodies, and target abundance. Therefore, the best indicator of success at this stage is that the yield of target of interest, i.e., H3K27me3, is greater than the IgG negative control sample. Usually, 10,000 EpSCs yield 15–20 ng of target DNA while IgG yields 3-6 ng of target DNA. We do not recommend analysis of fragment size distribution using an Agilent Bioanalyzer prior to library preparation.

Acknowledgments This work was supported by NIH grants P30 AR079200 and R01 AR069078 awarded to E.E. References 1. Chen Z, Li S, Subramaniam S, Shyy JYJ, Chien S (2017) Epigenetic regulation: a new frontier for biomedical engineers. Annu Rev Biomed Eng 19(1):195–219 2. Farrelly LA, Thompson RE, Zhao S, Lepack AE, Lyu Y, Bhanu NV et al (2019) Histone serotonylation is a permissive modification that enhances TFIID binding to H3K4me3. Nature 567(7749):535–539

˜ as-Potts C, Matunis MJ (2013) SUMO: 3. Cuben a multifaceted modifier of chromatin structure and function. Dev Cell 24(1):1–12 4. Strahl BD, Allis CD (2000) The language of covalent histone modifications. Nature 403(6765):41–45 5. Gelato KA, Fischle W (2008) Role of histone modifications in defining chromatin structure and function. Biol Chem 389(4):353–363

Cleavage Under Targets & Release Using Nuclease (CUT&RUN) of Histone. . . 6. Kouzarides T (2007) Chromatin modifications and their function. Cell 128(4):693–705 7. Hemberger M, Dean W, Reik W (2009) Epigenetic dynamics of stem cells and cell lineage commitment: digging Waddington’s canal. Nat Rev Mol Cell Biol 10(8):526–537 8. Jambhekar A, Dhall A, Shi Y (2019) Roles and regulation of histone methylation in animal development. Nat Rev Mol Cell Biol 20(10): 625–641 9. Flora P, Ezhkova E (2020) Regulatory mechanisms governing epidermal stem cell function during development and homeostasis. Dev Camb Engl 147(22):dev194100 10. Park PJ (2009) ChIP-Seq: advantages and challenges of a maturing technology. Nat Rev Genet 10(10):669–680 11. Skene PJ, Henikoff S (2017) An efficient targeted nuclease strategy for high-resolution mapping of DNA binding sites. eLife 6:e21856 12. Fuchs E (2016) Epithelial skin biology: three decades of developmental biology, a hundred questions answered and a thousand new ones to address. Curr Top Dev Biol 116:357–374 13. Fuchs E (2018) Skin stem cells in silence, action, and cancer. Stem Cell Rep 10(5): 1432–1438 14. Moltrasio C, Romagnuolo M, Marzano AV (2022) Epigenetic mechanisms of epidermal differentiation. Int J Mol Sci 23(9):4874 15. Katiyar SK, Singh T, Prasad R, Sun Q, Vaid M (2012) Epigenetic alterations in ultraviolet radiation-induced skin carcinogenesis: interaction of bioactive dietary components on

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epigenetic targets. Photochem Photobiol 88(5):1066–1074 16. Shen Y, Stanislauskas M, Li G, Zheng D, Liu L (2017) Epigenetic and genetic dissections of UV-induced global gene dysregulation in skin cells through multi-omics analyses. Sci Rep 7(1):42646 17. Li MY, Flora P, Pu H, Bar C, Silva J, Cohen I et al (2021) UV-induced reduction in Polycomb repression promotes epidermal pigmentation. Dev Cell 56(18):2547–2561.e8 18. Rheinwald JG, Green H (1977) Epidermal growth factor and the multiplication of cultured human epidermal keratinocytes. Nature 265(5593):421–424 19. Meers MP, Bryson TD, Henikoff JG, Henikoff S (2019) Improved CUT&RUN chromatin profiling tools. Parker S, Weigel D, editors. eLife 8:e46314 20. Langmead B, Salzberg SL (2012) Fast gappedread alignment with Bowtie 2. Nat Methods 9(4):357–359 21. Orlando DA, Chen MW, Brown VE, Solanki S, Choi YJ, Olson ER et al (2014) Quantitative ChIP-Seq normalization reveals global modulation of the epigenome. Cell Rep 9(3): 1163–1170 22. Tay RE, Olawoyin O, Cejas P, Xie Y, Meyer CA, Ito Y et al (2020) Hdac3 is an epigenetic inhibitor of the cytotoxicity program in CD8 T cells. J Exp Med 217(7):e20191453 23. Robinson JT, Thorvaldsdo´ttir H, Winckler W, Guttman M, Lander ES, Getz G et al (2011) Integrative genomics viewer. Nat Biotechnol 29(1):24–26

Methods in Molecular Biology (2023) 2736: 23–38 DOI 10.1007/7651_2023_498 © Springer Science+Business Media, LLC 2023 Published online: 25 August 2023

Targeted Gene Silencing by Using GapmeRs in Differentiating Human-Induced Pluripotent Stem Cells (hiPSC) Toward Pancreatic Progenitors Lucas Unger, Luiza Ghila, and Simona Chera Abstract Induced pluripotent stem cells as a source for generating pancreatic islet endocrine cells represent a great research tool for deciphering the molecular mechanisms of lineage commitment, a layered multi-step process. Additionally, targeted gene silencing by using GapmeRs, short antisense oligonucleotides, proved instrumental in studying the role of different developmental genes. Here we describe our approach to induce mTOR silencing by using specific GapmeRs during the differentiation of induced pluripotent stem cells toward pancreatic progenitors. We will describe our current differentiation protocol, the transfection procedure, and the quality control steps required for a successful experiment. Key words GapmeR, Gene silencing, Human iPSC, In vitro differentiation, Islet endocrine cells, Pancreatic endocrine progenitors

1

Introduction In recent years, significant progress has been made in the field of pancreatic beta cell differentiation protocols starting from pluripotent stem cells, with a special focus on optimization and increased functionality of the end-product, the differentiated beta cells [1– 6]. A notable facilitator represents the wide accessibility of pluripotent stem cells, following the discovery of the Yamanaka factors, which led to the relative ease of producing induced pluripotent stem cells from adult differentiated cells [7]. Using this resource, experiments could be conducted more frequently, testing numerous factors, small molecules, and culture conditions, which significantly contributed to the optimization of in vitro differentiation protocols [8–11]. Additionally, the development of complex threedimensional (3D) culture systems [12] bearing a closer resemblance to the natural environment during differentiation, are enhancing the maturation of in vitro differentiated beta cells. These 3D cultures provide more physiologically relevant context, leading to the production of stem cell islets with endocrine cell types that resemble their native counterparts closely in terms of

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Lucas Unger et al.

morphology, molecular landscape, and response to glucose stimulation [13–16]. Better differentiation protocols are also proving to be an additional tool to study the development of pancreatic endocrine cells more conveniently and efficiently than ever before [17, 18]. An important part of studying the complex mechanisms involved in the pancreatic islet development is to demultiplex the layered multistep lineage commitment process [19–21] and study its master regulators at individual level [22–24]. Various strategies are used to study these regulatory genes, including random mutagenesis; siRNA knockdowns; and transient, permanent, or inducible knockout cell lines. While it is often easier to mutate a gene or suppress its function, overexpression or activation can also reveal the function of a gene. One common problem with a lot of these strategies is achieving efficient delivery of the desired genetic material into iPSCs. iPSCderived cells are notoriously difficult to transfect due to their innate resistance to foreign DNA uptake [25, 26]. This low transfection efficiency can limit the success of introducing new genes or modifying existing ones in iPSCs. A common way of improving transfection efficiency is by increasing transfection agents and materials; however, this often leads to increased cell death and reduced cell viability. Depending on the experimental conditions, targeting transiently a gene over a specific stage during differentiation is highly desirable when studying its specific effect. An alternative way includes the utilization of GapmeRs to knock down mRNA from specific genes [27, 28] at specific timepoints along the differentiation protocol. GapmeRs are single-stranded antisense oligonucleotides with a length of 15–16 nucleotides that allow silencing of long noncoding RNA and mRNA [29, 30]. They consist of a central DNA segment, called “gap,” flanked by modified nucleotide called LNA (locked nucleic acids). When a GapmeR binds to its complementary RNA in the nucleus, the central DNA “gap” recruits the RNase H, which will induce a cleavage at the center of the target RNA sequence [31]. The resulting fragments are rapidly degraded by exonucleases, releasing the LNA GapmeR, which will continue to catalyze the degradation of additional RNA molecules [27] providing an efficient mechanism of RNA degradation [32]. Here, we will focus on the critical steps required for targeted gene silencing exemplified by mTOR using specific GapmeRs during the in vitro differentiation of induced pluripotent stem cells toward pancreatic endocrine progenitors.

Gene Silencing in Differentiating Stem Cells

2 2.1

25

Materials Equipment

1. Laminar flow cabinet. 2. Incubator at 37 °C, 5% CO2. 3. Cell counter (for example: NucleoCounter NC-200 from ChemoMetec with Via1-Cassette (#941-0012) using Reagent A100 (#910-0003) and B (#910-0002)). 4. Micropipette set. 5. Vacuum pump. 6. Cell culture microscope (equipped with 10× obj.) 7. Benchtop centrifuge. 8. Eppendorf tubes holder. 9. Real-Time PCR System (Bio-Rad CFX384). 10. Platform Rotator. 11. Swinging Bucket Plate Centrifuge.

2.2

Disposables

1. 100 mm Tissue Culture Dish (#353003, FALCON). 2. 6-Well Tissue Culture Plate (#353046, FALCON). 3. 12-Well Tissue Culture Plate (#353043, FALCON). 4. Costar® 6-well Clear Flat Bottom Ultra-Low Attachment (#3471, Corning). 5. Sterile filter pipette tips. 6. Sterile glass pipettes. 7. Sterile serological pipettes: 5 mL, 10 mL, 25 mL. 8. 15 mL and 50 mL falcon tube. 9. Sterile 1.5 mL Eppendorf tubes. 10. Coverslips 12 mm (#631-1577, VWR). 11. Fisherbrand™ Superfrost™ (#12727307, ThermoFisher).

Plus

Microscope

Slides

12. 6-well AggreWell 400 Microwell plates (Stemcell Technologies, #34425). 2.3

Reagents

1. Basal media (DMEM #31885023, DMEM/F12 #11330032, ThermoFisher). 2. mTeSR medium and supplement (#085850, Stem Cell Technologies). 3. Pen/Strep (#P4333-100ML, Sigma Aldrich). 4. CTS TrypLE Select Enzyme (#A1285901, ThermoFisher). 5. Dulbecco’s Phosphate Buffered Saline 500 mL (#D8537500ML, Sigma).

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6. 2-Mercaptoethanol (#21985023, Gibco). 7. Opti-MEM™ I Reduced Serum Medium (#31985062, Gibco). 8. TE, pH 8,0, RNase-free (#AM9849, Invitrogen). 9. Lipofectamine™ RNAiMAX (#13778075, Invitrogen). 10. QuantiNova SYBR GreenRT-PCR Kit (#208154, QIAGEN). 11. QuantiNova LNA PCR Assay (#249990, QIAGEN). 12. Antisense LNA GapmeRStandard (#339511, QIAGEN). 13. Geltrex™ LDEV-Free Reduced Growth Factor (#A1413202, Gibco). 14. RNeasy Mini Kit (#74104, QIAGEN). 15. Gibco™ CTS™ TrypLE™ Select Enzyme (#12093745, Gibco). 16. Anti-PDX1 antibody (#ab47308, abcam). 17. Anti-FOXA2 antibody (#ab23630, abcam). 18. Alexa Fluor 488® phalloidin (#A12379. ThermoFisher). 19. Bovine Serum Albumin Fraction (#03117332001, Roche). 20. Phosphate buffered saline (PBS) (#E404-100TABS, VWR). 21. ProLong™ Gold Antifade Mountant (#P36930, Invitrogen). 22. Triton ×100 (#85111, ThermoFisher). 23. Paraformaldehyde (#A11313.22, ThermoFisher). 24. DAPI (#D1306, Molecular Probes). 25. Activin A (Peprotech, # 120-14). 26. ALK5inhII M001).

(ENZO/AH

diagnostics,

#

ALX-270-445-

27. Betacellulin (Peprotech, #100-50). 28. BSA (Sigma-Aldrich, #A7030). 29. CHIR-99021(Tocris, #4423). 30. CMRL 1066 Medium (Corning, #15-110-CVR). 31. DMSO (Sigma-Aldrich, #D8418). 32. FGF-7 (Genscript, # Z03047). 33. GC1 (Tocris, #4554). 34. Glucose (Sigma-Aldrich, #G8769). 35. GlutaMAX (Life Technologies, #35050038). 36. GSiXX (Millipore, #565789). 37. hEGF (Peprotech, #AF-100-15). 38. Heparin Sodium Salt (Sigma-Aldrich, #H3149). 39. ITS-X (Thermo-Fisher, #51500056). 40. LDN-193189 (Selleckchem, # S2618).

Gene Silencing in Differentiating Stem Cells

27

41. Lipid Concentrate (Invitrogen, #11905-031). 42. MCDB131 Medium (Life Technologies, #10372-019). 43. N-acetylcysteine (Sigma-Aldrich, #A9165). 44. NaHCO3 (Sigma-Aldrich, #S5761). 45. Nicotinamide (Sigma-Aldrich, #N0636). 46. Pen/Strep (BioNordika, #BN-ECB3001D). 47. Retinoic Acid (Sigma-Aldrich, #R2625). 48. SANT1 (Sigma-Aldrich, #S4572). 49. Sodium Pyruvate (Lonza, 3BE13-115E). 50. TPB (Santa Cruz, #sc-204424). 51. Trace Elements A (Corning, # 25-021-CI). 52. Trace Elements B (Fisher Scientific, #25022CI). 53. Triiodothyronine (T3) (Sigma-Aldrich, #T6397). 54. TrypLE (Thermo-Fisher, #12563029). 55. Vitamin C (Sigma-Aldrich, #A4544). 56. Y-27632 (Selleckchem, #S1049). 57. ZM-447439 (Selleckchem, #S1103). 58. ZnSO4 (Sigma-Aldrich, #Z0251). 59. Anti-Adherence Rinsing Solution (Stemcell Technologies, #07010). 60. mTeSR medium and supplement (#085850, Stem Cell Technologies). 2.4 Reagent Preparation

1. Table 1. Differentiation Medium Mix 1. 2. Table 2. Differentiation Medium Mix 2. 3. Table 3. Differentiation Medium Mix 3.

Table 1 Differentiation Medium Mix 1 Differentiation Mix 1

Final concentration

Amount

MCDB131

n/a

500 mL

GlutaMAX (100×)

1×

5 mL

Glucose

10 mM

450 mg

NaHCO3

1.5 g/L

750 mg

BSA

0.5%

2.5 g

Pen/strep

0.1%

500 μL

Total

n/a

506.5 mL

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Lucas Unger et al.

Table 2 Differentiation Medium Mix 2 Differentiation Mix 2

Final concentration

Amount

MCDB131

n/a

500 mL

GlutaMAX (100×)

1×

5 mL

Glucose

10 mM

450 mg

NaHCO3

1.5 g/L

1250 mg

BSA

0.5%

10 g

ITS-X

0.5×

2.5 mL

Pen/strep

0.1%

500 μL

Total

n/a

508 mL

Table 3 Differentiation Medium Mix 3 Differentiation Mix 3

Final concentration

Amount

MCDB131

n/a

500 mL

GlutaMAX (100×)

1×

5 mL

Glucose

10 mM

130 mg

NaHCO3

1.5 g/L

7–50 mg

BSA

0.5%

10 g

ITS-X

0.5×

2.5 mL

ZnSO4 (10 mM)

10 μM

0.5 mL

Heparin (10 mg/mL)

10 μg/mL

0.5 mL

Pen/strep

0.1%

500 μL

Total

n/a

507.5 mL

4. Table 4. Differentiation Medium Mix 4. 5. Table 5. Differentiation Medium for each stage/day.

3

Methods Here, we present a method for targeted mTOR silencing during the differentiation of hiPSCs toward posterior foregut to pancreatic endoderm. An assessment of cell viability, silencing efficiency, and silencing effect on differentiation efficiency is also included. A scheme illustrating the differentiation steps and the timing of silencing is presented in Fig. 1.

Gene Silencing in Differentiating Stem Cells

29

Table 4 Differentiation Medium Mix 4 Differentiation Mix 4

Final concentration

Amount

CMRL 1066

n/a

500 mL

GlutaMAX (100×)

1×

5 mL

Sodium pyruvate (100 mM)

0.5 mM

2.5 mL

Lipid concentrate (2000×)

1×

0.25 mL

BSA

2%

10 g

ITS-X

0.5×

2.5 mL

ZnSO4 (10 mM)

10 μM

0.5 mL

Heparin (10 mg/mL)

10 μg/mL

0.5 mL

Pen/strep

0.1%

0.5 mL

Trace elements A (2000×)

1×

0.25 mL

Trace elements B (2000×)

1×

0.25 mL

Total

n/a

512.25 mL

Table 5 Differentiation medium for day 1 to 44 Day

Reagent

Stock concentration

Final concentration

Amount

1–3

Differentiation Mix 1

n/a

n/a

7 mL

1–3

Activin A

100 μg/mL

100 ng/mL

7 μL

1

CHIR-99021

3 mM

3 μM

7 μL

2

CHIR-99021

0.3 mM

0.3 μM

7 μL

4–6

Differentiation Mix 1

n/a

n/a

7 mL

4–6

Vitamin C

250 mM

0.25 mM

7 μL

4–6

FGF-7

100 μg/mL

50 ng/mL

3.5 μL

7–8

Differentiation Mix 2

n/a

n/a

7 mL

7–8

S3 Supplement SANT1 (0.625 mM) Retinoic acid (2.5 mM) LDN-193189 (0.25 mM) TPB (0.5 mM)

2500×

1×

2.8 μL

Stage 1:

Stage 2:

Stage 3:

(continued)

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Lucas Unger et al.

Table 5 (continued) Day

Reagent

Stock concentration

Final concentration

Amount

7–8

Vitamin C

250 mM

0.25 mM

7 μL

7–8

FGF-7

100 μg/mL

50 ng/mL

3.5 μL

Stage 4 (2D): 9–10

Differentiation Mix 2

n/a

n/a

7 mL

9–10

S4 Supplement SANT1 (0.625 mM) Retinoic acid (0.25 mM) LDN-193189 (0.5 mM) TPB (2.5 mM)

2500×

1×

2.8 μL

9–10

Vitamin C

250 mM

0.25 mM

7 μL

9–10

FGF-7

2 μg/mL

2 ng/mL

7 μL

9–10

hEGF (100 ng/μL)

100 μg/mL

100 ng/mL

7 μL

9–10

Activin A

1 mg/mL

10 ng/mL

0.7 μL

9–10

Y-27632

10 mM

10 μM

7 μL

9–10

Nicotinamide

1M

10 mM

70 μL

Stage 4 (Aggrewells): 11–12

Differentiation Mix 2

n/a

n/a

5 mL

11–12

S4 Supplement SANT1 (0.625 mM) Retinoic acid (0.25 mM) LDN-193189 (0.5 mM) TPB (2.5 mM)

2500×

1×

2 μL

11–12

Vitamin C

250 mM

0.25 mM

5 μL

11–12

FGF-7

2 μg/mL

2 ng/mL

5 μL

11–12

hEGF (100 ng/μL)

100 μg/mL

100 ng/mL

5 μL

11–12

Activin A

1 mg/mL

10 ng/mL

0.5 μL

11–12

Y-27632

10 mM

10 μM

5 μL

11–12

Nicotinamide

1M

10 mM

50 μL

13–16

Differentiation Mix 3

n/a

n/a

5 mL

13–16

S5 Supplement SANT1 (0.375 mM) Retinoic acid (75 μM) LDN-193189 (0.15 mM) GC1 (1.5 mM) GSiXX (0.15 mM) ALK5inhII (15 mM)

1500×

1×

3.3 μL

Stage 5:

(continued)

Gene Silencing in Differentiating Stem Cells

31

Table 5 (continued) Day

Reagent

Stock concentration

Final concentration

Amount

13–16

Betacellulin

100 μg/mL

20 ng/mL

1 μL

13–16

Y-27632

10 mM

10 μM

5 μL

17–23

Differentiation Mix 3

n/a

n/a

4 mL

17–23

S6 Supplement LDN-193189 (0.2 mM) GC1 (2 mM) GSiXX (0.2 mM) ALK5inhII (20 mM)

2000×

1×

2 μL

Stage 6:

Stage 7: 24–44

Differentiation Mix 4

4 mL

24–44

N-acetylcysteine

100 mM

1 mM

40 μL

24–44

T3

10 mM

10 nM

0.004 μL

24–44

ZM-447439

5 mM

0.5 μM

0.4 μL

GapmeR

GapmeR

GapmeR

D1-D3

hiPSC

Stage 2 (S2)

Stage 1 (S1)

Stage 0 (S0)

D4-D5

definitive endoderm

D6-D7

primitive gut tube

D9-D10

posterior foregut

Stage 5 (S5)

Stage 4 (S4)

Stage 3 (S3)

D11-D13

pancreatic endoderm

Pancreatic endocrine precursor cell

Stage 7 (S7)

Stage 6 (S6) D14-D20

D21-D27

Immature endocrine cell

Beta-like cell

Fig. 1 Scheme depicting a method in which specific gene is targeted for silencing by using GapmeRs during the differentiation protocol in stage 3 (posterior foregut stage) 3.1 Differentiation Toward Pancreatic Endocrine Cell Fate

The differentiation protocol used is based on Barsby et al. with the following modifications. 1. One day prior to the start of the differentiation, collect 1.2 × 107 stem cells in 7 mL mTeSR with 7 μL of 10 mM ROCK Inhibitor and plate on a Geltrex-coated 10 cm dish. In order to begin the differentiation process, the culture medium must be replaced daily with the corresponding basal medium from Tables 1 and 2 complemented as indicated in Table 5. 2. Once day 11 is reached, the Aggrewell plates must be prepared as follows:

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Lucas Unger et al.

(a) Add 2 mL of anti-adherence solution to each well and centrifuge at 1300 g for 6 min. (b) Remove the anti-adherence solution and wash with 2 mL DMEM per well. 3. The differentiating cells must then be transferred from the 10 cm dish into the Aggrewells using the following steps: (a) Wash the differentiating cells with 5 mL 1× PBS. (b) Add 5 mL TrypLE and incubate at 37 °C until cells start detaching. (c) Cease the trypsinization by adding 5 mL DMEM. (d) Determine the total amount of cells utilizing a cell counter and centrifuge the cells at 200 g for 3 min. (e) Resuspend the cell pellet in pre-warmed day 11 differentiation media at a cell density of 1.2 × 106/mL. (f) Add 5 mL of the cell suspension per well of the 6-aggrewell plate. 4. On day 15, the islet aggregates must be transferred from the Aggrewells to a rotating suspension culture in an ultra-low attachment (ULA) 6-well plate: (a) Transfer 2 mL of media of each well of the Aggrewell plate into the ULA plates. (b) Using a 10 mL serological pipette, gently transfer the aggregates into the ULA plates. (c) Swirl the plate so that the aggregates accumulate in the center and replace 4 mL of the media with the day 15 differentiation media. (d) Place the ULA plate on a platform rotator (95 RPM) in a cell culture incubator (5% CO2, 37 °C). 5. From this stage onward, the differentiation media only needs to be changed every 2–3 days. Swirl the plate so that the aggregates accumulate in the center and replace 4 mL of the media with the corresponding differentiation media. 3.2 Gene Silencing During Differentiation in 2D Conditions

1. The design parameters focus on achieving high potency by ensuring optimal target sequence accessibility through local secondary structure prediction, evaluating antisense off-target effects using ENSEMBL sequence alignments, and optimizing various factors such as length, Tm, gap size, self-complementarity, and LNA positions in the oligonucleotide design. We highly recommend utilizing the Antisense LNA GapmeR Custom Builder provided by Qiagen for GapmeR design purposes.

Gene Silencing in Differentiating Stem Cells

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2. Reconstitute the GapmeRs to a desired concentration of 50 nM in nuclease-free TE-Buffer (10 mM Tris–HCl, pH 7.5, 0.1 mM EDTA). See Note 1. 3. Progress with the differentiation protocol (see Subheading 3.1) in a 6-well plate following standard procedures until the desired stage of gene silencing is reached. See Note 2. 4. In two sterile microcentrifuge tube, combine 150 μL of OptiMEM with the 9 μL of RNAiMAX transfection reagent according to the manufacturer’s instructions. Mix gently and incubate the mixture at room temperature for 5 min. 5. Transfer 150 μL Opti-Mem per tube into two sterile microcentrifuge tubes, add 9 μL (450 pmol) of mTOR (in tube 1) or 9 μL control GapmeR working solution (in tube 2) and mix gently. See Note 3. 6. Mix each of the diluted GapmeR with the prepared working solution of RNAiMAX transfection reagent. Gently pipette up and down to ensure thorough mixing. Incubate the mixture for 10 min at room temperature to allow the formation of GapmerR-RNAiMAX complexes. 7. Remove the cell culture medium from the cells and add differentiation mix for the corresponding day. Add the GapmerRRNAiMAX complexes dropwise into the media. Ensure that the mixture is evenly distributed by gently moving the plate back and forth. 8. Place the cells in a 37 °C incubator with 5% CO2 for 24–48 h, to allow for GapmeRs uptake. 9. After the incubation period, carefully remove the transfection mixture from the wells without disturbing the cells. Replace it with corresponding differentiation mix according to the followed differentiation protocol. See Note 4. 3.3 Evaluation of Cell Viability

For the cell count and viability, we are using Via1-Cassette with Reagent A100 and B for NucleoCounter NC-200 from ChemoMetec, Denmark. The procedure presented below follows the recommendations of the supplier. 1. Collect 100 μL of cell suspension in one sterile 1.5 mL Eppendorf tube (1) and 200 μL in second Eppendorf tube (2). 2. In tube 1, add 100 μL solution A100, mix, add 100 μL solution B, mix, then load into one Via-1 cassette, and measure on NucleoCounter NC-200. 3. Load a second Via-1 cassette from tube 2 and measure viability and cell number. 4. Analyze the cell viability per condition. For mTOR GapmeRs, no obvious effect on cell viability was observed (Fig. 2a).

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Fig. 2 Quality controls for targeted silencing by specific mTOR GapmeRs. (a) Viability of transfected and non-transfected cells. (b) Knockdown of human mTOR in differentiating posterior foregut cells assessed by qPCR at 24 h and 72 h post-transfection. (c) Comparison of pancreatic differentiation markers, PDX1 and FOXA2, labeled at 24 h post-transfection with mTOR GapmeRs versus non-transfected control cells

3.4 Evaluation of Gene Silencing Efficiency (qPCR)

To evaluate the GapmeRs efficiency, we generated RNA extracts and used specific mTOR primers for measurements by qPCR. 1. Harvest the differentiating transfected cells using 5 mL TrypLE to generate a suspension of single cells and cleanse the trypsinization reagent by washing with 10 mL DMEM after cells start detaching (6–8 min). Transfer the cell suspension into a 15 mL falcon tube. 2. Count cells in cell suspension to aliquot 0.5 × 106 cells per tube and centrifuge at 200 g for 3 min. 3. Remove supernatant and wash cell pellet with 5 mL 1× DPBS followed by centrifugation at 200 g for 3 min.

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4. Remove supernatant and flash freeze in liquid nitrogen for long-term storage at -80 °C or continue to step 5. See Note 5. 5. Immediately after removing the frozen cell pellet form the liquid nitrogen, add 350 μL of 0.1% β-mercaptoethanol RLT buffer. 6. Extract RNA following the Qiagen RNeasy mini protocol. 7. With the obtained RNA, follow the QuantiNova SYBR GreenRT-PCR Kit to verify the gene silencing efficiency via qPCR. Evaluate the obtained qPCR data by calculating the relative expression of mTOR and the percentage of RNA decrease (Fig. 2b). 3.5 Evaluation of GapmeRs Impact on Differentiation Efficiency (Immunofluorescence)

To assess the impact of GapmeRs treatment on differentiation process, we investigated selected markers of pancreatic differentiation by immunofluorescence staining. Several time points can be selected for evaluating the differentiation efficiency. Here, we decided for 24 h after the transfection (Fig. 2c). 1. 24 h post transfection, transfer the coverslips by using tweezers into a clean 12-well plate filled with PBS. See Note 6. 2. Fix cell for 20 min at room temperature with 1 mL of 2% PFA. 3. Wash cells with 1 mL PBS three times for 5 min. 4. Permeabilize the fixed cells by removing the PBS and adding 500 μL 2% Triton ×100 in PBS for 20 min at room temperature and remove afterward. 5. Add 2% BSA in PBS for 30 min at room temperature to block against unspecific binding. 6. Prepare the primary antibody solution by calculating per slide a mix of 500 μL 2% BSA in PBS with 2.5 μL of PDX1 and FOXA2 antibody. 7. Replace the 2% BSA solution with primary antibody mix solution and incubate over night at 4 °C. 8. The following day, remove the antibody mix and wash three times with 500 μL 1× PBS for 5 min. 9. Prepare the secondary antibody solution by calculating per slide a mix of 500 μL 2% BSA in PBS with 1 μL of DAPI, 1 μL of corresponding secondary antibodies, and 5 μL of Phalloidin. 10. Replace the 1× PBS, wash with secondary antibody mix solution, and incubate for 1 h at room temperature in dark. 11. Wash coverslips three times with 500 μL 1× PBS for 5 min and mount cells onto microscope slides using 1 drop of mounting media. 12. Image the slides and document observations for subsequent analysis.

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Notes 1. Stock solutions of Antisense LNA GapmeRs should not be less than 10 μM. 2. The intracellular concentration of GapmeRs within the transfected cells can be influenced by cell confluency. A lower cell confluency can have a higher GapmeRs activity using the same concentration and transfection conditions. Similar confluency is needed to decrease inter-experimental variability. 3. Both the specific GapmeR sequences and the concentration should be optimized beforehand. From our experience, a threefold increase in the recommended amount of siRNA in the RNAiMAX protocol yielded optimal results. 4. In case the gene silencing should last longer than 72 h, we recommend a new transfection with GapmeRs every 72 h. 5. The cell pellet in 0.1% β-mercaptoethanol RLT buffer can be stored for long term at -80 °C. 6. In order to perform immunofluorescence staining at different differentiation stages, glass coverslips should be placed into the plate at the beginning of the differentiation protocol, just before coating the 6-well plates with Geltrex.

Acknowledgments The authors thank Max Pietschmann for expert advice and support on establishing the silencing method. The authors also acknowledge research funding received from the Research Council of Norway (NFR 251041 and 314397), Novo Nordic Foundation (NNF15OC0015054 and NNF21OC0067325), and the Norwegian Diabetesforbundet forskningfond. References 1. Pagliuca FW, Millman JR, Gurtler M, Segel M, Van Dervort A, Ryu JH, Peterson QP, Greiner D, Melton DA (2014) Generation of functional human pancreatic beta cells in vitro. Cell 159(2):428–439. https://doi.org/10. 1016/j.cell.2014.09.040 2. Russ HA, Parent AV, Ringler JJ, Hennings TG, Nair GG, Shveygert M, Guo T, Puri S, Haataja L, Cirulli V, Blelloch R, Szot GL, Arvan P, Hebrok M (2015) Controlled induction of human pancreatic progenitors produces functional beta-like cells in vitro. EMBO J 34(13):1759–1772. https://doi.org/10. 15252/embj.201591058

3. Balboa D, Otonkoski T (2015) Human pluripotent stem cell based islet models for diabetes research. Best Pract Res Clin Endocrinol Metab 29(6):899–909. https://doi.org/10. 1016/j.beem.2015.10.012 4. Rezania A, Bruin JE, Arora P, Rubin A, Batushansky I, Asadi A, O’Dwyer S, Quiskamp N, Mojibian M, Albrecht T, Yang YH, Johnson JD, Kieffer TJ (2014) Reversal of diabetes with insulin-producing cells derived in vitro from human pluripotent stem cells. Nat Biotechnol 32(11):1121–1133. https:// doi.org/10.1038/nbt.3033

Gene Silencing in Differentiating Stem Cells 5. Balboa D, Iworima DG, Kieffer TJ (2021) Human pluripotent stem cells to model islet defects in diabetes. Front Endocrinol (Lausanne) 12:642152. https://doi.org/10. 3389/fendo.2021.642152 6. Beydag-Tasoz BS, Yennek S, Grapin-Botton A (2023) Towards a better understanding of diabetes mellitus using organoid models. Nat Rev Endocrinol 19(4):232–248. https://doi.org/ 10.1038/s41574-022-00797-x 7. Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126(4):663–676. https://doi.org/10. 1016/j.cell.2006.07.024 8. Barsby T, Ibrahim H, Lithovius V, Montaser H, Balboa D, Vahakangas E, Chandra V, Saarimaki-Vire J, Otonkoski T (2022) Differentiating functional human isletlike aggregates from pluripotent stem cells. STAR Protoc 3(4):101711. https://doi.org/ 10.1016/j.xpro.2022.101711 9. Velazco-Cruz L, Song J, Maxwell KG, Goedegebuure MM, Augsornworawat P, Hogrebe NJ, Millman JR (2019) Acquisition of dynamic function in human stem cell-derived beta cells. Stem Cell Reports 12(2):351–365. https:// doi.org/10.1016/j.stemcr.2018.12.012 10. Balboa D, Barsby T, Lithovius V, SaarimakiVire J, Omar-Hmeadi M, Dyachok O, Montaser H, Lund PE, Yang M, Ibrahim H, Naatanen A, Chandra V, Vihinen H, Jokitalo E, Kvist J, Ustinov J, Nieminen AI, Kuuluvainen E, Hietakangas V, Katajisto P, Lau J, Carlsson PO, Barg S, Tengholm A, Otonkoski T (2022) Functional, metabolic and transcriptional maturation of human pancreatic islets derived from stem cells. Nat Biotechnol 40(7):1042–1055. https://doi.org/ 10.1038/s41587-022-01219-z 11. Ghila L, Legoy TA, Mathisen AF, Abadpour S, Paulo JA, Scholz H, Raeder H, Chera S (2021) Chronically elevated exogenous glucose elicits antipodal effects on the proteome signature of differentiating human iPSC-derived pancreatic progenitors. Int J Mol Sci 22(7). https://doi. org/10.3390/ijms22073698 12. Liu C, Oikonomopoulos A, Sayed N, Wu JC (2018) Modeling human diseases with induced pluripotent stem cells: from 2D to 3D and beyond. Development 145(5). https://doi. org/10.1242/dev.156166 13. Vegas AJ, Veiseh O, Gurtler M, Millman JR, Pagliuca FW, Bader AR, Doloff JC, Li J, Chen M, Olejnik K, Tam HH, Jhunjhunwala S, Langan E, Aresta-Dasilva S, Gandham S, McGarrigle JJ, Bochenek MA, Hollister-Lock J, Oberholzer J, Greiner DL,

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Weir GC, Melton DA, Langer R, Anderson DG (2016) Long-term glycemic control using polymer-encapsulated human stem cell-derived beta cells in immune-competent mice. Nat Med 22:306. https://doi.org/10.1038/nm. 4030 14. Legoy TA, Vethe H, Abadpour S, Strand BL, Scholz H, Paulo JA, Raeder H, Ghila L, Chera S (2020) Encapsulation boosts islet-cell signature in differentiating human induced pluripotent stem cells via integrin signalling. Sci Rep 10(1):414. https://doi.org/10.1038/ s41598-019-57305-x 15. Ghila L, Legoy TA, Chera S (2021) A method for encapsulation and transplantation into diabetic mice of human induced pluripotent stem cells (hiPSC)-derived pancreatic progenitors. Methods Mol Biol. https://doi.org/10. 1007/7651_2021_356 16. Carrasco M, Wang C, Soviknes AM, Bjorlykke Y, Abadpour S, Paulo JA, Tjora E, Njolstad P, Ghabayen J, Nermoen I, Lyssenko V, Chera S, Ghila LM, Vaudel M, Scholz H, Raeder H (2022) Spatial environment affects HNF4A mutation-specific proteome signatures and cellular morphology in hiPSC-derived beta-like cells. Diabetes 71(4): 8 6 2 – 8 6 9 . h t t p s : // d o i . o r g / 1 0 . 2 3 3 7 / db20-1279 17. Petersen MBK, Goncalves CAC, Kim YH, Grapin-Botton A (2018) Recapitulating and deciphering human pancreas development from human pluripotent stem cells in a dish. Curr Top Dev Biol 129:143–190. https://doi. org/10.1016/bs.ctdb.2018.02.009 18. Veres A, Faust AL, Bushnell HL, Engquist EN, Kenty JH, Harb G, Poh YC, Sintov E, Gurtler M, Pagliuca FW, Peterson QP, Melton DA (2019) Charting cellular identity during human in vitro beta-cell differentiation. Nature 569(7756):368–373. https://doi.org/10. 1038/s41586-019-1168-5 19. Petersen MBK, Azad A, Ingvorsen C, Hess K, Hansson M, Grapin-Botton A, Honore C (2017) Single-cell gene expression analysis of a human ESC model of pancreatic endocrine development reveals different paths to beta-cell differentiation. Stem Cell Reports 9(4): 1246–1261. https://doi.org/10.1016/j. stemcr.2017.08.009 20. Sharon N, Vanderhooft J, Straubhaar J, Mueller J, Chawla R, Zhou Q, Engquist EN, Trapnell C, Gifford DK, Melton DA (2019) Wnt signaling separates the progenitor and endocrine compartments during pancreas development. Cell Rep 27(8):2281–2291 e2285. https://doi.org/10.1016/j.celrep. 2019.04.083

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21. Legoy TA, Ghila L, Vethe H, Abadpour S, Mathisen AF, Paulo JA, Scholz H, Raeder H, Chera S (2020) In vivo hyperglycaemia exposure elicits distinct period-dependent effects on human pancreatic progenitor differentiation, conveyed by oxidative stress. Acta Physiol 228(4):e13433. https://doi.org/10.1111/ apha.13433 22. Legoy TA, Mathisen AF, Salim Z, Vethe H, Bjorlykke Y, Abadpour S, Paulo JA, Scholz H, Raeder H, Ghila L, Chera S (2020) In vivo environment swiftly restricts human pancreatic progenitors toward mono-hormonal identity via a HNF1A/HNF4A mechanism. Front Cell Dev Biol 8:109. https://doi.org/10. 3389/fcell.2020.00109 23. Ghila L, Bjorlykke Y, Legoy TA, Vethe H, Furuyama K, Chera S, Raeder H (2020) Bioinformatic analyses of miRNA-mRNA signature during hiPSC differentiation towards insulinproducing cells upon HNF4alpha mutation. Biomedicine 8(7). https://doi.org/10.3390/ biomedicines8070179 24. Vethe H, Ghila L, Berle M, Hoareau L, Haaland OA, Scholz H, Paulo JA, Chera S, Raeder H (2019) The effect of Wnt pathway modulators on human iPSC-derived pancreatic Beta cell maturation. Front Endocrinol (Lausanne) 10:293. https://doi.org/10.3389/fendo. 2019.00293 25. Fontes A, Lakshmipathy U (2013) Advances in genetic modification of pluripotent stem cells. Biotechnol Adv 31(7):994–1001. https://doi. org/10.1016/j.biotechadv.2013.07.003 26. Yamoah MA, Thai PN, Zhang XD (2021) Transgene delivery to human induced pluripotent stem cells using nanoparticles. Pharmaceuticals (Basel) 14(4). https://doi.org/10. 3390/ph14040334 27. Stein CA, Hansen JB, Lai J, Wu S, Voskresenskiy A, Hog A, Worm J,

Hedtjarn M, Souleimanian N, Miller P, Soifer HS, Castanotto D, Benimetskaya L, Orum H, Koch T (2010) Efficient gene silencing by delivery of locked nucleic acid antisense oligonucleotides, unassisted by transfection reagents. Nucleic Acids Res 38(1):e3. https:// doi.org/10.1093/nar/gkp841 28. Shen X, Beasley S, Putman JN, Li Y, Prakash TP, Rigo F, Napierala M, Corey DR (2019) Efficient electroporation of neuronal cells using synthetic oligonucleotides: identifying duplex RNA and antisense oligonucleotide activators of human frataxin expression. RNA 25(9):1118–1129. https://doi.org/10.1261/ rna.071290.119 29. Deleavey GF, Damha MJ (2012) Designing chemically modified oligonucleotides for targeted gene silencing. Chem Biol 19(8): 937–954. https://doi.org/10.1016/j. chembiol.2012.07.011 30. Bennett CF, Swayze EE (2010) RNA targeting therapeutics: molecular mechanisms of antisense oligonucleotides as a therapeutic platform. Annu Rev Pharmacol Toxicol 50:259– 293. https://doi.org/10.1146/annurev. pharmtox.010909.105654 31. Liang XH, Sun H, Nichols JG, Crooke ST (2017) RNase H1-dependent antisense oligonucleotides are robustly active in directing RNA cleavage in both the cytoplasm and the nucleus. Mol Ther 25(9):2075–2092. https:// doi.org/10.1016/j.ymthe.2017.06.002 32. Matsuzono K, Imamura K, Murakami N, Tsukita K, Yamamoto T, Izumi Y, Kaji R, Ohta Y, Yamashita T, Abe K, Inoue H (2017) Antisense oligonucleotides reduce RNA foci in spinocerebellar ataxia 36 patient iPSCs. Mol Ther Nucleic Acids 8:211–219. https://doi. org/10.1016/j.omtn.2017.06.017

Methods in Molecular Biology (2023) 2736: 39–51 DOI 10.1007/7651_2023_480 © Springer Science+Business Media, LLC 2023 Published online: 05 May 2023

Retinoic Acid-Mediated Differentiation of Mouse Embryonic Stem Cells to Neuronal Cells Sangeeta Dutta, Debosree Pal, and M. R. S. Rao Abstract The capability of pluripotent embryonic stem cells (ESCs) to proliferate and differentiate into specific lineages makes them an important avenue of research in the field of cell therapy as well as a useful model to study patterns of differentiation and gene expression, recapitulating many events that occur during the very early stages of development of the mammalian embryo. With striking similarities that exist between inherently programmed embryonic development of the nervous system in vivo and the differentiation of ESCs in vitro, they have already been used to treat locomotive and cognitive deficits caused by brain injury in rodents. A suitable differentiation model thus empowers us with all these opportunities. In this chapter, we describe a neural differentiation model from mouse embryonic stem cells using retinoic acid as the inducer. This method is among the most commonly used one to acquire a homogeneous population of neuronal progenitor cells or mature neurons as desired. The method is scalable, efficient, and results in production of ~70% neural progenitor cells within 4–6 days. Key words E14TG2a cell line, Embryoid bodies, Mature neurons, Mouse embryonic stem cells (mESCs), Neuronal differentiation, Neuronal progenitor cells (NPCs), Retinoic acid (RA), Stem cell culture

1

Introduction An appropriate differentiation culture system can provide unlimited quantities of homogeneous material in vitro, which is vital for any study related to understanding the molecular, biochemical, and physiological process of differentiation. A good model system allows us to alter the cellular physiology, specifically and rapidly, giving us the chance to study the impact on cell fate specification process under predefined conditions. One of the first lineages to be developed from embryonic stem cells (ESCs) were neuronal cells [1, 2]. Several subsequent studies effectively differentiated ESCs into different cellular subtypes like neurons and glial cells. Over the last three decades, neural cells differentiated from pluripotent stem cells (PSCs), including ESCs and induced pluripotent stem cells (iPSCs) have become a powerful tool for basic understanding, disease modeling, drug screening and regenerative medicine.

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An applicable in vitro model for development is hypothesized as the one in which neural differentiation can be induced that reflects the earliest fate of neuronal development as it occurs in vivo in ectoderm. With recent advances, a similarity in pattern of neuronal differentiation from ESCs in vitro and neural development in vivo has been witnessed. This property has provided promising outcome in stem cell research, especially in the field of biomedical application and regenerative medicine. There are several inducers and pathways through which neural differentiation can be achieved like retinoic acid, Wnt/β-catenin, transforming growth factor/bone morphogenetic protein, Notch, fibroblast growth factor, Hedgehog, c-Jun N-terminal kinase/mitogen-activated protein kinase, to mention few [3]. Retinoic acid (RA)-induced neural differentiation model system is one of the most commonly used systems, which was first described by Bibel et al. in 2004 [4, 5]. When ESCs are cultured as cellular aggregates and treated with RA, they commit to a neural fate to form neuronal progenitors. These progenitors have the capacity to give rise to glutamatergic neurons under specific conditioned media, forming a dense axon network within 7–9 days [4–6]. Retinoic acid (RA) is an active derivative of vitamin A which plays a key role in cell growth and differentiation by activating nuclear receptors, RARs (α, β, and γ), which are ligand-dependent regulators of transcription [7–9]. We now have much, but not full, understanding of how RA drives the differentiation process; however, it is certainly clear from several reports that RA has genomic [10, 11] as well as non-genomic effects [12, 13]. RARγ subtype (particularly RARγ2 isoform) was shown to be essential for RA-induced commitment of mouse ESCs into the neuronal lineage. RARγ2 is rapidly phosphorylated after RA addition and appears to activate a specific subset of key genes involved in RA metabolism (Cyp26a1, Cyp26b1, Cyp26c1, and Dhrs3), patterning genes exemplified by the Hox genes (Hoxa1, Hoxa3, Hoxa5, Hoxb1, Hoxb4), other genes encoding homeobox proteins (Meis2, Cdx1, Gbx2, Dlx3 and HNF1β) and genes with a wide variety of functions such as Lefty1, Arg1 and Stra8. Mechanism of activation of these genes was found to be epigenetically regulated via binding specifically to retinoic acid response elements (RAREs) with direct repeats with a 5 and 7 base pair spacer [14, 15]. Non-genomic effect of RA has been demonstrated in several studies where it rapidly and transiently activates several kinase cascades exemplified by the MAPK pathways. In ESCs, RA activates p42/p44MAPKs (also called Erks) which translocate to nucleus where they phosphorylate several targets like MSK1 (mitogen- and stress-activated protein kinase). Downstream, MSK1 phosphorylates histones and several nuclear proteins involved in the transcription of the RA-target genes, including RARs themselves [14–16].

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Fig. 1 Schematic representation of the timeline showing differentiation of mouse ES cells, first into NPCs and then to neurons using RA as neuronal differentiation inducer and by using precise conditioned media at specific points of neuronal differentiation

In this chapter, we describe the cell culture technique to obtain neuronal progenitor cells (NPCs) and mature neurons from mouse ESCs with induction of RA in approximately 2 weeks’ time (Fig. 1). We follow the original method as described by Bible et al. with few modifications [17, 18]. For the present methodology, we used rapidly growing ES-E14TG2a feeder–independent mouse ESCs line derived from the blastocyst of the embryo of a 129/Ola strain mouse. Initial ESCs cultures are allowed to form embryoid bodies and are guided to differentiate into NPCs under the influence of RA and specific conditioned media (Fig. 1).

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Materials Cell culture grade reagents should be used to prepare all the chemicals in sterilized milli Q/ultra-pure water (prepared by purifying deionized water to attain a sensitivity of 18 MΩ-cm at 25 °C). Prepared reagents should be autoclaved/filter (using 0.2-μm syringe filter) as mentioned. Store all the prepared reagents at room temperature unless mentioned otherwise.

2.1

Chemicals

1. DMEM (Dulbecco’s modified Eagle medium) high glucose (Sigma, Catalog No. D1152), store at 4 °C. 2. Fetal bovine serum (FBS) (Gibco, Catalog No. 16000-044), aliquot in 50 mL vials and store at -20 °C (see Note 1). 3. 1× non-essential amino acids (NEAA) (Sigma, Catalog No. M7145). NEAA is supplied as 100×. Aliquot 10 mL of 100× NEAA in 15 mL vials and store at -20 °C (see Note 2).

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4. 1× Penicillin-streptomycin solution (Sigma, Catalog No. P4333). The concentration of solution is supplied as 100×. Aliquot 1 mL of 100× solution in 1.5 mL vials and store at -20 °C. 5. 1× Leukemia Inhibitory Factor (LIF) (ESGRO® Recombinant Mouse LIF Protein, Merck Millipore, Catalog No. ESG1107). Stock supplied is 1000× and does not need any dilution. 6. 0.1 mM β-mercaptoethanol (Sigma, Catalog No. 63689). Filter and sterilize the solution and store at 4 °C (see Note 3). 7. 0.2% Gelatin (HiMedia, Catalog No. TC041). Dissolve 0.2 g gelatin in 100 mL water, autoclave, and store at 4 °C (see Note 4). 8. 0.05% of Trypsin (Sigma, Catalog No. T4799). Store at 4 °C (see Note 5). 9. 5 μM Retinoic acid (Sigma, Catalog No. R2625). Prepare 10 mM stock by dissolving 3 mg in 1 mL of DMSO, sterilize by using syringe filter inside a laminar hood, aliquot 100 μL, and store at -20 °C (see Note 6). 10. 1× PBS (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.47 mM KH2PO4, pH of 7.4) (see Note 7). 11. Laminin (5 μg/mL) (Roche, Catalog number: 11243217001). Comes as a solution of 0.5 mg/mL, aliquot it into 500 μL and store at -20 °C. Dilute with sterile water to make a concentration of 5 μg/mL (for 10 mL, take 100 μL of 0.5 mg/mL laminin and add 9.9 mL PBS, invert mix). 12. Poly-L-ornithine (100 μg/mL or 0.01% w/v) (Sigma, Catalog No. A-004-M). Make 10 mL aliquot and store at -20 °C. 13. DMEM-F12 (ThermoFisher, Catalog No. 11320033). Store at 4 °C. 14. N2 Supplement (ThermoFisher, Catalog No. 17502048). Stock is supplied as 100×; aliquot into 500 μL and store at -20 °C. 15. B27 Supplement (ThermoFisher, Catalog No. 17504044). Stock is supplied as 50×; aliquot into 1 mL and store at -20 °C. 16. DPBS (ThermoFisher, Catalog No. 14040133) and store at 4 °C. 2.2

Cell Line

2.3

Miscellaneous

E14TG2a feeder-independent mouse ES cell line (ATCC, CRL-1821™). 1. Cell/tissue culture grade plates (90 mm, 60 mm, 35 mm) (see Note 8). 2. Bacterial grade petri plates (90 mm, 30 mm) (see Note 8).

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3. 40 μm syringe filters (Falcon Catalog No. 352340). 4. 0.2 μm syringe filters (mdi, pre-sterilized PES membrane syringe filter).

3

Methods

3.1 Initiation of ESC Culture

1. Prepare the ESC media. (DMEM high glucose supplemented with 15% FBS, 1× non-essential amino acids, 0.1 mM β-mercaptoethanol, and 1× penicillin-streptomycin along with 1× LIF) (see Notes 9 and 10). 2. Coat the cell culture grade plate with 0.2% gelatin. Warm the gelatin to room temperature (RT). Depending on the size of plate to be used, 90 mm or 35 mm, pour 5 mL or 2 mL of 0.2% gelatin into the plates, respectively. Keep at 37 °C for at least 10 min. Discard the excess by tilting the plate to one side and pipetting out excess of gelatin from periphery. Immediately pour a layer of ES media to the coated plate to avoid drying and keep them at 37 °C. 3. Take out stocks of ESCs stored in liquid nitrogen. Thaw the cells by putting the vial in a beaker containing water at 37 °C for 3–5 min. 4. Once thawed, resuspend the cells in 5 mL of freshly prepared ESC media. 5. Spin down at 91–101 g for 4 min at RT to collect the cells. Discard the media. 6. Resuspend the cell pellet in 1 mL of fresh media. Pipette well to evenly distribute the cells into a single cell suspension. 7. Pour the cell suspension in gelatin-coated plates and make up the volume (see Note 11). Swirl the plates gently 8–10 times for equal distribution of cells throughout the plate. 8. Keep the plates in incubator with 37 °C temperature and 5% CO2 and allow the cells to grow. 9. After 24 h of incubation, cells should be attached to the plate. Discard the media to get rid of dead and floating cells. Pour fresh ESC media and keep the plates back in incubators according to the above-mentioned conditions. 10. Keep changing media every alternate day till the cells become 70–80% confluent (Fig. 2a).

3.2 Subculturing ESCs

1. Subculturing should be done once the cells reach 70–80% confluency (see Note 12). 2. For subculture, discard the media. Wash the cells by pouring 2–3 mL of PBS (at RT) to the dish. Swirl around few times and

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Fig. 2 Various stages of RA-induced neuronal differentiation as observed under microscope. (a) Fully confluent ESCs. (b–d) Floating globular embryoid bodies (EBs) at days 2, 4, and 8 after initiation. There is a visible increase in size of the EBs with time. (e, f) Neurons as observed after 2 h and 24 h after initiation with a section zoomed to show the cells. Scale bar = 100 μm

discard the PBS by tilting the plate to one side and pipetting out from the periphery (see Note 13). 3. Add 1 mL of freshly prepared 0.05% trypsin (at RT) to the plate. Keep at 37 °C for 30 s to 1 min (see Note 14). 4. Add 1–2 mL of ES media to stop or neutralize the trypsinization process. 5. Collect the cells by pipetting around the dish few times and transferring the cell suspension into a falcon. Spin down the cells at 91–101 g for 4 min at RT. 6. Discard the supernatant and resuspend the cells in 1 mL of fresh media. Pipette thoroughly to break all clumps and till the solution comprises mostly single cells (see Note 15). 7. Passage ESCs at a ratio of 1:5. Take 200 μL of cell suspension and add it to a new gelatin-coated 90 mm plate containing 7 mL of ESC media. Keep the plates in incubator with 37 °C temperature and 5% CO2 and allow the cells to grow to confluency (see Notes 16 and 17). 3.3 Initiation of Embryonic Bodies (EB) Culture

1. Prepare the EB media (DMEM high glucose supplemented with 10% FBS, 0.1 mM β-mercaptoethanol, and 1× penicillinstreptomycin) (see Note 18). 2. Trypsinize the ESCs. Collect the cell pellet and resuspend them in 1 mL of EB media as described above in Subheading 3.2, steps 1 to 6.

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3. Count the number of cells using a hemocytometer. 4. Take bacterial grade plates for EB culture. For a 90 mm plate, resuspend 4 × 106 cells in 7 mL of EB media, and for a 35 mm plate, resuspend 2.5 × 105 cells in 2 mL EB media. 5. Change the media every alternate day using gravity sedimentation. Collect all the media (containing floating globular EBs, Fig. 2b) from the plate into a 15 mL falcon. Keep the falcon tube upright on a stand and allow the EBs to settle down for 5–10 min. Carefully discard the media without disturbing the sedimented EBs. Resuspend the EBs in fresh EB media, invert mix once, and pour back into the culture plate (see Note 19). 3.4 Differentiation of EBs into Neuronal Progenitor Cells (NPCs).

1. On the 4th day after initiation of EBs, change the media as described in Subheading 3.3, step 5. (Fig. 2c) 2. Add retinoic acid (RA) to the culture to differentiate EBs into NPCs (see Note 20). 3. Continue the EB culture for 4 more days under the influence of RA. Change media every alternate day along with addition of RA as described in step 2 and Subheading 3.3, step 5 (Fig. 2d).

3.5

Harvesting EBs

1. Eight days after initiating of EBs (4 days without RA and 4 days with RA, or 4+/-RA, collect the cells by gravity sedimentation. 2. Discard the media and wash once with 1× PBS. One can score for neuronal marker using various techniques in EBs at this stage (Fig. 3a, b). 3. To collect NPCs, trypsinize the RA-treated EBs with 0.5 mL of 0.05% freshly prepared trypsin (see Note 5). 4. Incubate at 37 °C for 3–5 min. During incubation, shake the falcon tube once or twice. EBs should appear to disintegrate. 5. Add EB medium to inactivate the trypsin. Dissociate EBs by pipetting, they should dissociate within 10 times of pipetting up and down. 6. Pass the solution drop-by-drop through a 40 μm filter to remove any undissociated EBs. 7. Centrifuge at 140 g for 5 min to collect the NPCs. 8. Further processing of NPCs will depend upon the aim of the next experimental plan (see Note 21). For further differentiation of NPCs to neurons, follow the below-mentioned steps.

3.6 Differentiation of NPCs to Neurons

1. Coating the plates. (a) Add PLO solution (100 μg/mL) to plates (for example, 1 mL for each well of a 6-well dish). Evenly distribute and leave at 37 °C inside an incubator overnight.

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Fig. 3 Evaluation of various markers at different stages of RA-induced differentiation of ESCs to NPCs and neurons. (a) Immunofluorescence (IF) for NPC marker NESTIN and maturing neuron marker TUJ1 in cryosections of RA- versus vehicle-treated EBs. (b) qPCR analysis of various markers for neuronal lineage in RA-treated versus vehicle-treated EBs confirming our in vitro neuronal lineage differentiation system. PAX6 and NESTIN are well-known NPC markers, ASCL1 is a neuronal lineage marker, TUJ1 represents maturing neuron marker while GFAP is an astrocyte lineage marker. (c) qPCR analysis of various markers for NPCs and neurons at different hours of differentiation of NPCs to neurons. A reduction in NPC markers and increase in neuron marker can be observed with time. (d, e) IF for NPC marker NESTIN and maturing neuron marker TUJ1 in neurons at different hours of differentiation respectively. Error bars indicate s.d. from three independent experiments, with technical duplicates in each (n = 6). *p < 0.05, **p < 0.01, ***p < 0.001, student’s t-test; Scale bar = 10 μm

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(b) Next day, remove surplus PLO solution by tilting the plate and removing the excess with the help of a pipette. Wash plates thrice with DPBS (see Note 22). (c) Add laminin (5 μg/mL) to each well/plate, evenly distribute, and incubate at 37 °C for 2–3 h. (d) Retrieve laminin-coated dish immediately before plating NPCs. Aspirate excess laminin solution and immediately add medium/cells. Care must be taken so that laminincoated plates do not dry out. 2. Prepare neurobasal media (or N2) medium (DMEM-F12 along with 1× N2 supplement and 1× penicillin-streptomycin solution). Mix well without shaking. Store the media at 4 °C for up to 3 weeks. 3. Prepare N2B27 media (N2 media with 1× B27 supplement) (see Note 23). 4. Harvest the NPCs by following steps 2 to 7 under Subheading 3.5. Wash once with DPBS (see Note 24). 5. Resuspend the washed pellet of NPCs in N2 medium. Count the cell number using a hemocytometer. 6. Plate the cells at a density of 90,000 cells/cm2 onto PLO + laminin-coated plates in N2 medium. Change medium after 2 h and then again after 24 h with N2 medium. 7. Second day after plating, add N2B27 medium. The morphology of cells should change from spindle shape (visible when you will give media change at 2 h) to more neuronlike with projections from 16–20 h onward (Fig. 2e, f). 8. Neurons can be harvested as per the requirement of the experiment (for example at 12 h, 24 h, 2 days, and 4 days). Scoring for neuronal markers can be done at various time points (Fig. 3c–e).

4

Notes 1. Running vial can be stored at 4 °C for short duration of 1–2 weeks. 2. Running vial can be stored at 4 °C for short duration of 1–2 weeks. 3. To prepare 10 mL of the 100 mM solution, take a fresh 15 mL falcon, first pour the required amount of water, i.e., 9.930 mL, and then add 70 μL of 14.3 M solution of β-mercaptoethanol (commercially available concentration of β-mercaptoethanol is 14.3 M). Invert mix to mix the solution properly before filter sterilization.

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4. All the autoclaved reagents should be opened inside the laminar hood only to avoid contamination. 5. Should be prepared fresh every time by diluting supplied stock solution of 0.25% in autoclaved PBS (phosphate buffer saline). Make small volumes or as required to avoid wastage. To make 2 mL of working solution of 0.05% trypsin, take 1.6 mL of PBS and add 0.4 mL of 0.25% trypsin. 6. Further dilute the 10 mM stock to a second stock of 1 mM stock by adding sterilize water inside the laminar hood. Aliquot and store at -20 °C. Working vial of RA solution (1 mM) can be stored at 4 °C up to a week. Since RA and DMSO both are light sensitive chemicals, one should be cautious during its preparation and in its storage. 7. For 1 L of 1× PBS, start with 800 mL of distilled water and add 8 g of NaCl, 0.2 g of KCl, 0.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. Add distilled water to a total volume of 1 L. Sterilize by autoclaving and store at room temperature. Open only inside the hood. 8. We use cell culture grade plates from Corning and bacterial grade plates from Tarsons. 9. Before preparing the ESC media, all the components should be at room temperature (DMEM high glucose, FBS, penicillinstreptomycin solution, NAEE, β-mercaptoethanol) except LIF (which should be kept at 4 °C all the time). 10. ESC media should be prepared fresh every time before use. If required, it can be stored at 4 °C; however, it should be used within a week. Preparing small volume or as required works best as it avoids storage as well as wastage. We usually prepare a minimum volume of 50 mL. For that take a fresh 50 mL falcon, pour 41.5 mL of DEM (high glucose) followed by 7.5 mL of FBS, 500 μL of 100× penicillin-streptomycin solution, 500 μL of 100× NAEE, 50 μL of 100 mM β-mercaptoethanol, finally supplemented by 50 μL of 1000× LIF. Invert mix the prepared media and keep at 37 °C for minimum half an hour before use. 11. The plate size to be used for the revival of cells will depend upon the number of the cells in the stock. Use a final volume of 7 mL and 3 mL ES media for 90 mm or 35 mm dish, respectively. 12. We also found that the cells take more time to reach 70–80% confluency after revival of culture (usually 3–4 days). However, the growth rate is much faster once they are subcultured (usually 2 days). 13. Washing with PBS is important step. Without washing the remnant media containing even trace amount of FBS will interfere with trypsinization process.

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14. The amount of trypsin used is given for 90 mm plate. Use 200–500 μL for 35 mm plate. Usually, cells are let loose from the surface within 30 s but if you can see that cells are not full detached, keep for some more time, but not more than 1 min as it may cause cell damage and death. 15. To prevent cells from differentiating, they should be dissociated down to single cells during their passage. If they are not thoroughly dissociated, they are likely to form large clumps after passage, and the cells within these clumps will then differentiate or die. 16. Although the ratio during passage of cell that we generally use is 1:5, one should take care that the cell density is not too low. In that case, 1:3 or 1:4 ratio can be used as suited. Low cell density plating may cause them to form large clumps which contain differentiated cells prior to reaching confluence. 17. Also transferring cells from smaller to bigger plate, 35 mm to 90 mm (which have surface area of 9.5cm2 and 58 cm2), is equivalent to splitting the cells in 1:5. So, without any further splitting, the whole cell suspension can be transferred to a bigger plate and grown till confluent. 18. All the reagents should be at room temperature at the time of preparation of EB media. Media should be prepared fresh in small quantities to avoid storage and wastage. Excess media prepared should be stored at 4 °C for not more than a week. For 50 mL EB media, pour 44.5 mL of DMEM high glucose in a fresh 50 mL falcon, followed by 5 mL of FBS, 500 μL of 100× penicillin-streptomycin solution, and 50 μL of 100 mM β-mercaptoethanol. Invert well to mix and keep at 37 °C at least half an hour before use. 19. EBs are floating 3D globular structures. One can observe increase in the size of the globules as the culture progresses. For media change during the whole process of EB culture, gravity sedimentation (i.e., collecting EBs by letting them sediment under the force of gravity) should be used. Also, do not pipette or mix vigorously, otherwise the globular structure of EBs will break apart, not allowing cells to differentiate properly as intended. 20. For 90 mm plate containing 7 mL EB media, add 35 μL of 1 mM RA stock solution to get working concentration of 5 μM while 10 μL of 1 mM RA for 35 mm dish containing 2 mL of EB media. 21. NPC pellets obtained can be either flash frozen in liquid N2 and stored in -80 °C for future RNA isolation or/and protein extraction or could be used for these processes’ straight way. Alternatively, for experimental procedures like ChIP, NPCs can be cross-linked using 1% formaldehyde (Sigma, F8775) for

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10 min at room temperature followed by quenching with glycine at a final concentration of 0.125 M for 10 min at room temperature. Cross-linked cells are then centrifuged at 2000 g for 10 min at 4 °C and washed twice with 1× ice-cold PBS. Finally, the pellets can either be flash frozen in liquid N2 and stored at -80 °C or processed directly for chromatin extraction. 22. Ensure that the plates don’t dry out in between washes. Some reports say PLO is toxic to neurons and hence the three washes with DPBS are essential. 23. The pH of N2 medium is critical, discard when media looks pink. Mix well without shaking. No need to filter. Store the media at 4 °C for up to 3 weeks. Avoid exposure to light. 24. Washing with DPBS will include resuspending the pellet in 1 mL DPBS, centrifugation at 140 g for 5 min at room temperature, and discarding the supernatant completely. This step is important as trace amount of serum can inhibit neuronal differentiation.

Acknowledgments M.R.S Rao acknowledges the Department of Science and Technology, Government of India, for SERB Distinguished Fellowship and SERB-YOS (Year of Science Chair Professorship). Debosree Pal thanks the University Grants Commission, Government of India, and JNCASR, India, for her PhD fellowship. Sangeeta Dutta thanks the Department of Biotechnology, Government of India, for her postdoctoral fellowship.

Funding This work was financially supported by the Department of Biotechnology, Government of India (Grant Numbers: BT/01/COE/07/ 09 and DBT/INF/22/SP27679/2018). M.R.S.R. acknowledges the Department of Science and Technology for J. C. Bose and S.E.R.B. Distinguished fellowships and the Year of Science Chair professorship. References 1. Evans MJ, Kaufman MH (1981) Establishment in culture of pluripotential cells from mouse embryos. Nature 292(5819):154–156 2. Martin GR, Evans MJ (1975) Differentiation of clonal lines of teratocarcinoma cells:

formation of embryoid bodies in vitro. Proc Natl Acad Sci 72(4):1441–1445 3. Chuang JH, Tung LC, Lin Y (2015) Neural differentiation from embryonic stem cells in vitro: an overview of the signaling pathways. World J Stem Cells 7(2):437

RA Induced Neural Differentiation of mESCs 4. Bibel M, Richter J, Schrenk K, Tucker KL, Staiger V, Korte M, Goetz M, Barde YA (2004) Differentiation of mouse embryonic stem cells into a defined neuronal lineage. Nat Neurosci 7(9):1003–1009 5. Plachta N, Bibel M, Tucker KL, Barde YA (2004) Developmental potential of defined neural progenitors derived from mouse embryonic stem cells. Development 131(21): 5449–5456 6. Bibel M, Richter J, Lacroix E, Barde YA (2007) Generation of a defined and uniform population of CNS progenitors and neurons from mouse embryonic stem cells. Nat Protoc 2(5): 1034–1043 7. Kin Ting Kam R, Deng Y, Chen Y, Zhao H (2012) Retinoic acid synthesis and functions in early embryonic development. Cell Biosci 2:1– 14 8. Mark M, Ghyselinck NB, Chambon P (2009) Function of retinoic acid receptors during embryonic development. Nucl Recept Signal 7(1):nrs-07002 9. Zile MH (1998) Vitamin A and embryonic development: an overview. J Nutr 128(2): 455S–458S 10. Moutier E, Ye T, Choukrallah MA, Urban S, Osz J, Chatagnon A, Delacroix L, Langer D, Rochel N, Moras D, Benoit G (2012) Retinoic acid receptors recognize the mouse genome through binding elements with diverse spacing and topology. J Biol Chem 287(31): 26328–26341 11. Rosenfeld MG, Lunyak VV, Glass CK (2006) Sensors and signals: a coactivator/corepressor/epigenetic code for integrating signaldependent programs of transcriptional response. Genes Dev 20(11):1405–1428

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12. Piskunov A, Rochette-Egly C (2012) A retinoic acid receptor RARα pool present in membrane lipid rafts forms complexes with G protein αQ to activate p38MAPK. Oncogene 31(28):3333–3345 13. Stavridis MP, Collins BJ, Storey KG (2010) Retinoic acid orchestrates fibroblast growth factor signalling to drive embryonic stem cell differentiation. Development 137(6):881–890 14. Rochette-Egly C (2015) Retinoic acid signaling and mouse embryonic stem cell differentiation: cross talk between genomic and non-genomic effects of RA. Biochim Biophys Acta 1851(1):66–75 15. Al Tanoury Z, Gaouar S, Piskunov A, Ye T, Urban S, Jost B, Keime C, Davidson I, Dierich A, Rochette-Egly C (2014) Phosphorylation of the retinoic acid receptor RARγ2 is crucial for the neuronal differentiation of mouse embryonic stem cells. J Cell Sci 127(9):2095–2105 16. Piskunov A, Al Tanoury Z, Rochette-Egly C (2014) Nuclear and extra-nuclear effects of retinoid acid receptors: how they are interconnected. In: The biochemistry of retinoic acid receptors I: Structure, activation, and function at the molecular level, pp. 103–127 17. Pal D, Dutta S, Iyer DP, Shriram D, Bhaduri U, Rao MRS (2022) Identification of PAX6 and NFAT4 as the transcriptional regulators of the long noncoding RNA Mrhl in neuronal progenitors. Mol Cell Biol 42(11):e00036– e00022 18. Pal D, Neha CV, Bhaduri U, Zenia Z, Dutta S, Chidambaram S, Rao MRS (2021) LncRNA Mrhl orchestrates differentiation programs in mouse embryonic stem cells through chromatin mediated regulation. Stem Cell Res 53: 102250

Methods in Molecular Biology (2023) 2736: 53–64 DOI 10.1007/7651_2022_472 © Springer Science+Business Media, LLC 2023 Published online: 08 February 2023

Isolation and Functional Analysis of Myoepithelial Cells from Adult Mouse Submandibular Glands Rika Yasuhara, Seya Kang, Rino Tokumasu, and Kenji Mishima Abstract Salivary gland myoepithelial cells regulate salivary secretion and have been implicated in the histological diversity of salivary gland tumors. However, isolation of myoepithelial cells has been difficult owing to a lack of detailed functional analysis and cell surface markers. Therefore, we aimed to isolate myoepithelial cells from adult mouse submandibular glands using the epithelial marker EpCAM and cell adhesion factor CD49f as indicators and characterize them via sphere-forming culture. Functional analysis of specific gene expression in myoepithelial cells is possible via cell transfection experiments using the piggyBac transposon vector system. Here, we describe detailed methods and tips for the isolation and functional analysis of myoepithelial cells. Key words Myoepithelial cells, piggyBac transposon vector system, Primary culture, Salivary glands, Sphere

1

Introduction Salivary myoepithelial cells are terminally differentiated cells that regulate the secretion of saliva from acinar cells. Furthermore, they are thought to be involved in tissue diversity in salivary gland tumors [1, 2]. To identify the myoepithelial cell properties in exocrine glands such as mammary glands, the specific fraction expressing several cell surface markers as seen in stem cells, including CD24, CD29, CD10, CD44, and CD49f, was used [3– 6]. However, it is still controversial whether myoepithelial cells have multipotency in a physiological condition. Myoepithelial cells have both epithelial and myoepithelial properties and are located near the basement membrane [2, 7]. Therefore, we identified them using the epithelial cell adhesion molecule (EpCAM) and heterodimeric integral membrane proteins CD49f/ITGA6 (laminin receptor) [5, 8]. The comprehensive transcriptomic analysis from RNA-seq showed that EpCAM-low and CD49f-high cells expressed myoepithelial specific markers, including αSMA, Krt14, Tp63, and Snai2 [8]. Here, we report the detailed protocol of primary and secondary cell line culture of myoepithelial cells from mouse submandibular glands. Additionally, we describe specific

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gene expression in myoepithelial cells using notch intercellular domain (NICD), which is highly expressed in salivary duct carcinoma, by using the piggy-bac transposon vector system. Salispheres, which are cell aggregates enriched with salivary gland stem cells, are utilized in a self-renewal capacity to expand salivary gland stem cells [6]. In contrast, organoids, which are three-dimensional mini organs, are developed to assess cell differentiation from stem/progenitor cells [9]. These cultures use similar methods but different extracellular matrices and niche factors. A recent study showed a salivary gland organoid culture with ductal-, acinar-, and myoepithelial-cell differentiation that regulated Wnt, BMP/TGFβ, FGF, and Notch signals from these salivary gland cells [9]. Our current myoepithelial single culture is yet to develop a form; however, the application of various signaling factors may enhance culture development. Here, we describe the basic setup for a spheroid culture of myoepithelial cells, including some tips and tricks for successful culture management.

2 2.1

Materials Reagents

2.1.1 Mice

2.1.2 Culture

1. C57BL/6 J female mice were purchased from Sankyo Laboratory (Tokyo, Japan). 2. Trp53-null (p53-/-) mice were provided by the RIKEN BioResource Center (RBRC01361; Tsukuba, Ibaraki, Japan). 1. Dulbecco’s modified Eagle’s medium (DMEM)/F12HAM (D8437, Sigma–Aldrich). 2. Keratinocyte SFM (#17005042, Gibco). 3. Fetal bovine serum (FBS, #SH30084.03, HyClone, Thermo Fisher). 4. Penicillin–Streptomycin (#15140122, Gibco). 5. HBSS (#14175095, Gibco). 6. PBS (#T9181, Takara). 7. HEPES (#15630-080, Gibco). 8. Collagenase type I (#03517604, Wako). 9. Hyaluronidase (#H3506-5G, Sigma–Aldrich). 10. DNaseI (#DN25-1G, Sigma–Aldrich). 11. TrypLE Express (#12605-010, Gibco). 12. Dispase (2.6 U/mL, #354235, Corning). 13. Bovine serum albumin (BSA; #A1470-10G, Sigma–Aldrich). 14. Cholera toxin (Fujifilm Corporation). 15. ROCK inhibitor (Y-27632) (#SCM075, Sigma–Aldrich).

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16. Matrigel growth factor reduced (#354230, Corning). 17. G418/Geneticin (#11811031, Gibco). 18. Doxycycline (#D5897, LKT Labs). 19. Epidermal growth factor (EGF; #AF-100-15, Peprotech). 20. Fibroblast growth Peprotech).

factor

2

(FGF-2;

#AF-100-18B,

21. N2 supplement (#17502048, Gibco). 22. Insulin (#I0516, Sigma–Aldrich). 23. Dexamethasone (#D4902, Sigma–Aldrich). 24. Hydrocortisone (#H4001-1G, Sigma–Aldrich). 2.1.3 DNA Cloning

1. Gateway LR Invitrogen).

Clonase

II

Enzyme

mix

(#11791020,

2. One Shot®OmniMAX™2 T1 Phage-Resistant Cells (#C854003, Invitrogen). 3. pENTR-NICD1 (#46048, Addgene). 4. PB-TAC-ERN (#80475, Addgene). 5. Super PiggyBac Transposase Expression Vector (#PB210PA-1, System Biosciences). 2.1.4 FACS

1. Fluorescein isothiocyanate (FITC)-conjugated anti-TER-119 antibody (#116205, BioLegend). 2. FITC-conjugated anti-CD31 antibody (#102405, BioLegend). 3. FITC-conjugated anti-CD45 antibody (#103107, BioLegend). 4. PE/Cy7-conjugated BioLegend).

anti-EpCAM

antibody

(#118215,

5. APC-conjugated anti-CD49f antibody (#313615, BioLegend). 6. Fixable Viability Dye eFluor 450 (#65-0863-14, eBioscience). 2.1.5 Immunofluorescence

1. Cell recovery solution (#354253, Corning). 2. 4% PFA (#16320145, Wako). 3. Sucrose (#19300025, Wako). 4. Optimal cutting temperature; OCT compound (#4583, Sakura Finetek). 5. Rabbit anti-Ki-67 Bioscience). 6. Mouse anti-Krt18 Biotechnik).

antibody antibody

(1:200;

#M3060,

Spring

(1:50;

#61028,

Progen

7. Rabbit anti-Krt5 antibody (1:200; #ab52635, Abcam). 8. Rabbit anti-NKCC antibody (1:200; #8351, Cell Signaling Technology).

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9. Rabbit anti-Aqp5 Alomone Labs).

antibody

(1:200;

#AQP-005,

10. Rat anti-E-cadherin antibody (1:200; #114420, R&D Systems). 11. Mouse anti-αSMA antibody (1:200; #ab7817, Abcam). 12. 4′,6-diamidino-2-phenylindole; DAPI (1:1000; #349-91331, Dojindo). 2.2

Equipment

1. Dissecting forceps—sterilize by autoclave. 2. Dissecting scissors—sterilize by autoclave. 3. 90 × 20 mm Petri dishes (#SH9020, IWAKI). 4. 100-mm tissue culture dishes (#93100, TPP), 6-well (#92006, TPP), 12-well (#92012, TPP), and 96-well (#92096, TPP) plates. 5. Nunclon Sphera 12-well plates (#174931, Thermo) *ultra-low attachment plate. 6. 5 mL disposable pipettes (#606160, Greiner Bio-One), 10 mL (#607160,Greiner Bio-One). 7. Pathtool pipettes (#731202, Iwaki). 8. Millex-GP Filter, 0.22 μm pore size (#SLGVM33RS, Millipore). 9. 0.7 μm cell strainer (#542070, Greiner Bio-One) and 0.4 μm cell strainer (#542040, Greiner Bio-One). 10. Cell Sorter SH1800 (Sony Biotechnology, Tokyo, Japan). 11. Neon Transfection System (Invitrogen). 12. CryoStar NX70 (PHC). 13. Microtome Scientific).

blades

(HP35

Ultra,

#3153735,

Thermo

14. Micro slides (#5116, Muto Pure Chemicals). 2.3

Reagent Setup

1. Mixture of three anesthetic agents A mixture of 0.375 mL domitor (1 mg/mL), 1 mL midazolam (5 mg/mL), 1.25 mL butorphanol (5 mg/mL), and saline was prepared up to 10 mL. 2. Matrigel coating the dish (1:60) Prepare a mixture of 50 μL Matrigel and 3 mL PBS (1:60 dilution). Halfway filling the diluted Matrigel into the well of 6-well tissue culture plates and incubate for 1 h at 37 °C. 3. Enzymatic digestion medium (10 mL) Dissolve a mixture of 7500 U collagenase type I (final conc. 750 U/mL), 5000 U hyaluronidase type IV (final conc. 500 U/mL), and DNase I (final conc. 0.1 mg/mL) in 10 mL of DMEM/F12HAM filtered through 0.22 μm pores.

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4. 2% FBS in HBSS (10 mL) Prepare a mixture of 0.2 mL FBS, 0.1 mL HEPES, and 9.7 mL HBSS. 5. Primary culture medium [5] Prepare a mixture of DMEM/F12HAM containing 10% FBS, hydrocortisone (0.5 μg/mL), cholera toxin (10-10 M), EGF (10 ng/mL), insulin (5 μg/mL), ROCK inhibitor (10 μM), antibiotics, and antimicrobials (see Note 1). 6. Cell line culture medium Prepare a mixture of Keratinocyte SFM supplemented with EGF and BPE, antibiotics, and antimicrobials. 7. Sphere culture medium Prepare a mixture of DMEM/F12HAM supplemented with EGF (20 ng/mL), FGF-2 (20 ng/mL), N2 supplement (1/100), insulin (10 μg/mL), dexamethasone (1 μM), antibiotics, and antimicrobials.

3

Methods

3.1 Primary Culture of Myoepithelial Cells from Adult Mouse Submandibular Glands 3.1.1 Preparation of Submandibular Glands [Timing: 10 min]

1. For mouse anesthesia, inject 100 μL per 10 g weight of a mixture of three anesthetic agents in mice via intraperitoneal injection (see Note 2). 2. Make a 10 mm vertical incision in the neck skin with surgical scissors and carefully detach fibrous connecting tissues around the submandibular glands (SMG) (Fig. 1a). Excise SMGs after making sure the entire structure. 3. Transfer SMGs to a 100 mm petri dish with 10 mL PBS on ice and wash three times with PBS, before mincing to approximately 5 mm pieces with scissors.

3.1.2 Isolation of Single Cells with Enzymatic Digestion [Timing: 90 min]

4. Transfer the minced SMGs into a 50 mL conical tube containing digestion medium (10 mL per three mice) with collagenase type I (750 U/mL), hyaluronidase (500 U/mL), and DNase I (0.1 mg/mL) for 45 min at 37 °C with shaking (at a speed of 100/min) in a water bath. Pipette the tissues up and down every 10 min with a path tool pipette. 5. Centrifuge at 100 × g for 20 sec at 4 °C. 6. Aspirate the supernatant and wash twice with 10 mL of DMEM/F12HAM and repeat step 5. 7. After washing the cells, add 5 mL of TrypLE Express and 50 μL of DNaseI(10 mg/mL), carefully pipette with P200, and dissociate for 10 min at 37 °C with shaking (at 100/min speed) in the water bath. Add 50 μL DNase I (10 mg/mL) every 2 min (see Note 3).

Fig. 1 (a) Gross appearance of mouse salivary glands; (b) scatter plot of flow cytometry; (c) culture of primary cells (day 1); (d) culture of primary cells (day 7); (e) PiggyBac Expression vector; (f) sphere culture; (g) sphere (hematoxylin and eosin staining)

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8. Centrifuge at 300 × g for 5 min at 4 °C. 9. Aspirate the supernatant and wash twice with 10 mL of DMEM/F12HAM and repeat step 8. 10. After washing the cells, add 1 mL of Dispase (2.6 U/mL), 20 μL of DNase I (10 mg/mL), and 9 mL of DMEM/ F12HAM (10% FBS) to the culture tube and carefully pipette with P200 for 1 min. 11. Centrifuge at 350 × g for 5 min at 4 °C. 12. Aspirate the supernatant and wash with 10 mL DMEM/ F12HAM containing 10% FBS, before centrifugation at 350 × g for 5 min at 4 °C. 13. Aspirate the supernatant and resuspend cells in 10 mL DMEM/F12HAM containing 10% FBS. Filter through a 70 μm filter into a new 50 mL tube and then filter through a 40 μm cell strainer into a new 50-mL tube. 14. Count cell numbers. 3.1.3 Flow Cytometry Analysis and Cell Sorting (FACS) [Timing: 90 min]

15. Adjust concentration to 1 × 106 cells per mL with HBSS containing 2% FBS. 16. Stain cells with fluorescent-conjugated antibodies (anti-TER119, anti-CD31, anti-CD45, anti-EpCAM, and anti-CD49f antibodies) at 1:500 dilution for 30 min on ice (see Note 4). 17. Centrifuge at 350 × g for 5 min. Carefully remove the staining buffer and wash twice with 1 mL PBS. 18. Incubate with Fixable Viability Dye eFluor 450 at 1:1000 dilution for 30 min on ice in a light-proof condition. 19. Centrifuge at 350 × g for 5 min. 20. Aspirate the supernatant before adding 500 μL HBSS containing 2% FBS. 21. Sort the myoepithelial cell fractions (EpCAMlow CD49fhigh) after gating the living cells (eFluor 450-negative) and stromal cell populations (TER-119-CD31-CD45-) via Cell Sorter SH800 (Fig. 1b). 22. Transfer sorted cell suspension into a new 1.5 mL tube with fresh medium. 23. Centrifuge at 300 × g for 5 min. 24. Aspirate the supernatant, resuspend with 1 mL of primary culture medium, and count the cell number. At this stage, the cells are used for gene expression analysis of myoepithelial markers such as αSMA and Krt 5.

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3.1.4 Culture of Myoepithelial Cells (Fig. 1c and 1d)

25. Seed the sorted cells at a concentration of 1 × 105 cells per well in Matrigel (GFR)-coated 6-well culture plate and incubate at 37 °C in a 5% CO2 incubator (Passage 0). 26. Keep the culture at 37 °C in 5% CO2 for 48 h. Change the medium every 2–3 days (see Note 5). 27. Expand the culture of primary myoepithelial cells when the cells reach 80–90% confluency. 28. Aspirate the medium and wash the cells with 5 mL PBS. Add 1 mL of TrypLE Express and incubate at 37 °C in 0.5% CO2 for 2 min. 29. Add 9 mL culture medium before centrifugation at 300 × g for 5 min. 30. Aspirate the supernatant and resuspend with 10 mL of culture medium before seeding in a 100-mm tissue culture dish. Incubate at 37 °C in a 5% CO2 incubator until the cells reach 80–90% confluency (Passage 1). Change the medium every 2–3 days (see Note 6).

3.2 Establishment of a Myoepithelial Cell Line from Trp53-Null Mice

Preparation of myoepithelial cells from Tp53-null mice (see Note 7). See Subheading 3.1, steps 1–29. Discard the supernatant and resuspended with 10 mL of Keratinocyte SFM and then seed it into a 100-mm tissue culture dish. Incubate at 37 ° C in a 5% CO2 incubator until the cells reach 80–90% confluency (Passage 1). Change the medium every 2–3 days (see Note 8). 31. Preserve cells with 1 mL of cell banker 2 at a concentration of 1 × 106 cells per cryotube as stock at Passage 2–3.

3.3 Establishment of Dox-Inducible Gene Expression Cell Line 3.3.1 Preparation of Expression Vector (Fig. 1e) (See Note 9)

32. Prepare a mixture of 1 μL of 100 ng/μL entry vector (pENTRNICD1), 1 μL of 150 ng/μL destination vector (PB-TAC-ERN), and 6 μL of TE buffer (pH 8.0) in a 1.5 mL tube. 33. Add 2 μL of LR clonase II and incubate at 25 °C for 1 h. 34. Add 1 μL of Proteinase K solution and incubate at 37 °C for 10 min. 35. Incubate 1 μL of LR reaction from step 34 with 50 μL of One Shot®OmniMAX™2 T1 Phage-Resistant Cells in a new 1.5 mL tube on ice for 30 min. 36. Heatshock the mixture at 42 °C for 30 sec and then quickly place it on ice for 5 min. 37. Add 250 μL of SOC medium to the tube and incubate at 37 °C for 1 h. 38. Spread 100 μL of mixture onto the LB agar plate with calbenicillin and incubate the plate upside-down at 37 °C overnight. 39. On the next day, select suitable colonies and store 1 μg/μL purified DNA.

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40. Cultured myoepithelial cells should be at 80–90% confluency in a 100-mm culture dish on the day of transfection. 41. Harvest cells by adding 2 mL TrypLE (see steps 28–30) and adjust the cell concentrations to 1 × 106 cells/mL in Keratinocyte SFM without antibiotics. 42. Transfer 10 μL of the cell suspension (1 × 104 cells) to a 1.5-mL tube and 10 μL Buffer R was replaced. 43. Add 1.5 μL of the expression vector (PB-NICD) along with 1.5 μL Super PB transposase expression vector. 44. Transfer the total volume of the mixture with a 10-μL NeonTip and set it to a Neon tube filled with 3 mL Buffer E. 45. Electroporate at 1275 V for 20 ms at 1 pulse. 46. Immediately seed the transfected cells in a 12-well culture plate filled with 500 μL pre-warmed Keratinocyte SFM without antibiotics. Maintain culture for 48 h (see Note 10).

3.3.3 Drug Selection

47. Change the medium to Keratinocyte SFM with G418 (0.4 mg/mL). Isolate the stably transfected cell lines via antibiotic selection for 7 days (see Note 11). 48. Change the medium every 2–3 days. 49. Preserve the batch culture cells as stock.

3.3.4 Isolation by Limiting Dilution and Expansion

50. Plate 100 μL of 10 and 100 cells/mL in a 96-well tissue culture plate. 51. After 4 days of culture, extract single-clone wells. 52. Expand the selected single-colony wells in a 96-well tissue culture plate to 80–90% confluency and then culture in 12-well tissue culture plate. 53. Passage the clones into a 6-well tissue culture plate after the cells reach 80–90% confluency. 54. Preserve the monoclonal selections as stocks.

3.3.5 Validation of Inducible Expression

55. Plate the monoclonal selections at a density of 1 × 105 cells/ well in a 12-well tissue culture plate. 56. On the next day, add doxycycline (2 μg/mL) and keep the culture for 48 h (see Note 12). 57. Monitor gene expression with mCherry fluorescent protein using a fluorescent microscope (Keyence) (Fig. 1d). At this stage, the cells are deemed ready for RNA isolation to check for gene expression. 58. Select the clones showing the highest gene expression.

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Assays

3.4.1 Sphere Culture (Fig. 1f and 1g)

59. Aliquot the cultured cells at a density of 1 × 104 cells per 1.5 mL tube. 60. Then, add 50 μL Matrigel GFR and pipette immediately on ice. 61. Drop the cell mixture into the well of 12-well ultra-low attachment plates (Fig. 1f). 62. Place the plate upside down for 20 min at 37 °C. 63. Add 1 mL of DMEM/F12HAM, supplemented with EGF (20 ng/mL), FGF-2 (20 ng/mL), N2 supplement (1/100), insulin (10 μg/mL), and dexamethasone (1 μM) and culture for 7 days or longer. Change the medium every 2–3 days.

3.4.2 Immunohistochemistry

64. Add cell recovery solution to the cultures and keep on ice for 30 min. Then, collect spheres before centrifugation at 350 × g for 5 min. 65. Fix the spheroid in 4% paraformaldehyde overnight. 66. The next day, add 10% sucrose and leave for 1 h, followed by 20% sucrose for 1 h, and finally, 30% sucrose overnight. 67. Embed the fixed spheres in the OCT compound (see Note 13). 68. Section frozen tissues (4 μm). 69. Either stain the sections with hematoxylin–eosin (Fig. 1g) or subject them to immunofluorescence. Counterstain with DAPI (1 μg/mL). Image using a fluorescent microscope (Keyence). Calculate positive areas and signaling intensities automatically using a hybrid cell count application (BZ-H4C, Keyence) in the BZ-X Analyzer software (BZ-H4A, Keyence) (see Note 14).

3.4.3 Tissue-Specific Markers

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E-cadherin, Pan-cytokeratin

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αSMA, Snai2

Basal cells

Krt5, Krt14

Ductal cells

Krt18, Krt19

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Proliferating cells

ki-67, PCNA

Notes 1. ROCK inhibitor (Y-27632) is used for inhibiting apoptosis after single-cell dissociation. It was also reported in stem cell research for usage during single-cell cloning after gene transfection.

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2. SMGs were resected following blood withdrawal to avoid blood contamination in the tissue rather than by cervical dislocation (without prior anesthesia). 3. DNase I was used in assisting single-cell separation, as cells come out sticky after digestion. 4. As CD49f is also expressed in pericyte-like cells, EpCAM/ CD326 was used during selection to assist in cell exclusion. 5. The cells should become visible colonies around day 7. 6. The primary culture of the myoepithelial cells at passage 2 is used for additional assays. 7. p53 gene is frequently mutated in sporadic cancer but p53 deficiency is not thought to affect normal development [10]. 8. The myoepithelial cell line from Tp53-null mice until passage 10 is used for additional assays. 9. The PB transposon system was used for the stable expression of NICD in the cells [8]. 10. Transfected cells can be monitored via GFP fluorescence from the PBase vector. 11. Please do not use other antibiotics with geneticin/G418 to avoid competitive or cross-reaction. Geneticin/G418 selective antibiotic is used for positive cell selection expressing the neomycin resistance (neo) gene. In mammalian cells, geneticin/ G418 is usually achieved in 3–7 days with concentrations ranging from 400 to 1000 μg/mL. It is recommended the optimal concentration of the product should be re-evaluated. 12. The PB-TetOn vector carried a tetracycline response element to drive doxycycline induction. 13. Spheres are kept in the OCT compound for 30 min to preserve the formation of the tissue. 14. Please refer to the original paper for the immunofluorescence results [8]. References 1. Makarenkova HP, Dartt DA (2015) Myoepithelial cells: their origin and function in lacrimal gland morphogenesis, homeostasis, and repair. Curr Mol Biol Reports 1:115–123. https://doi.org/10.1007/s40610-0150020-4 2. Emmerson E, Knox SM (2018) Salivary gland stem cells: a review of development, regeneration and cancer. Genesis 56:e23211. https:// doi.org/10.1002/dvg.23211 3. Shackleton M et al (2006) Generation of a functional mammary gland from a single stem

cell. Nature 439:84–88. https://doi.org/10. 1038/nature04372 4. Sale S, Lafkas D, Artavanis-Tsakonas S (2013) Notch2 genetic fate mapping reveals two previously unrecognized mammary epithelial lineages. Nat Cell Biol 15:451–460. https:// doi.org/10.1038/ncb2725 5. Prater D et al (2014) Mammary stem cells have myoepithelial cell properties. Nat Cell Biol 16: 942–950 6. Centonze A et al (2020) Heterotypic cell-cell communication regulates glandular stem cell

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multipotency. Nature 584:608–613. https:// doi.org/10.1038/s41586-020-2632-y 7. Mattingly A, Finley JK, Knox SM (2015) Salivary gland development and disease. Wiley Interdiscip Rev Dev Biol 4:573–590. https:// doi.org/10.1002/wdev.194 8. Yasuhara R et al (2022) Role of Snai2 and notch signaling in salivary gland myoepithelial cell fate. Lab Investig 102:1245–1256. https://doi.org/10.1038/s41374-02200814-7

9. Yoon YJ et al (2022) Salivary gland organoid culture maintains distinct glandular properties of murine and human major salivary glands. Nat Commun 13:3291. https://doi.org/10. 1038/s41467-022-30934-z 10. Donehower LA et al (1992) Mice deficient for p53 are developmentally normal but susceptible to spontaneous tumours. Nature 356:215– 221. https://doi.org/10.1038/356215a0

Methods in Molecular Biology (2023) 2736: 65–76 DOI 10.1007/7651_2022_477 © Springer Science+Business Media, LLC 2023 Published online: 08 February 2023

Chromosomal Analysis in Lineage-Specific Mouse Hematopoietic Stem Cells and Progenitors Nur Afizah Yusoff, Zariyantey Abd Hamid, Paik Wah Chow, Salwati Shuib, Izatus Shima Taib, and Siti Balkis Budin Abstract Hematopoiesis is maintained throughout life from the hematopoietic stem cell niche in which hematopoietic stem cells and lineage-specific hematopoietic progenitors (HSPCs) reside and regulate hematopoiesis. Meanwhile, HSPCs behavior is modulated by both cell intrinsic (e.g., transcriptional factors) and cell extrinsic (e.g., cytokines) factors. Dysregulation of these factors can alter HSPCs function, leading to disrupted hematopoiesis, cellular changes, and subsequent hematological diseases and malignancies. Moreover, it has been reported that chromosomal aberration (CA) in HSPCs following exposure to carcinogenic or genotoxic agents can initiate leukemia stem cells (LSCs) formation which lays a fundamental mechanism in leukemogenesis. Despite reported studies concerning the chromosomal integrity in HSPCs, CA analysis in lineage-specific HSPCs remains scarce. This indicates a need for a laboratory technique that allows the study of CA in specific HSPCs subpopulations comprising differential hematopoietic lineages. Thus, this chapter focuses on the structural (clastogenicity) and numerical (aneugenicity) form of CA analysis in lineage-specific HSPCs comprised of myeloid, erythroid and lymphoid lineages. In this protocol, we describe how to perform CA analysis in lineage-specific HSPCs derived from freshly isolated mouse bone marrow cells (MBMCs) using the combined techniques of colony-forming unit (CFU) and karyotyping. Prior to CA analysis, lineage-specific HSPCs for myeloid, erythroid, and lymphoid were enriched through colony-forming unit (CFU) assay. CFU assay assesses the proliferative ability and differentiation potential of an individual HSPC within a sample. About 6 to 14 days of cultures are required depending on the type of HSPCs lineage. The optimal duration is crucial to achieve sufficient colony growth that is needed for accurate CFU analysis via morphological identification and colony counting. Then, the CA focusing on clastogenicity and aneugenicity anomalies in respective HSPCs lineage for myeloid, erythroid and Pre-B lymphoid were investigated. The resulted karyotypes were classified according to the types of CA known as Robertsonian (Rb) translocation, hyperploidy or complex. We believe our protocol offers a significant contribution to be utilized as a reference method for chromosomal analysis in lineage-specific HSPCs subpopulations. Key words CFU, Chromosomal analysis, Hematopoietic lineages, Hematopoietic stem cells and progenitors, Karyotyping

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Introduction Bone marrow is the primary site for hematopoietic stem cells (HSCs) niche in which hematopoiesis is regulated [1]. The niche is resided by population of hematopoietic stem cells and lineagecommitted hematopoietic progenitors (HSPCs) that possess the ability to self-renew and differentiate into adult blood cells [2]. Hematopoietic stem cells (HSCs) are a type of primitive cell that has infinite self-renewal ability and pluripotency property, distinguishing them from other hematopoietic cells. In contrast to HSCs, hematopoietic progenitor cells possess short-term hematopoietic repopulation capacity, limited self-renewal ability, and restricted capacity for lineage differentiation. The hematopoietic colony-forming unit (CFU) assay, also known as clonogenic assay, is an in vitro protocol to identify and quantitate the presence of HSPCs with multipotential or restricted lineage differentiation potency to produce hematopoietic cells that consist of myeloid, erythroid, and lymphoid lineages [3–6]. Proliferative and differentiation capacities of HSPCs are defined using CFU assay through colony formation that can be morphologically analyzed [7]. Hematological disorders and malignancies vary according to hematopoietic lineages, for example, syndrome of myelodysplasia (myeloid lineage), aplastic anemia (erythroid lineage), lymphoma (lymphoid lineage), and leukemia (myeloid and lymphoid lineages). Leukemia is a clonal hematological malignancy in which HSPCs undergo uncontrolled cell proliferation and disrupted differentiation leading to accumulation of immature blood-forming cells known as blast cells [8, 9]. It occurs due to genomic alteration such as chromosomal aberration (CA) or epigenetic modification affecting HSPCs [10–12]. Up to our knowledge, molecular studies focusing on CA using lineage-directed strategy targeting HSPCs remain limited and deserve further exploration (Fig. 1). CA status can be analyzed through karyotyping via structural and numerical chromosomal abnormalities [13]. Deletions, duplications, translocations, insertions, and inversions are examples of structural abnormalities, while polyploidy, hyperploidy, and aneuploidy are examples of numerical abnormalities [13]. In order to obtain a reliable result, metaphase spreading is essential technique for CA study as metaphase chromosomes is used for cytogenetics analysis particularly in cancer studies. In conclusion, this chapter describes protocol on how to perform CA analysis in lineagespecific HSPCs from freshly isolated mouse bone marrow cells (MBMCs) using the combined techniques of colony-forming unit (CFU) and karyotyping.

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Fig. 1 Elucidation of the genotoxicity effects in the respective population of lineage-specific HSPCs comprising myeloid, lymphoid, and erythroid lineages focusing on CA in the form of structural and numerical anomalies. Heterogeneous responses of bone marrow cells including HSPCs toward carcinogenic and genotoxic agents have been reported. However, homogeneous responses of lineage-specific HSPCs population remain unexplored. The study focusing on CA using lineage-directed strategy is fundamental to uncover a novel mechanism linking carcinogenic and genotoxic agents to the disruption of genomic integrity affecting HSPCs populations in mediating leukemic stem cells formation and subsequent development of hematological malignancies and disorders

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Materials

2.1 Preparation of Cell Samples

Prepare all solutions with deionized water or ultrapure water (obtained by purifying deionized water to a sensitivity of 18 MΩ-cm at 25 °C) and reagents of analytical grade. Unless otherwise stated, prepare and keep all reagents at room

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temperature. All procedures for cell processing and setup of CFU assays should be performed using sterile technique and universal handling precautions. When disposing of waste materials, follow all safety standards, including the proper trash disposal technique. Pay great attention to any potentially dangerous materials and adhere to the SOP as specified in the lab. We do not add sodium azide to reagents. 2.2 Mouse Bone Marrow Cells (MBMCs) Isolation and Culture

Materials for this section are as follows: 1. Dulbecco’s Modified Eagle Medium (DMEM). 2. 10% Fetal bovine serum (FBS). 3. Penicillin/streptomycin. 4. Stem cells factor (SCF), interleukin-6 (IL-6), and interleukin-3 (IL-3). 5. T-25 or T-75 culture flask or a 6-well culture plate. 6. 40 μM nylon mesh cell strainer.

2.3 Colony-Forming Unit (CFU) Assay

Materials for this section are as follows: 1. Methylcellulose-based medium for hematopoietic CFUs. We advise using methylcellulose-based medium enriched with cytokines that are needed for the growth of lineage-specificHSPCs (myeloid, erythroid, and lymphoid). In current protocol, our CFUs medium are from STEMCELL Technologies. 2. Sterile syringes. 3. 35 mm culture dishes or 6-well culture plates.

2.4 Karyotyping Study

Materials for this section are as follows: 1. KaryoMAX colcemid solution. 2. Trypsin EDTA. 3. Leishman stain. 4. Potassium chloride (KCl). 5. Methanol. 6. Acetic acid. 7. Phosphate-buffered saline (PBS), pH 7.4.

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Methods

3.1 Isolation of Mouse Bone Marrow

Isolate mouse bone marrow cells using flushing technique through tibia and femur [14–16] in laminar flow hood or biosafety cabinet (see Note 1). Next, by using a nylon mesh cell strainer with a size of 40 μM (BD Biosciences, San Diego, CA, USA), filter the collected bone marrow cells and suspend them in DMEM medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/ streptomycin as well as growth factors such as 10 ng/mL interleukin-6 (IL-6), 5 ng/mL interleukin-3 (IL-3), and 100 ng/ mL stem cells factor (SCF) [17–19]. Culture mouse bone marrow cells overnight prior to the experiment in a humidified incubator at 37 °C and 5% CO2. Step-by-step protocols are as illustrated (Fig. 2).

3.2 Preparation of Methylcellulose Media

Allow the addition of cells at 1:10 (v/v) ratio to make up a complete methylcellulose medium to maintain the optimal viscosity of the medium. 1. Thaw complete MethoCult™ medium bottle overnight at 2–8 °C or at room temperature (15–25 °C) until it is completely thawed (approximately within 1 h).

Fig. 2 Procedure for isolation of mouse bone marrow cells using flushing method

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2. Before aliquoting, shake ferociously for 1–2 min and then let the bottle stand for at least 5 min so that the bubbles can reach the top. 3. Using a 3 mL or 6 mL luer lock syringe fitted to a 16-gauge blunt-end needle, dispense methylcellulose medium into 14 mL (17 × 95 mm) sterile tubes (see Note 2). 4. Add 3 mL or 4 mL per tube for 1.1 mL duplicate or triple cultures, respectively (see Note 3 and 4). 5. Vortex tubes to thoroughly mix. Complete medium can be kept at -20 °C for extended storage, 2–8 °C for immediate use, or 2–8 °C for up to 1 month. Mix thoroughly and use immediately after thawing. Avoid refreezing. 3.3 Enrichment of HSPCs Using ColonyForming Unit (CFU)

Maintain the growth of respective lineage-specific HSPCs using specialized methylcellulose-based medium enriched with specific growth factors (StemCells Technologies, Vancouver, BC, Canada). Culture three lineages of HSPCs in selective medium that support the growth of specific lineage as follows: (i) erythroid [(MethoCult media #03334) (colony-forming uniterythroid (CFU-E) and mature burst-forming unit erythroid (BFU-E)], (ii) myeloid [(MethoCult media #03534) (colonyforming unit-granulocyte macrophage (CFU-GM), colony-forming unit-granulocyte (CFU-G), and colony-forming unit macrophage (CFU-M)], and (iii) Pre-B lymphoid [(MethoCult #03630) (colony-forming unit-Pre-B (CFU-Pre-B)]. Perform the CFU assay according to the manufacturer’s instructions (STEMCELL Technologies). 1. Briefly, collect the 24 h MBMCs and centrifuge at 2500 rpm for 7 min. Then, for erythroid, myeloid, and Pre-B lymphoid CFU assays, prepare 100 μL of cell suspension with a viable cell number of 1 × 105, 2 × 104, and 5 × 104 through cell counting respectively. 2. Then, after mixing completely, transfer the mixture of cell suspension with 1 mL of lineage-specific methylcellulose culture media into a 6-well plate (see Notes 5 and 6). 3. Culture MBMCs according to specific progenitors such as 7 days (CFU-E, mature BFU-E, and Pre-B lymphoid) and 14 days (CFU-GM, CFU-G, and CFU-M) at 37 °C with 5% CO2 in incubator as per recommendation (STEMCELL Technologies, Vancouver, BC, Canada) to obtain desirable colony growth suitable for analysis (Fig. 3) (see Notes 7 and 8).

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Fig. 3 Colony-forming unit (CFU) for differential lineages of mouse HSPCs (myeloid, Pre-B lymphoid, and erythroid) as observed under an inverted light microscope. Representative images are for (a) CFU-E (erythroid), (b) BFU-E (burst-forming unit erythroid), (c) CFU-GM (granulocyte and macrophage), (d) CFU-G (granulocyte), (e) CFU-M (macrophage), and (f) CFU-Pre-B (Pre-B lymphoid). (Magnification: 40×)

3.4 Analysis of Lineage-Specific HSPCs from CFU Assay

Colony for respective lineage-specific HSPCs derived from CFUs assay can be visualized under an inverted light microscope. Analyze the colony for respective lineage-specific HSPCs based on morphological characteristic as describe in Table 1.

3.5 Slide Preparation for Karyotyping Analysis

Establish colonies for respective lineage-specific HSPCs following 7 or 14 days of CFU assays. Then proceed to downstream protocols as follows: 1. Arrest cells in metaphase by using colcemid treatment. To do this, mix 0.05 μg/mL of colcemid in a 3 mL of DMEM and add the mixture into 6-well plates containing CFU-derived colonies (see Note 9). 2. Continue incubation of culture overnight prior to colonies harvesting. 3. Harvest colonies of respective lineage-specific HSPCs by centrifuging the cells at 2500 rpm for 7 min, followed by 20 min of incubation with 0.075 M KCl hypotonic solution in a 37 °C water bath (see Note 10). 4. Centrifuge the cells at 2500 rpm for 5 min and fix in 2 mL of methanol: acetic acid at 3:1 ratio (fixative).

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Table 1 A brief overview of morphological characteristics for respective CFUs CFU

Morphological characteristics

CFU-E

CFU-E produces a colony containing 8–200 erythroblasts with 1–2 cell clusters

BFU-E

BFU-E is more immature erythroid progenitor than CFU-E and produces a colony containing >200 erythroblasts in a single or multiple clusters

CFU-GM CFU-GM is more primitive myeloid progenitor than CFU-G and CFU-M and produces a colony with >40 cells of granulocytes and macrophages in a single or multiple clusters CFU-M

CFU-M produces a colony containing macrophages with irregular shape and size

CFU-G

CFU-G produces a colony containing round and small granulocytes with denser colony as compared to CFU-M

CFU-Pre- CFU Pre-B produces colonies with variation in size and morphology. Colony consists of tiny, B round, or oval-shaped cells and may appear as compact colonies or diffuse colonies with peripheral cells

Fig. 4 Slide preparation for karyotyping analysis

5. On pre-cleaned microscopic slides, drop the suspended cells carefully (see Note 11). 6. Then stain the slides with Leishman’s stain (see Notes 12 and 13) (Fig. 4).

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Score twenty evenly distributed metaphases from the provided slides for each lineage-specific HSPC using the following procedures: 1. Using a light microscope, observe the slides and capture each metaphase by a camera that was attached to the microscope. 2. Then, use GenASIs Bandview Software (Applied Spectral Imaging, Carlsbad, CA, USA) to analyze the images of the chromosomes (see Note 14). 3. Score CA according to structural and numerical anomalies. Examples of structural and numerical chromosomal anomalies as presented by Robertsonian (Rb) translocation, hyperploidy, and complex karyotypes are as shown in Fig. 5 (see Note 15). 4. Calculate the percentage of CA as per the following formula: Total CA × 100 Total metaphase

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Notes 1. As closely as possible to the joints, cut both ends of the bone in a laminar flow hood while using sterile equipment to prevent contamination. 2. In order to remove the air from the syringe, insert the needle just below the methylcellulose medium’s surface and draw up around 1 mL. Expel the medium entirely by gently depressing the plunger. Repeat until no visible air space remains. 3. When aliquoting, do not expel the medium to the “0” point on the syringe. Instead of measuring from 3.0 mL to 0 mL, measure from 3.5 mL to 0.5 mL. 4. To prevent the bottle from being frozen and thawed again, it is advisable to pour the entire contents of the bottle into several tubes. 5. To avoid cross-contamination, use a new sterile, disposable 3 mL syringe connected with a new 16-gauge blunt-end needle for each tube plated. 6. Allow the mixture to evenly separate between the compartments of the flask for 3–5 min on a vertical stand. This ensures that the entire bottom layer of the flask receives an equal volume of the mixture, which will cover all surfaces. When setting the flask horizontally, move quickly yet gently to avoid detaching the cells. It might not completely cover the surface of the top compartment if it moves too slowly.

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Fig. 5 The spread and karyotyped metaphase chromosome. Representative images of (a) normal chromosome followed by chromosomal anomalies with (b) Robertsonian (Rb) translocation, (c) hyperploidy, and (d) complex karyotypes

7. Myeloid progenitors need about 14 days to fully grow into colonies that can be morphologically identified and well differentiated for their respective myeloid series (granulocytes and macrophages). Meanwhile, the erythroid (CFU-E and mature BFU-E) and Pre-B lymphoid progenitors were collected on day 7, as colonies’ quality would be deteriorated after this time due to nutrients depletion and pH changes caused by the buildup of cellular metabolic product.

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8. Proper CFU growth requires optimal culture conditions. It is recommended to use humidified incubators with a water pan in the chamber and to monitor temperature and CO2 levels on a regular basis. 9. Inadequate colcemid incubation times result in fewer metaphase spreads and overlapping chromosomes. Longer colcemid incubation times result in shorter and thicker chromosomes, making analysis challenging. 10. The molarity of the hypotonic solution is an additional crucial factor. A 0.075 M KCl solution swells the cells just enough to allow adequate chromosomal spreading while not lysing them. 11. When dropping the resuspended cell pellet onto slides, ensure that the slide is slanted at a 45° angle and that there is adequate space (at least 2 inches) between the dropper and the slide for the chromosomes to spread onto the slide for analysis. 12. Another method to promote chromosomal spread is to quickly cover the slide in fixative once the cells have been placed on it. 13. Depending on the methodology, the temperature during drying the slides can range from 20 °C to 75 °C. A greater temperature may cause the fixative to dry faster. A cold temperature may generate an increase in wetness on slides. As a result, the fixative may dry too slowly. Because drying time affects chromosome spreading, it is critical to determine the right temperature and humidity combination to optimize chromosomal spreading for the cells. 14. Based on our previous study [20], hyperploidy can be notable in the negative control group (untreated with 1,4-BQ) of erythroid progenitor. Ex vivo culture condition may cause the cells to have proliferative stress. Thus, bear in mind that there is a potential of acquired DNA damage in this progenitor when cultured under conditions of proliferative stress. 15. The total number of 40 chromosomes including chromosomes X and Y were classified as normal chromosomes and represented as 40, XY. On the other hand, the chromosome was classified as CA when one of the following CA composed of Rb (structural CA), hyperploidy (numerical CA), and complex karyotype (presence of both structural and numerical CA) was detected.

Acknowledgments This work was funded by the Ministry of Higher Education of Malaysia under the Fundamental Research Grants Scheme (FRGS) with grant number: FRGS/1/2021/SKK06/UKM/03/1. The work has been carried out at the Centre of Diagnostic, Therapeutic & Investigative Studies, Universiti Kebangsaan Malaysia, Kuala Lumpur, Malaysia.

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References 1. Passegue´ E, Jamieson CH, Ailles LE et al (2003) Normal and leukemic hematopoiesis: are leukemias a stem cell disorder or a reacquisition of stem cell characteristics? Proc Natl Acad Sci U S A 100:11842–11849 2. Huang X, Cho S, Spangrude GJ (2007) Hematopoietic stem cells: generation and selfrenewal. Cell Death Differ 14:1851–1859 3. Eaves C (1995) Assays of hematopoietic progenitor cells. In: Beutler E, Lichtman MA, Coller BS, Kipps TJ (eds) Williams hematology, 5th edn. McGraw-Hill, New York, pp 22–66 4. Klug CA, Jordan CT (eds) (2002) Hematopoietic stem cell protocols, vol 63. Humana Press, Totowa 5. Miller CL, Lai B (2005) Human and mouse hematopoietic colony-forming cell assays. Methods Mol Biol 290:71–89 6. Franken NA, Rodermond HM, Stap J et al (2006) Clonogenic assay of cells in vitro. Nat Protoc 1:2315–2319 7. Eaves CJ (2015) Hematopoietic stem cells: concepts, definitions, and the new reality. Blood 125:2605–2613 8. Thomas X (2009) Targeting leukemia stem cells: the new goal of therapy in adult acute myeloid leukemia. World J Stem Cells 1:149– 154 9. Apidi E, Wan Taib WR, Hassan R et al (2018) A review on effect of genetic features on treatment responses in acute myeloid leukemia. Meta Gene 18:31–38 10. Pei-Shen A, Ramasamy R, Hussin NH et al (2016) Differential expression patterns of leukaemia associated genes in leukaemia cell lines compared to healthy controls. Malaysian J Med Health Sci 12:33–45 11. Tsiftsoglou AS, Bonovolias ID, Tsiftsoglou SA (2009) Multilevel targeting of hematopoietic stem cell self-renewal, differentiation and apoptosis for leukemia therapy. Pharmacol Ther 122:3264–3280 12. Zhang L, Lan Q, Ji Z et al (2012) Leukemiarelated chromosomal loss detected in

hematopoietic progenitor cells of benzeneexposed workers. Leukemia 26:2494–2498 13. Ozkan E, Lacerda MP (2022) Genetics, cytogenetic testing and conventional karyotype. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing. https://www.ncbi.nlm. nih.gov/books/NBK563293/. Accessed 10 Nov 2022 14. Chow PW, Rajab NF, Chua KH et al (2018) Differential responses of lineage-committed hematopoietic progenitors and altered expression of self-renewal and differentiation-related genes in 1,4-benzoquinone (1,4-BQ) exposure. Toxicol In Vitro 46:122–128 15. Faiola B, Fuller ES, Wong VA et al (2004) Exposure of hematopoietic stem cells to benzene or 1,4-benzoquinone induces genderspecific gene expression. Stem Cells 22:750– 758 16. Tian JF, Peng CH, Yu XY et al (2012) Expression and methylation analysis of p15 and p16 in mouse bone marrow cells exposed to 1,4-benzoquinone. Hum Exp Toxicol 31: 718–725 17. Chow PW, Abdul Hamid Z, Chan KM et al (2015) Lineage-related cytotoxicity and clonogenic profile of 1,4-benzoquinone-exposed hematopoietic stem and progenitor cells. Toxicol Appl Pharmacol 284:8–15 18. Hamid ZA, Tan HY, Chow PW et al (2018) The role of N-Acetylcysteine supplementation on the oxidative stress levels, genotoxicity and lineage commitment potential of ex vivo murine haematopoietic stem/progenitor cells. Sultan Qaboos Univ Med J 18:130–136 19. Chan CY, Hamid Z, Taib IS et al (2018) Effects of n-acetyl-cysteine supplementation on ex-vivo clonogenicity and oxidative profile of lineage-committed hematopoietic stem/ progenitor cells. J Teknol 80 20. Chow PW, Abd Hamid Z, Mathialagan RD et al (2021) Clastogenicity and aneugenicity of 1,4-benzoquinone in different lineages of mouse hematopoietic stem/progenitor cells. Toxics 9:10

Methods in Molecular Biology (2023) 2736: 77–84 DOI 10.1007/7651_2023_479 © Springer Science+Business Media, LLC 2023 Published online: 05 May 2023

Easy and Rapid Methods for Human Umbilical Cord Blood– Derived Mesenchymal Stem Cells and Human Umbilical Wharton’s Jelly–Derived Mesenchymal Stem Cells Figen Abatay Sel, Ayse Erol, Mediha Suleymanoglu, Durdane Serap Kuruca, and Fatma Savran Oguz Abstract These protocols describe modified methods that use Ficoll-Paque density gradient for umbilical cord blood–derived mesenchymal stem cells and explant method for Wharton’s jelly–derived mesenchymal stem cells. The Ficoll-Paque density gradient method allows to obtain mesenchymal stem cells while eliminating monocytic cells. In this method, precoating the cell culture flasks with fetal bovine serum helps remove the monocytic cells and instruct more pure mesenchymal stem cells. On the other hand, the explant method for Wharton’s jelly–derived mesenchymal stem cell is user-friendly and cost-effective than enzymatic methods. In this chapter, we provide a collection of protocols to obtain mesenchymal stem cells from human umbilical cord blood and Wharton’s jelly. Key words Explant method, Ficoll-Paque density gradient, Mesenchymal stem cell, Umbilical cord blood, Wharton’s jelly

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Introduction In an attempt to bring mesenchymal stem cells (MSCs) closer to reaching its application in cell therapy, many researchers have focused on obtaining MSC protocols over the years. In the beginning of 1970s, the first MSCs were characterized [1]. In recent years, cell based-therapies have been an exciting field to diminish the pain and symptoms especially for immune-mediated disorders [2–6]. Obtaining MSCs has advantages compared to other stem cells such as induced pluripotent stem cells and hematopoietic stem cells. The isolation of umbilical cord–derived MSC can be achieved by noninvasive procedures during the delivery of an infant. In addition to this advantage, MSC can be obtained from many autologous and allogeneic tissues such as adipose [7], dental pulp [8], cord blood [9], and Wharton’s jelly [10]. They can be

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distinguished with the features for ability to adhere to plastic inherently, and surface markers such as CD29+, CD44+, CD73+, CD90+, CD105+ and lack of CD14-, CD34-, CD45- and HLA (human leukocyte antigen)-DR from other stem cells [11]. The differentiation features of MSC make itself a particularly unique cell group for regenerative medicine. MSC can differentiate into different cell lineages such as osteocytes [12], chondrocytes [13], adipocytes [14], myocytes [15], astrocytes [16] and epithelia [17]. This potential of MSC makes it a valuable tool not only for regenerative medicine but also for tissue engineering. In addition to the differentiation features, MSC can secrete cytokines, anti-inflammatory, and pro-inflammatory molecules. Many researchers have reported that MSC can ensure immunomodulation for many diseases [18–20]. It also has low immunogenicity due to the lack of major histocompatibility complex (MHC) and T-cell recruitment. Thus, the low immunogenicity of MSC makes it safe for allogeneic transplantation. Furthermore, MSC has been used for cancer therapy as a “Trojan horse” [21, 22]. It is believed that MSC can change tumor microenvironments with chemical agent secretion and can be used as drug or drug carrier. In particular, there are numbers of studies for leukemia [23–25]. We have recently reported the advantages of mesenchymal stem cell application for several diseases and cancer [26]. Here, we describe in detail the obtaining of human cord blood and human Wharton’s jelly derived–MSC protocols.

2

Materials 1. Human umbilical cord blood (see Note 1).

2.1 Biological Materials

2. Human umbilical cord (Wharton’s jelly) (see Note 1).

2.2

1. 100-μL PIPPETMAN.

Equipment

2. 1000-μL PIPPETMAN. 3. Biosafety cabinet (Fagus). 4. Hemocytometer (Neubauer). 5. Inverted microscope (Motic, AE21). 6. Culture incubator (New Brunswick, Galaxy 170R). 7. Falcon tube, 15 mL (Corning, Sigma Aldrich). 8. Falcon tube, 50 mL (Corning, Sigma Aldrich). 9. Pasteur pipette, glass, 5 mL (Thomas Scientific). 10. Pasteur pipette bulb for glass pipette (Isolab). 11. Disposable Pasteur pipette, polyethylene (Isolab). 12. Dissecting scissors, sharp pointed (Thermo Fisher Sci).

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13. Dissecting scissors, curved pointed (Thermo Fisher Sci). 14. Sterile disposable surgical blade (Miltex). 15. Stainless steel tissue tweezers (Aktion TWE-6). 16. Tissue forceps stabilizing pin (Stille). 17. Storage and transport sample container (FıratMed). 18. Heparin tubes (BD). 19. 75 cm2 cell culture flask (Sarstedt). 20. Centrifuge (Heraeus-Labofuge-400R). 21. Gloves. 22. Petri dishes. 2.3 Chemicals and Solutions

1. Distilled water. 2. RPMI-1640 (HyClone, AF29498412). 3. Dulbecco’s Modified Eagle Medium (DMEM) (BIOSERA MS00FG100E). 4. Fetal bovine serum (FBS) (HyClone-RE00000006). 5. Penicillin-Streptomycin (HyClone). 6. Amphotericin B (Sigma A2942). 7. Ficoll-Paque (Biosera-MS00ET100J, Lymphocyte Separation Media). 8. Phosphate buffered saline (PBS) (HyClone-AE29431651). 9. Hanks’ balanced salt solution (HBSS) (Hyclone). 10. Trypsin 0.25%/EDTA (HyClone).

3

Methods

3.1 Preparation of DMEM Medium Solution

1. Add 50 mL FBS into 450 mL L-glutamine-enriched DMEM.

3.1.1 Human Umbilical Cord Blood Derived– Mesenchymal Stem Cell Procedures

1. As a source of mesenchymal stem cells, human cord blood was taken in heparin tubes containing anticoagulant.

Obtaining of Human Umbilical Cord Blood

2. Add 5 mL penicillin + streptomycin into the medium. 3. Store FBS and antibiotic-included DMEM at +4 °C (see Note 2).

2. Transfer the heparin tubes immediately to the laboratory. 3. Mononuclear cells were isolated in the cell culture laboratory within about 3 h.

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Isolation of Human Umbilical Cord Blood– Derived Mesenchymal Stem Cells

1. Wear gloves while handling all procedures. 2. Before the procedure, turn on the UV lights of the biosafety cabinet and laboratory for 45 min at least. 3. Clean the equipment and work surfaces with ethanol. 4. Mix HBSS solution and blood samples into 50 mL falcon tubes at a ratio of 1:1. 5. Add 5 cc Ficoll-Paque solution into 15 mL falcon tubes. 6. Slowly and carefully add the mixture of HBSS and cord blood to the 15 mL falcon tubes with Ficoll-Paque. 7. Centrifuge the mixture loaded onto Ficoll-Paque at 2600 rpm for 20 min with the centrifuge brake closed. 8. After centrifugation, collect buffy coat and transfer to the new sterile 15 mL falcon tubes. 9. Fill the buffy coat-added falcon tubes with PBS until it is 15 mL. Centrifuge buffy coat-PBS mixture at 1500 rpm for 10 min. 10. After centrifugation, discard the supernatant. 11. Add 5 mL HBSS on the pellet and centrifuge at 1000 rpm for 5 min. 12. After centrifugation, discard the supernatant. 13. Add 1 mL DMEM and mix cells and DMEM. 14. Label 75 cm2 culture flasks and precoat the flasks with FBS to remove the monocytic cells 15. Transfer cells and DMEM mixture into the labeled flasks carefully. 16. Add 9 mL DMEM into the flasks and incubate them in the culture incubator. 17. After 10 h, discard 10 mL DMEM.

the

DMEM

mixture

and

add

18. Incubate the cells at 37 °C in a humidified atmosphere with 5% CO2 and change the medium when required. 19. It is highly recommended to use the 3rd or 4th passage for MSC characterization (Fig. 1). 3.1.2 Human Umbilical Wharton’s Jelly Derived– Mesenchymal Stem Cell Procedures Preparation of Sterilization Solution for Human Umbilical Cord

1. Add 25 mL DMEM into 50 mL falcon tube. 2. After DMEM, add 250 μL penicillin and 250 μL amphotericin B and pipette carefully. 3. Incubate the solution in the incubator.

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Fig. 1 Cord blood samples in the heparin tubes are placed into 50 mL falcon tube and mixed with HBSS solution at 1:1 ratio (a). 5 cc Ficoll-Paque is added into the 15 mL falcon tubes (b). The mixture of HBSS and blood samples is loaded carefully onto the 15 mL falcon tubes with Ficoll-Paque (c). Samples are centrifuged at 2600 rpm for 20 min with centrifuge brake closed (d–f). After centrifugation, buffy coat is collected in new sterile falcon and washed twice (g, h). The supernatant is discarded and 1 mL DMEM is added to the cells, and finally the mixture of cells and DMEM is transferred to a 75 cm2 labeled flask (i) Obtaining of Human Umbilical Cord

1. As a source of mesenchymal stem cells, approximately 15–20 cm of human umbilical cord was taken into storage sample container. 2. Transfer the sample containers immediately to the laboratory. 3. Divide into two pieces and place them into to prepared sterilization solution falcon tube before. 4. Incubate the falcon tubes in the incubator for one and half hour (Fig. 2).

Isolation of Human Umbilical Wharton’s Jelly– Derived Mesenchymal Stem Cells

1. After incubation, sterilize the surface of the falcon tube and place into the biosafety cabinet. 2. Place the piece of cord in a petri dish and add 1–2 mL PBS. 3. Remove arterial and vein vessels from the cord using scissors and forceps. 4. After removing the veins, cut the cord into shreds using scissors and a surgical blade. 5. Place the Wharton’s jelly pieces into labeled 75 cm2 flasks at regular intervals with the help of tweezers. 6. Incubate the cells at 37 °C in a humidified atmosphere with 5% CO2 and change the medium when required. 7. It is highly recommended to use the 3rd or 4th passage for MSC characterization (Fig. 3).

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Fig. 2 The cord sample in the storage container (a) is placed into prepared sterilization solution and incubated for one and half hour (b, c)

Fig. 3 The cord sample is placed into a petri dish (a). The veins are removed away from the tissue and shred using scissors and forceps (b, c). Wharton’s jelly pieces are placed into the flask (d)

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Notes 1. Our current study was approved by the Ethics Committee of Istanbul University, Istanbul Faculty of Medicine, on 27 May 2020 (No: 62261). The human umbilical cord blood samples used in the study were collected from individuals who had healthy pregnancies and were participants in the study with informed consent at the Obstetrics and Gynaecology Polyclinic of Istanbul Bakırkoy Dr Sadi Konuk Training and Research Hospital. This study was funded by a grant from Istanbul University Scientific Research Projects Unit (BAP project ID no: 37059) and Council of Higher Education of Turkish Republic 100/2000 PhD Program. 2. The prepared DMEM solution is used for culture cells when needed.

References 1. Friedenstein AJ, Deriglasova UF, Kulagina NN et al (1974) Precursors for fibroblasts in different populations of hematopoietic cells as detected by the in vitro colony assay method. Exp Hematol 2(2):83–92 2. Markov A, Thangavelu L, Aravindhan S et al (2021) Mesenchymal stem/stromal cells as a valuable source for the treatment of immunemediated disorders. Stem Cell Res Ther 12(1): 192 3. Zhu R, Yan T, Feng Y et al (2021) Mesenchymal stem cell treatment improves outcome of COVID-19 patients via multiple immunomodulatory mechanisms. Cell Res 31(12): 1244–1262 4. Lanzoni G, Linetsky E, Correa D et al (2021) Umbilical cord mesenchymal stem cells for COVID-19 acute respiratory distress syndrome: a double-blind, phase 1/2a, randomized controlled trial. Stem Cells Transl Med 10(5):660–673 5. Xu F, Fei Z, Dai H et al (2022) Mesenchymal stem cell-derived extracellular vesicles with high PD-L1 expression for autoimmune diseases treatment. Adv Mater 34(1):e2106265 6. Riazifar M, Mohammadi MR, Pone EJ et al (2019) Stem cell-derived exosomes as nanotherapeutics for autoimmune and neurodegenerative disorders. ACS Nano 13(6): 6670–6688 7. Rautiainen S, Laaksonen T, Koivuniemi R (2021) Angiogenic effects and crosstalk of adipose-derived mesenchymal stem/stromal cells and their extracellular vesicles with endothelial cells. Int J Mol Sci 22(19):10890

8. Aydin S, S¸ahin F (2019) Stem cells derived from dental tissues. Adv Exp Med Biol 1144: 123–132 9. Kim HJ, Cho KR, Jang H et al (2021) Intracerebroventricular injection of human umbilical cord blood mesenchymal stem cells in patients with Alzheimer’s disease dementia: a phase I clinical trial. Alzheimers Res Ther 13(1):154 10. Ranjbaran H, Abediankenari S, Mohammadi M et al (2018) Wharton’s jelly derivedmesenchymal stem cells: isolation and characterization. Acta Med Iran 56(1):28–33 11. Dominici M, Le Blanc K, Mueller I et al (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315–317 12. Atashi F, Modarressi A, Pepper MS (2015) The role of reactive oxygen species in mesenchymal stem cell adipogenic and osteogenic differentiation: a review. Stem Cells Dev 24(10): 1150–1163 13. Li M, Yin H, Yan Z et al (2022) The immune microenvironment in cartilage injury and repair. Acta Biomater 140:23–42 14. Chen Q, Shou P, Zheng C et al (2016) Fate decision of mesenchymal stem cells: adipocytes or osteoblasts? Cell Death Differ 23(7): 1128–1139 15. Chen Y, Shen H, Ding Y et al (2021) The application of umbilical cord-derived MSCs in cardiovascular diseases. J Cell Mol Med 25(17):8103–8114

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16. Marrelli M, Paduano F, Tatullo M (2015) Human periapical cyst-mesenchymal stem cells differentiate into neuronal cells. J Dent Res 94(6):843–852 17. Samaeekia R, Rabiee B, Putra I et al (2018) Effect of human corneal mesenchymal stromal cell-derived exosomes on corneal epithelial wound healing. Invest Ophthalmol Vis Sci 59(12):5194–5200 18. Cruz FF, Rocco PRM (2020) The potential of mesenchymal stem cell therapy for chronic lung disease. Expert Rev Respir Med 14(1): 31–39 19. Marofi F, Alexandrovna KI, Margiana R et al (2021) MSCs and their exosomes: a rapidly evolving approach in the context of cutaneous wounds therapy. Stem Cell Res Ther 12(1):597 20. Dong L, Wang Y, Zheng T et al (2021) Hypoxic hUCMSC-derived extracellular vesicles attenuate allergic airway inflammation and airway remodeling in chronic asthma mice. Stem Cell Res Ther 12(1):4 21. Hmadcha A, Martin-Montalvo A, Gauthier BR, Soria B, Capilla-Gonzalez V (2020) Therapeutic potential of mesenchymal stem cells for cancer therapy. Front Bioeng Biotechnol 8:43

22. Niess H, Thomas MN, Schiergens TS et al (2016) Genetic engineering of mesenchymal stromal cells for cancer therapy: turning partners in crime into Trojan horses. Innov Surg Sci 1(1):19–32 23. Zhu Y, Sun Z, Han Q et al (2009) Human mesenchymal stem cells inhibit cancer cell proliferation by secreting DKK-1. Leukemia 23(5):925–933 24. Vianello F, Villanova F, Tisato V et al (2010) Bone marrow mesenchymal stromal cells non-selectively protect chronic myeloid leukemia cells from imatinib-induced apoptosis via the CXCR4/CXCL12 axis. Haematologica 95(7):1081–1089 25. Yu K, Yin Y, Ma D et al (2020) Shp2 activation in bone marrow microenvironment mediates the drug resistance of B-cell acute lymphoblastic leukemia through enhancing the role of VCAM-1/VLA-4. Int Immunopharmacol 80: 106008 26. Sel FA, Oguz FS (2022) Regenerative medicine application of mesenchymal stem cells. In: Turksen K (ed) Cell Biology and Translational Medicine, Volume 16. Advances in Experimental Medicine and Biology, vol 1387. Springer, Cham

Methods in Molecular Biology (2023) 2736: 85–93 DOI 10.1007/7651_2023_484 © Springer Science+Business Media, LLC 2023 Published online: 24 May 2023

Encapsulation of MSCs in PRP-Derived Fibrin Microbeads € O¨zge Lalegu¨l-Ulker, S¸u¨kran S¸eker, Ays¸e Eser Elc¸in, and Yas¸ar Murat Elc¸in Abstract Platelet-rich plasma (PRP) is a highly concentrated platelet-containing blood plasma that incorporates a significant amount of growth factors and cytokines needed to accelerate the tissue repair process. PRP has been used effectively for many years in the treatment of various wounds by direct injection into the target tissue or impregnation with scaffold or graft materials. Since autologous PRP can be obtained by simple centrifugation, it is an attractive and inexpensive product for use in repairing damaged soft tissues. Cellbased regenerative approaches, which draw attention in the treatment of tissue and organ injuries, are based on the principle of delivering stem cells to damaged sites by various means, including encapsulation. Current biopolymers used in cell encapsulation have some advantages with some limitations. By adjusting its physicochemical properties, PRP-derived fibrin can become an efficient matrix material for encapsulating stem cells. This chapter covers the fabrication protocol of PRP-derived fibrin microbeads and their use to encapsulate stem cells as a general bioengineering platform for prospective regenerative medical applications. Key words Blood-derived materials, Cell encapsulation, Fibrin, Mesenchymal stem cells, Personalized medicine, Platelet-rich plasma, Regenerative medicine, Tissue engineering

1

Introduction Platelet-rich plasma (PRP) is a concentrated platelet suspension obtained by centrifugation of whole blood, containing significant amounts of growth factors and cytokines responsible for wound healing. Platelets, also called thrombocytes, are non-nucleated cells derived from megakaryocytes, the largest hematopoietic cells of the bone marrow. They have a characteristic discoid shape in a size range from 1 to 3 μm [1]. The main functions of platelets are to prevent acute blood loss by repairing damaged blood vessel walls and adjacent tissues after injury. Platelets circulating in the bloodstream in an inactive state become active when they come into contact with collagen following endothelial damage to perform their physiological functions. Platelets contain a number of bioactive molecules within the cytoplasmic granules [2]. After activation, platelets’ shape changes and aggregation typically occurs, followed by the release of growth factors in the α-granule content [3]. Some of these factors include platelet-derived growth factor (PDGF), transforming growth factor-beta (TGF-β), vascular endothelial 85

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growth factor (VEGF), epithelial growth factor (EGF) and insulinlike growth factor (IGF-I) [4]. Autologous PRP was first introduced in the 1970s and was initially used as a transfusion product to treat patients with thrombocytopenia [5]. Since that time, autologous PRP has become an attractive blood-derived product for a wide spectrum of medical fields, including dentistry, maxillofacial surgery, sports medicine, orthopedics, plastic and reconstructive surgery, dermatology, ophthalmology, and urology. PRP is traditionally prepared by two-stage centrifugation of an anticoagulated blood sample (citrated, EDTA, or heparinized blood). The whole blood is first centrifuged at low speed to remove erythrocytes. Then, the supernatant is centrifuged at high speed in order to discard a part of the acellular plasma that is called plateletpoor plasma (PPP). After second centrifugation, the platelets (pellet) obtained are suspended in a small amount of blood plasma to obtain mainly a platelet concentrate [6]. The resulting mixture is the PRP that contains a high concentration of platelet-derived growth factors and healing proteins [7]. This is a very simple and low-cost method for preparing a concentrated platelet suspension. A healthy person has ~150,000-350,000/μL of platelets in their blood; that is, at least 1,000,000 platelets can be obtained from 5 mL of PRP. The concentration of human platelets in PRP is several times higher than in whole blood [8]. Besides growth factors, PRP also contains various clotting factors, adhesion molecules, cytokines, chemokines, integrins, and various plasma proteins [9]. Of these plasma proteins, fibrin and fibronectin are cell adhesion molecules, and vitronectin promotes cell migration. These proteins and biologically active molecules in PRP act as messengers and regulators influencing a variety of cell-cell and cell-extracellular matrix interactions [10]. In recent years, stem cell therapy has gained momentum in the field of tissue regeneration and is widely used in wound healing. Multipotent stromal cells, also called mesenchymal stem cells (MSCs), have the ability to differentiate into several mesenchymal cell lineages, such as muscle, cartilage, bone, tendon, and fat [11]. MSCs can stimulate tissue regeneration in many types of injuries, but are not always effective for complete healing [12]. For example, direct injection of MSCs into the wound site may not provide an effective treatment due to the very low cell survival rate and insufficient density of cellular localization. It can be predicted that encapsulation systems [13] that localize stem cells in a specific microenvironment and control the delivery of bioactive molecules secreted by cells will be useful in overcoming this limitation. PRP can be considered as a suitable candidate for the creation of an encapsulation microenvironment, as a bioactive product containing high amounts of growth factors involved in tissue regeneration. The combined use of MSCs and PRP has recently been shown to be an effective approach to repair injured skeletal muscles

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[14]. This chapter describes the fabrication of PRP-derived fibrin microbeads and MSC encapsulation, which can be considered as a platform for encapsulating desired types of cells through a plateletderived growth factor secreting matrix. With its prominent properties, PRP-derived fibrin microbeads have great potential in tissue engineering and regenerative medicine, especially in the context of personal medicinal applications.

2

Materials

2.1 Equipment and Consumables

1. Centrifuge. 2. Axial rotator. 3. Magnetic stirrer. 4. Inverted microscope. 5. Automated cell counter. 6. Carbon dioxide incubator. 7. Laminar flow cabinet. 8. Microplate reader. 9. Pipette pump. 10. Tissue culture polystyrene (TCPS) flasks. 11. Centrifuge tubes. 12. Syringe with 21 G needle tips.

2.2

Solutions

The chemicals used in the encapsulation process were purchased from Sigma-Aldrich (St. Louis, MO); cell culture chemicals were obtained from Lonza (Basel, Switzerland). 1. Bone marrow mesenchymal stem cell growth medium (Solution 1): α-MEM + 10% fetal bovine serum (FBS) + 2 mM L-glutamine + % 1 penicillin/streptomycin solution (Pen/Strep) (see Note 1). 2. Trypsin/ethylenediaminetetraacetic acid (EDTA) solution (Solution 2): 0.05% trypsin and 1.0 mM EDTA in PBS pH 7.4 (see Note 1). 3. Cell suspension buffer (Solution 3): 140 mM NaCl, 5 mM KCl, 10 mM HEPES, 10 mM glucose, and 0.02% EDTA (see Note 2). 4. Crosslinking solution (Solution 4): 0.9 M calcium chloride (in 0.85% NaCl), 10 U/mL thrombin (see Note 2). 5. Alginate removal solution (Solution 5): 55 mM sodium citrate, 150 mM NaCl, and 30 mM EDTA (pH 6.8) (see Note 2). 6. Fixative solution for SEM analysis (Solution 6): 2.5% glutaraldehyde solution in 0.1 M PBS (pH 7.4).

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Methods

3.1 Preparation of PRP

1. Transfer whole blood samples to centrifuge tubes containing anticoagulant (sodium citrate 3.8%) and centrifuge at 1800 rpm for 10 min (see Note 1). 2. Carefully separate the blood plasma (supernatant) from the red blood cells (bottom layer) and transfer to another centrifuge tube and centrifuge at 3600 rpm for 10 min. 3. Gently discard the upper two-thirds portion of the plasma and separate the lower one-third into another tube to obtain PRP. 4. Store the PRP at room temperature by agitating on the axial rotator for fresh use.

3.2 Culture of BMMSCs

1. Seed BM-MSCs into cell culture flasks containing the growth medium (Solution 1) (see Note 1). 2. Incubate cells inside the carbon dioxide incubator at 37 °C, 5% CO2, 95% air, and 90% humidity. 3. When cells reach approximately 70% confluence, passage them using trypsin/EDTA (Solution 2) by following the steps below: (a) Aspirate the growth medium from the flasks. (b) Gently wash the cultured cells twice with sterile 10 mL of 1 × PBS to remove the residual medium. (c) Add the solution containing 0.05% trypsin and 1.0 mM EDTA to the flasks and keep inside the incubator at 37 °C for 2–3 min (see Note 3). (d) Add 10 mL of serum-containing medium to inactivate trypsin. (e) Pipette the medium to collect the detached cells and transfer the cell suspension into a 50 mL conical tube and centrifuge at 250 × g for 5 min. (f) Aspirate the supernatant and resuspend the cells by adding 1 mL of the growth medium (Solution 1). (g) Pipette the resuspended cells and seed into a flask to continue passages. 4. Use the BM-MSCs in the second or third passage for encapsulation after confirming their MSC character (see Note 4).

3.3 Cell Microencapsulation Within PRP-Derived Fibrin

A schematic representation of the encapsulation steps of MSCs in PRP-derived fibrin microbeads is given in Fig. 1. 1. Add 0.2 g of sodium alginate to 10 mL of PRP obtained as described in Subheading 3.1 to attain 2% by weight PRP-alginate solution; then stir on a magnetic stirrer until dissolved (see Note 5).

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Fig. 1 Schematic representation of the encapsulation steps of MSCs in PRP-derived fibrin microbeads

2. Meanwhile, trypsinize the cultured cells up to Passage 2 or 3 following the steps in Subheading 3.2, steps 3(a–e). 3. Next, aspirate the supernatant and resuspend the cells in 100 μL of cell suspension buffer (Solution 3) (see Note 6). 4. Add 100 μL of cell suspension to 3,900 μL of PRP-alginate solution (as described in Subheading 3.3, step 1). 5. Gently pipette the BM-MSCs/PRP-alginate pre-gel mixture until homogeneous and then carefully transfer the suspension into the sterile cartridge of a closed encapsulation device (Nisco, Zurich, Switzerland) (see Note 7). 6. By turning on the droplet-forming device, transfer the cellcontaining suspension to the crosslinking solution (Solution 4) through an electromagnetically driven vibrating 200 μm nozzle via a syringe pump. Then, stir for 15 min to complete the thrombin-mediated ionotropic gelation process (see Note 8). 7. Examine the morphology of the resulting cell-containing fibrin-alginate microbeads (size, uniformity, and homogeneous distribution of cells) under an inverted microscope (Fig. 2a).

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Fig. 2 (a) Phase-contrast micrograph of MSC-encapsulated fibrin microbeads with uniform sphericity. SEM images showing the sphericity (a) and surface topography (b) of a microbead

8. Filter the resulting fibrin-alginate microbeads through a mesh strainer and wash with cell suspension buffer (Solution 3). 9. Next, keep the fibrin-alginate microbeads in the alginate removal solution (Solution 5) for 15 min to obtain alginatefree fibrin microbeads. 10. Wash the fibrin microbeads with the growth medium and culture the encapsulated cells for subsequent use. 3.4 Scanning Electron Microscopy (SEM)

1. Fix MSC-encapsulated fibrin microbeads in fixative solution overnight (Solution 6). 2. Wash microbeads using distilled water to remove excess fixatives. 3. Next, dehydrate the microbeads by passing them through a series of ethanol solutions (50–95%). 4. Sputter-coat the samples with a thin layer of gold-palladium after mounting on stubs, and acquire images by a SEM device operating at 20 kV (Fig. 2b, c).

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1. Maintain the culture of encapsulated BM-MSCs in growth medium in a multi-well plate (6-, 12-, or 24-well) suitable for your processing capacity for a predetermined time interval (e.g., 7, 14, 21 days) (see Note 1). 2. At time points, remove the culture medium from the wells and wash the microbeads with a serum-free medium. 3. Add ((3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide)) (MTT) solution to each well, diluting the culture medium 1:10 (v/v) and incubate at 37 °C for 4 h (see Note 9). 4. After 3–4 h of incubation, observe the formation of insoluble blue-purple formazan crystals with an inverted microscope (see Note 10). 5. Separate the formed formazan crystals and dissolve in 200 μL of MTT solvent. Then measure the absorbance values for each well at a wavelength of 570 nm using a microplate reader (see Note 11).

4

Notes 1. Handling of cell cultures should be performed under a laminar flow cabinet. It is essential to sterilize all materials used for cell culture to avoid any contamination. 2. Solutions prepared under nonsterile conditions such as cell suspension buffer and alginate removal buffer should be sterilized prior to use by filtering through 0.22 μm syringe filter inside laminar flow hood. 3. The trypsin-EDTA solution should be stored at -20 °C and given sufficient time to reach the appropriate temperature before use. 4. Before encapsulation studies, the cell type should be verified by flow-cytometry. For BM-MSCs, CD29+, CD44+, CD90+, CD34–, and CD45– immunophenotype should be demonstrated. 5. Alginate powder should be sterilized under a UV light source (254 nm) for 30 min before being used in cell encapsulation studies. 6. Count cells using a hemocytometer and resuspend the required amount for cell encapsulation experiments. We optimized the cell density as 1 × 106 cells per mL of PRP-alginate solution, but reoptimization of cell density may be required when working with different cell types. 7. It is essential to avoid the formation of bubbles and obtain a homogeneous mixture. If a product size >1,000 μm is not a

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disadvantage for your application, it is also possible to create microbeads manually with the aid of an injector under sterile conditions. 8. Fibrin is obtained by thrombin and calcium-mediated activation of fibrinogen in blood plasma. When using a 200 μm nozzle, the final size range of the microbeads will be ~450–500 μm. 9. The ratio of MTT solution varies according to the well size used. It is appropriate to add 50 μL of MTT solution into 450 μL of culture medium for each well of a 24-well plate. 10. Yellow colored MTT salt reduced to blue-purple colored formazan crystals by mitochondrial dehydrogenase activity of living cells. 11. MTT solvent is generally included in the MTT test kit. However, MTT solvent can also be obtained by preparing a 0.1 N HCl solution in isopropanol.

Competing Interests The authors have intellectual properties related to PRP-derived fibrin biomaterials. Y.M.E. is the founder and director of Biovalda, Inc. (Ankara, Turkey). References 1. Scully D, Naseem KM, Matsakas A (2018) Platelet biology in regenerative medicine of skeletal muscle. Acta Physiol 223(3):e13071 2. Phadke A, Singh B, Bakti N (2019) Role of platelet rich plasma in rotator cuff tendinopathy-clinical application and review of literature. J Clin Orthop Trauma 10(2): 244–247 3. Bakogiannis C, Sachse M, Stamatelopoulos K, Stellos K (2019) Platelet-derived chemokines in inflammation and atherosclerosis. Cytokine 122:154157 4. Chicharro-Alca´ntara D, Rubio-Zaragoza M, Damia´-Gime´nez E, Carrillo-Poveda JM, Cuervo-Serrato B, Pela´ez-Gorrea P, SopenaJuncosa JJ (2018) Platelet rich plasma: new insights for cutaneous wound healing management. J Funct Biomater 9(1):10 5. Gupta M, Barman KD, Sarkar R (2021) A comparative study of microneedling alone versus along with platelet-rich plasma in acne scars. J Cutan Aesthet Surg 14(1):64–71 6. Zhang W, Guo Y, Kuss M, Shi W, Aldrich AL, Untrauer J, Kielian T, Duan B (2019) Platelet-

rich plasma for the treatment of tissue infection: preparation and clinical evaluation. Tissue Eng Part B Rev 25(3):225–236 7. S¸eker S¸, Elc¸in AE, Elc¸in YM (2020) Macroporous elastic cryogels based on platelet lysate and oxidized dextran as tissue engineering scaffold: in vitro and in vivo evaluations. Mater Sci Eng C 110:110703 8. Pavlovic V, Ciric M, Jovanovic V, Stojanovic P (2016) Platelet rich plasma: a short overview of certain bioactive components. Open Med (Wars) 11(1):242–247 9. S¸eker S¸, Elc¸in AE, Elc¸in YM (2019) Autologous protein-based scaffold composed of platelet lysate and aminated hyaluronic acid. J Mater Sci Mater Med 30(12):127 ˆ C, Santana 10. Rodrigues AA, Lana JF, Luzo A MH, Perez AG, Lima-Silva DB, Belangero WD (2014) Platelet-rich plasma and tissue engineering. In Platelet-Rich Plasma, pp. 139. Lana JFSD, Santana MHA, Belangero WD, Luzo ACM (eds), Springer, Berlin 11. Elc¸in YM (2004) Stem cells and tissue engineering. Adv Exp Med Biol 553:301–316

MSC Encapsulation in Fibrin Microbeads 12. Sagaradze GD, Basalova NA, Efimenko AY, Tkachuk VA (2020) Mesenchymal stromal cells as critical contributors to tissue regeneration. Front Cell Dev Biol 8:576176 13. Durkut S, Elc¸in AE, Elc¸in YM (2015) In vitro evaluation of encapsulated primary rat hepatocytes pre- and post-cryopreservation at-80 degrees C and in liquid nitrogen. Artif Cells Nanomed Biotechnol 43(1):50–61

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€ ¨ , S¸eker S¸, Elc¸in AE, Elc¸in YM 14. Lalegu¨l-Ulker O (2019) Encapsulation of bone marrow-MSCs in PRP-derived fibrin microbeads and preliminary evaluation in a volumetric muscle loss injury rat model: modular muscle tissue engineering. Artif Cells Nanomed Biotechnol 47(1):10–21

Methods in Molecular Biology (2023) 2736: 95–103 DOI 10.1007/7651_2023_481 © Springer Science+Business Media, LLC 2023 Published online: 08 June 2023

Assessing Neuronogenic Versus Astrogenic Bias of Neural Stem Cells Via In Vitro Clonal Assay Laura Rigoldi and Antonello Mallamaci Abstract Within the developing cerebral cortex, neural stem cells (NSCs) give rise to neurons and glial cells, according to complex spatio-temporal trajectories. In this respect, a key issue is how NSCs are committed to different neural lineages in time and space. Clonal assays are a powerful tool to address this issue. Here we describe an easy clonal assay protocol employable to dissect NSCs lineage commitment and molecular mechanisms underlying it. NSCs of distinctive spatio-temporal origin, and/or having undergone different molecular manipulations, are plated at low density and allowed to differentiate for a few days. Then, systematic immunoprofiling of the resulting clones allows to quantify commitment of their NSC ancestors to neuronal and astroglial fates. Key words Astrogenesis, Clonal, Commitment, Fate choice, Lineage, Neurogenesis, NSC

1

Introduction Within the developing cerebral cortex, neural stem cells (NSCs) give rise to neurons and astrocytes according to specific schedules, which depend on the region taken into account, the developmental stage, and the genotype [1]. A key parameter affecting the final outcome of the process is the rate at which NSCs progress toward lineage-committed progenitors, neuronal as well as astroglial (aka neuronoblasts and astroblasts) [2]. This parameter can be evaluated in vivo as well as in vitro by clonal assays. In these assays, clones originating from neural precursors are categorized into neuronal, mixed, and astroglial, and changes of the corresponding frequencies are used as an index of altered NSC commitment to distinctive neural fates. Albeit preferable as run in a physiological milieu, in vivo clonal assays classically require a huge number of animals, due to the relatively low number of geometrically isolated clones identifiable in each brain. More recent use of tagged libraries fixes this issue; however, it remarkably increases the complexity of immunohistological data collection and analysis [3]. Despite limits of reductionistic approaches, performing clonal analysis in vitro

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circumvents these issues, allowing to get a rapid and cheap insight into the impact that a given factor exerts on NSCs’ histogenetic choices. In a typical in vitro clonal assay, NSCs may be prompted to selfrenew for a convenient time as floating neurospheres, in a growth factor (GF)-supplemented medium, where, if due, they are differentially exposed to the factor of interest (e.g., the activation of a transgene) or its control. Next, NSCs and committed progenitors originating from dissociation of these neurospheres are transferred to a pro-differentiative, serum-supplemented medium and let to attach as isolated cells to a matrix substrate. Finally, isolated clones originating from these cells are immunoprofiled, and frequencies of mixed, neuronal, and astroglial clones (originating from NSCs, neuronoblasts, and astroblasts, respectively) are used to infer the histogenetic bias of NSCs [4]. While running in vitro clonal assays, a critical parameter requiring careful consideration is the superficial density at which isolated, NSC-derivative precursors are plated and “interrogated” for their histogenetic properties. At one side of the spectrum, the use of super-low densities (e.g., one cell per Terasaki plate well) [5] guarantees the geometrical segregation among clones. However, it fully abrogates community effects occurring in vivo, and it may result logistically and economically demanding. On the contrary, operating at very high densities may mitigate the deficit in community effects as well as logistical and financial issues. However, within the timeframe allocated to neural differentiation, this approach may lead to substantial inter-clonal coalescence, which artifactually reduces the prevalence of pure clones, compresses their dynamics, and, therefore, jeopardizes the sensitivity of the assay [2]. In this respect, plating differentiating precursors at intermediate densities (e.g., 8000 cells/cm2) and immunoprofiling their derivatives four days after can be an acceptable compromise, obviously provided that the two or more batches of NSCs interrogated do not display any differential bias to clonal coalescence [2]. In such a case, using very low plating densities would be mandatory. Based on these considerations, we recommend to preliminary assess (see Note 1) the congruence between coalescence rates peculiar to distinct NSCs subject of examination, and, only after that, to execute the clonal assay procedure detailed below.

2 2.1

Materials Reagents

2.1.1 Media Proliferative

DMEM/F-12, GlutaMAX (Gibco#31331028) supplemented with 1× N2 Supplement (Gibco 17502048), 1 mg/mL BSA, 0.6% w/v glucose, 2 μg/mL heparin (Sigma H3393), 20 ng/mL bFGF (Invitrogen #PHG0261), 20 ng/mL EGF (Invitrogen #PHG0311), 1× Pen/Strept (Invitrogen #15140122), 10 pg/mL fungizone (Invitrogen #15290026).

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Differentiative

Neurobasal A (Gibco #10888022) supplemented with 1× B27 (Gibco #17504044), 5% of fetal bovine serum (FBS, Euroclone #ECS0180L), 1× GlutaMAX (Gibco #35050038), 1× Pen/Strept (Invitrogen #15140122), 10 pg/mL fungizone (Invitrogen #1529 0026).

2.1.2 Blocking Solution

1× PBS; 10% fetal bovine serum (FBS), 1 mg/mL BSA; 0.1% Triton X-100.

2.1.3 Antibodies

The following primary antibodies were used: α-Tub-β3, mouse monoclonal (clone Tuj1; Covance MMS-435P), 1:1000. α-GFAP rabbit polyclonal (DAKO#Z0334), 1:400. α-S100β, rabbit monoclonal (Abcam #52642), 1:200. α-EGFP, chicken polyclonal (Abcam #137970), 1:800. α-RFP, mouse monoclonal (Chromotek 5f8-100), 1:500. The following secondary antibodies were used: Alexa 488 Goat α-Rabbit (Invitrogen), 1:500. Alexa 594 Goat α-Mouse (Invitrogen), 1:500. Alexa 488 Goat α-Chicken (Invitrogen), 1:800. Alexa 594 Donkey α-Rat (Invitrogen), 1:500.

2.1.4 Other

DNase I (Roche #10104159001). Trypsin-EDTA (0.5%), no phenol red (Gibco #15400054). Soybean Trypsin Inhibitor, powder (Gibco #17075029). Poly-D-lysine (Sigma #P7405). Leukemia Inhibitory Factor (Sigma #ESG1106). Vectashield Mounting Medium (Vector #H-1000).

2.2

Equipment

1. Stereomicroscope. 2. Biological hood. 3. Scissors. 4. Tweezers. 5. 1× phosphate-buffered saline. 6. 30% glucose. 7. Ice. 8. p2, p20, p200, p1000 pipettes. 9. p2, p20, p200, p1000 tips. 10. 1.5 mL tube. 11. Trypan blue.

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12. 12 mmØ glass coverslip. 13. 24-well plate. 14. 12-well plate. 15. 4% paraformaldehyde. 16. DAPI (4′, 6′-diamidino-2-phenylindole). 17. Microscope slide. 18. Epifluorescence microscope.

3

Methods The following steps are performed under non-aseptic conditions.

3.1 Embryonic Dissection

1. Sacrifice a pregnant mouse at 11 dpc (day post-coitum) (see Note 2) by cervical dislocation. 2. Spray briefly the mouse womb with 70% (v/v) and dissect out the uterine horns. Gently collect them into a petri dish containing sterile 1× PBS + 0.6% (w/v) glucose. The following steps are carried out in a tissue culture hood under aseptic conditions and using sterile tools. All the reagents used in this protocol are optimized for 12–13 embryos; if your conditions are different, the results can vary. 3. Under a stereomicroscope, separate each embryo from its placenta and embryonic sac. 4. Dissect the cortex (see Note 3) of each embryo in sterile 1× PBS + 0.6% (w/v) glucose and transfer all the cortices (from 12 embryos circa) to a 1.5 mL Eppendorf, kept on ice.

3.2 Setting Up of the Neurosphere Culture

1. Remove the excess of PBS/glucose solution, paying attention not to touch the cortices, and add 300 μL of complete proliferative medium. 2. Gently pipette up and down 10 times using a 200 μL pipette, to mechanically dissociate the cortices to single neural precursor cells, and transfer the Eppendorf to ice. 3. Wait 3 min in order to allow chunks of undissociated tissue to get down by gravity. 4. Transfer 200 μL of supernatant containing single neural precursor cells to a 1.5 mL Eppendorf, kept on ice. 5. Add 200 μL of complete proliferative medium to undissociated tissue and repeat steps 2–4, transferring dissociated precursor cells to the 1.5 mL Eppendorf referred to in step 4.

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6. Repeat step 5 2–3 times, and discard the residual undissociated tissue. 7. Count viable dissociated cells by trypan blue exclusion and resuspend them at a density of 500,000 cells/mL. 8. Plate 500 μL of cell suspension at per well of a 24-well plate. 9. Apply the “factor of interest” (e.g., a drug, or a recombinant lentivirus for somatic transgenesis) (see Note 4). 10. Place the plate in an incubator at 37 °C and 5% CO2 for 4 days (see Note 5). 11. The following day, check under the microscope the presence of neurospheres. 3.3

Clonal Assay

1. At day in vitro 4 (DIV 4), collect the medium containing the neurospheres of each well in a 1.5 mL Eppendorf and centrifuge at 180 g for 5 min at room temperature (RT). 2. Discard the supernatant and add 2 μL of 1 mg/mL DNAse I and 200 μL of 0.05% (w/v) trypsin-EDTA to each Eppendorf. 3. Gently pipette few times (approx. 3) and incubate for 5 min at RT. 4. Add 300 μL of trypsin inhibitor solution (see Note 6) to stop trypsin activity. 5. Pipette up and down 15–20 times with p200 pipette to generate a single cell suspension, paying special attention not to make air bubbles and keeping the pipette tip far (≥2 mm) from the Eppendorf bottom (see Note 7). 6. Centrifuge at 180 g for 5 min at RT. 7. Discard the supernatant. Resuspend cells in 1 mL of complete differentiative medium, count viable cells by trypan blue exclusion, and dilute each sample to 80,000 cells/mL. 8. Plate 8000 cells per cm2 on 12 mmØ coverslips coated by polyD-lysine (0.2 mg/mL) (see Note 8) and pre-placed in a 24-well plate (see Note 9). 9. Keep the plate in an incubator at 37 °C and 5% CO2 for 3 more days. 10. At DIV 7, 24 h before blocking the assay, add 1 × 106 U/mL mouse Leukemia Inhibitory Factor (mouse LIF) (see Note 10).

3.4 Clonal Assay Immunocytochemistry

1. 24 h later, at DIV 8, fix cells in 4% paraformaldehyde (PFA) for 15 min at 4 °C. 2. Gently wash cells 3 times in 1× PBS. 3. Incubate cells in blocking solution for at least 1 h at RT.

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4. Incubate the samples with primary antibodies, α-Tub-β3 and α-GFAP (the latter replaceable by α-S100β), pre-diluted in blocking solution as detailed in Subheading 2, overnight at 4 °C. 5. The following day, wash samples for 5 min in 1× PBS – 0.1% Triton X-100. Repeat this step 2 times. 6. Incubate samples with secondary antibodies, pre-diluted in blocking solution as detailed in Materials, for 2 h at RT. 7. Wash the samples for 5 min in 1× PBS. Repeat this step for 2 times. 8. Counterstain cells phenylindole).

with

DAPI

(4′,

6′-diamidino-2-

9. Mount the samples on microscope slides with Vectashield Mounting Medium. 3.5 Clonal Assay Analysis

1. Acquire immunofluorescence images in epifluorescence with a 20× air objective. 2. Randomly choose clones based on only DAPI signal (see Note 11). Here, a clone is defined as a group of at least 2 cells whose reciprocal distance is less than two average cell-soma diameters (Fig. 1). 3. Then, categorize clones (Fig. 2) based on neuronal and astroglial markers (see Note 12): “pure neuronal clones” – clones including Tub-β3 positive cells, but not GFAP/S100β (see Note 13) positive ones; “pure astroglial clones” – clones including GFAP/S100β positive cells, but not Tub-β3 positive ones; “mixed clones” – clones including both Tub-β3 positive cells and GFAP/S100β positive cells. 4. Evaluate clone frequencies and perform statistical analysis of results (see Note 14).

Fig. 1 Example of clones selection. Scalebar 20 μm

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Fig. 2 Clones phenotyping. Scalebar 20 μm

4

Notes 1. To compare coalescence biases peculiar to cells exposed to factor A and control factor, we suggest to run an ad hoc test, as detailed below. First, alternatively transduce two halves of the starting NSCs pool (prepared as in Methods, Subheading 3.2, step 7) by two lentiviral vectors, driving constitutive Egfp or mCherry expression, and make aliquots of each transduced hemi-pool (as described in Methods, Subheading 3.2, step 8). Immediately afterward, apply “factor A” or control factor (e.g., A-transgene vs control-transgene) to both Egfp+ and mCherry+ aliquots (as described in Methods, Subheading 3.2, step 9). Next, further follow the protocol up to Methods, Subheading 3.3, step 7. Then, mix precursors originating from dissociation of DIV4, Egfp+, and mCherry+, neurospheres having been exposed to the same transgene (A or control) at 1:1 ratio, and plate the resulting suspensions at (cumulative) 8000 cells/cm2 in complete differentiative medium (as described in Methods, Subheading 3.3, step 8). Complete the clonal assay as described in this method, however replacing α-Tub-β3 and α-GFAP (and their corresponding secondary antibodies) by α-Egfp and α-mCherry (and their corresponding secondary antibodies). Finally, evaluate the frequency of A-transgene+ and control-transgene+ clones co-including Egfp+ and mCherry+ cells, as an index of clone coalescence. 2. It is also possible to run this assay starting later during embryonic development (e.g., embryonic day 12 to 15). 3. Microdissection of cortices has been performed including only the mouse neonatal pallium (e.g., no signaling centers – septum, hem, and antihem – have been included). In this method,

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we dissect a specific region of the brain (e.g., the cortex), but this assay can be run using other brain regions of interest. 4. In case of lentiviral somatic transgenesis, NCSs are acutely infected with dedicated lentiviral vector (LV) sets at multiplicity of infection (MOI) = 8 per each LV, having the lentiviral titer previously been evaluated on HEK293T reference cells. As previously reported [6] and consistently with our experience (not shown), if cell density ≥ 200 cells/μL, these conditions are sufficient to effectively co-transduce almost the totality of neural cells. Moreover, murine NSCs can tolerate cumulative MOIs up to 30. To note, transduction of murine NSCs by recombinant LVs (preferably generation ≥3 LVs) is performed as described [6], in full compliance with Biological Safety Level 2 (BSL2) rules [7]. 5. Murine neurospheres in proliferation need to be periodically passaged, in order to prevent their exaggerated growth (typically every 3–4 days). In this respect, 2–3 days after cell plating, should neurospheres have become too big, gently transfer them in a 12-well plate, pipetting them 2 times by a p1000 pipette (do not passage them by trypsin, to prevent digestion of growth factors’ receptors). 6. The trypsin inhibitor solution is composed of 5 mL of DMEM/F-12, 5 μL of 1 mg/mL DNAse I, and 70 μL of 10 mg/mL trypsin inhibitor. 7. Before the differentiation step, it is critical to generate a single cell suspension, to be sure that every clone analyzed at the end of the assay has been generated starting from one single neural precursor cell plated. 8. Perform poly-D-lysine coverslip coating as follows. Dilute the 1 mg/mL poly-D-lysine stock 1:5 with sterile H2O to a final concentration of 0.2 mg/mL. Cover the coverslip on the bottom of each well with 400 μL of the diluted solution. Incubate over night at 4 °C. Aspirate the solution, wash 3 times with H2O, and let wells dry before adding the medium. 9. After plating for the differentiation step, check under the microscope to be sure that cells are homogeneously distributed on the well plate. 10. Stimulation with LIF permits to unmask early astroglial committed precursors. If the “factor” applied to NSCs is expected to intrinsically bias NSCs toward astrogenesis, this step may be omitted, in order to increase the resolving power of the assay. In such a case, the subsequent PFA fixation step can be preferably anticipated by one day, i.e., to DIV7 [8]. 11. This step must be run by an operator blind to “antibody signals” and unaware of samples identities.

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12. This step must be run by an operator unaware of samples identities. 13. Pay attention: αGFAP selectively labels murine neopallial astrocytes; however, within the murine archipallium and the primate pallium, it also labels NSCs. As for S100β, its E11.5 + DIV8 signal is weak, so that its detection may result challenging. 14. ≥300 clones (collected from about 40 randomly assorted photographic fields) are preferably scored per each experimental condition and each biological replicate. At least three biological replicates are analyzed per each experimental condition. Statistical significance of results is evaluated by t-test analysis.

Acknowledgments We thank Manuela Santo who contributed to set the protocol. This work was supported by intramural SISSA funding. References 1. Mallamaci A (2013) Developmental control of cortico-cerebral astrogenesis. Int J Dev Biol 57(9–10):689–706 2. Santo M, Rigoldi L, Falcone C, Tuccillo M, ˜ o V, Mallamaci Calabrese M, Martı´nez-Cerden A (2023) Spatial control of astrogenesis progression by cortical arealization genes. Cereb Cortex 33(6):3107–3123 ˜ ate M, 3. Cerrato V, Parmigiani E, Figueres-On Betizeau M, Aprato J, Nanavaty I, Berchialla P, Luzzati F, de Sperati C, Lo´pez-Mascaraque L, Buffo A (2018) Multiple origins and modularity in the spatiotemporal emergence of cerebellar astrocyte heterogeneity. PLoS Biol 16(9): e2005513 4. Falcone C, Santo M, Liuzzi G, Cannizzaro N, Grudina C, Valencic E, Peruzzotti-Jametti L, Pluchino S, Mallamaci A (2019) Foxg1 antagonizes neocortical stem cell progression to astrogenesis. Cereb Cortex 29(12):4903–4918

5. Temple S (1989) Division and differentiation of isolated CNS blast cells in microculture. Nature 340(6233):471–473 6. Brancaccio M, Pivetta C, Granzotto M, Filippis C, Mallamaci A (2010) Emx2 and Foxg1 inhibit gliogenesis and promote neuronogenesis. Stem Cells 28(7):1206–1218 7. Stanford Environmental Health & Safety. Lentivirus Fact Sheet. https://ehs.stanford.edu/refer ence/lentivirus-fact-sheet. Accessed 1 April 2023 8. Frisari S, Santo M, Hosseini A, Manzati M, Giugliano M, Mallamaci A (2022) Multidimensional functional profiling of human neuropathogenic FOXG1 alleles in primary cultures of murine pallial precursors. Int J Mol Sci 23(3):1343

Methods in Molecular Biology (2023) 2736: 105–114 DOI 10.1007/7651_2022_473 © Springer Science+Business Media, LLC 2023 Published online: 08 February 2023

Stem Cell-Based Modeling Protocol for Parkinson’s Disease Babak Arjmand, Shayesteh Kokabi-Hamidpour, Hamid Reza Aghayan, Sepideh Alavi-Moghadam, Rasta Arjmand, Mostafa Rezaei-Tavirani, Parisa Goodarzi, Ensieh Nasli-Esfahani, and Mohsen Nikandish Abstract Parkinson’s disease is a progressive neurodegenerative disorder, which is mainly characterized by unintended or uncontrollable body movements. Pathophysiologically, disturbances in the neurotransmission system of the brain like dopaminergic system and synaptic dysfunction are classified as top-rated causes of the onset of Parkinson’s disease, which symptoms can be different according to the involvement of neurotransmission system type and the effect of the disease on the motor and non-motor systems. Although some pharmacological and non-pharmacological approaches have been applied to control and slow down the progression of the disease, a definitive cure has not yet been discovered. One of the factors involved in this issue is the lack of appropriate laboratory models to investigate the pathological mechanisms involved in the disease as well as various aspects of candidate drugs, which ultimately leads to the failure of drug discovery and development pipelines. To deal with these challenges, the application of stem cells, especially cellular reprogramming of somatic cells to human pluripotent stem cells, also known as induced pluripotent stem cells, has been able to promise a new chapter in the modeling of Parkinson’s disease. Induced pluripotent stem cells have the stemness capability; therefore, they can differentiate into any type of cell such as nerve cells. Also, since these cells are obtained from the reprogramming of somatic cells in the patient’s body, they maintain the patient’s genetic content, which can play an important role in increasing the quality of disease modeling and the validity of the results of laboratory studies. Therefore, the procedure for modeling induced pluripotent stem cells for Parkinson’s disease is explained in this chapter. Key words Disease modeling, Embryoid body, Human-induced pluripotent stem cells, Midbrain dopaminergic neurons, Parkinson’s disease, Stem cell-based models

Abbreviations BDNF bFGF DA DMEM/F-12 EB FBS FGF8a GDNF hiPSCs iPSCs

Brain-derived neurotrophic factor Basic fibroblast growth factor Dopaminergic neurons Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 Embryoid body Fetal bovine serum Fibroblast growth factor-8a Glial-derived neurotrophic factor Human-induced pluripotent stem cells Induced pluripotent stem cells

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KOSR NEAAs PBS PD PNS ROS SN SOP

1

Knockout serum replacement Nonessential amino acids Phosphate-buffered saline Parkinson’s disease Peripheral nervous system Reactive oxygen species Substantia nigra Standard operating procedure

Introduction Parkinson’s disease (PD) refers to a pathophysiological condition, which is triggered by the degeneration of the brain and progress through the demolition of motor and non-motor systems [1-4] [1-4][6-71722] [1–4]. Etiopathogenically, the loss of dopaminergic neurons (DA) in the substantia nigra (SN) region of the brain is among the main initial mechanisms involved in PD motor deficits, which is triggered as a result of the production of reactive oxygen species (ROS) caused by the oxidative stress [5, 6]. Consequently, PD motor deficits can be accompanied by various PD symptoms such as slowed movements, postural instability, rigidity of muscles, and tremors [7]. In addition to the motor system, non-motor system dysfunctionality can also lead to the onset of various symptoms such as orthostatic hypotension, constipation, urogenital disorders, cognitive and behavioral disorders, sleeping problems, and mood symptoms [8], which can be appeared as a result of disturbances in the noradrenergic, serotonergic, and cholinergic systems as well as neurodegeneration of peripheral nervous system (PNS) [7]. It is also worth mentioning that symptoms of PD appear progressively and deteriorate through the years [9]. However, the severity of symptoms varies among patients [10]. Nevertheless, the progression and development of PD can be life-threatening and highlight the need for prompt medical attention [11]. As yet, many pharmacological (e.g., levodopa and selegiline) [12] and non-pharmacological approaches (e.g., occupational therapy and physiotherapy) have been represented for patients, which can relieve symptoms or slow down the progression of PD [13]. Although in recent years, many steps have been taken in the field of drug discovery and development of PD [14], no definitive treatment has been reported for patients that have led to a notable outcome as a definite cure [12]. In addition, some pharmaceutical approaches have been associated with undesirable effects and challenges [15]. Observations indicate that the lack of appropriate laboratory models is strongly related to drug discovery and development failure for patients with PD. Accordingly, the mentioned challenges highlighted the importance of seeking suitable models

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to scrutinize the pathological mechanisms underlying PD and elucidate all aspects of candidate drugs in this field [16]. In this context, stem cell technology has been proposed not only as an effective treatment approach for PD [17, 18] but also as a novel in vitro modeling approach, which can provide a biological platform to achieve in-depth knowledge of PD pathogenesis and pave the way for discovering new drugs with higher therapeutic potential [19]. Induced pluripotent stem cells (iPSCs) are one of the stem cell types, which have attracted the attention of many researchers in the field of PD modeling and treatment due to many advantages [20–23]. For instance, iPSCs are obtained through the reprogramming of somatic cells derived from the patient’s body [24] Accordingly, they can preserve the genetic content of the patient, which is of great value in the field of disease research [25]. In addition, they can differentiate into different cell types due to their stemness capability [26]. Therefore, they can be used to develop biological platforms like organoids in the field of modeling diseases related to the nervous system [24]. Herein, the current protocol described the process of producing embryoid bodies (EB) and subsequently dopaminergic neurons from iPSCs, which can be applied for drug discovery and development of PD.

2

Materials

2.1 Induced Pluripotent Stem Cells Culturing

1. Human-induced pluripotent stem cells (hiPSCs) (Royan Institute, Iran). 2. 70% ethanol (Kimia Alcohol Zanjan, Iran). 3. Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12; Gibco™, 11320033). 4. 50% TrypLE™ Select (Thermo Fisher Scientific, USA) (see Note 1). 5. 2-Mercaptoethanol (50 mM) (Invitrogen, 21985023). 6. Knock out serum replacement (KOSR; Gibco, 10828-028). 7. Nonessential amino acids (NEAAs; Invitrogen, 11140-035). 8. L-glutamine (Invitrogen, 25030-024). 9. Penicillin-streptomycin (Invitrogen, 15140122). 10. Basic fibroblast growth factor (bFGF; Thermo Fisher Scientific). 11. Phosphate-buffered saline (PBS; Biowest, France). 12. iPSC medium: StemFit AK02N (Ajinomoto). 13. ROCK inhibitor Y-27632 (Sigma-Aldrich). 14. iMatrix-511MG (Thermo Fisher Scientific). 15. Trypan blue solution 0.4% (Invitrogen, USA).

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16. LM511-E8-coated dishes. 17. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 18. 6-well plates. 19. Cell Scraper (Falcon, Cat. no. 353085). 20. Filter cap cell culture flasks (300, 175, 75, and 25 cm2 (TPP, Switzerland)). 21. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea). 22. Sterile conical tubes 15 and 50 mL (SPL, Korea). 23. Hemocytometer and cover glass. 24. Microcentrifuge tube (SPL, Korea). 25. Inverted microscope with phase contrast (Nikon, Japan). 26. NucleoCassette™ (Chemometec, Denmark). 27. NucleoCounter® NC-100™ (Chemometec, Denmark). 28. Biological safety cabinet (Esco, Singapore). 2.2 Embryoid Body (EB) Formation

1. Aggrewell plates Technologies).

(10,000

cells

per

EB;

Stem

Cell

2. Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12; Gibco™, 11320033). 3. Knockout serum replacement (KOSR; Gibco, 10828-028). 4. L-glutamine (Invitrogen, 25030-024). 5. Nonessential amino acids (NEAAs; Invitrogen, 11140-035). 6. Penicillin-streptomycin (Invitrogen, 15140122). 7. 2-Mercaptoethanol (50 mM) (Invitrogen, 21985023). 8. ROCK inhibitor Y-27632 (Sigma-Aldrich). 9. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 10. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea). 11. Sterile conical tubes 15 and 50 mL (SPL, Korea). 12. Cell Scraper (Falcon, Cat. no. 353085). 13. 0.1% Gelatin-coated plates (24-well plate of Sigma-Aldrich). 14. 0.1% Gelatin-coated plates (96-well plate of Sigma-Aldrich). 15. Fetal bovine serum (FBS) Biopharm—EDQM certified (Biowest, USA). 2.3 In Vitro Differentiation of iPSCs into Midbrain Dopaminergic Neurons (mDMA)

1. Neural induction medium: 47 mL DMEM/F-12, 1.0 mL 50 B27 Supplement, 0.5 mL GlutaMAX, 0.5 mL 100 N2 Supplement, 0.5 mL P/S, 0.5 mL NEAA, 100 μL of 100 μg/mL Noggin, and 1 μL of 25 mM DM. 2. L-glutamine (Invitrogen, 25030-024).

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3. N-2 Supplement (100×) (Gibco™, 17502048). 4. Glial-derived neurotrophic factor (GDNF, Gibco™). 5. N6, 29-O-dibutyryl adenosine 39, 59 cyclic monophosphate sodium salt (dCAMP, Sigma-Aldrich). 6. Laminin (Thermo Fisher). 7. Antibiotic-Antimycotic (100×) (Gibco™, 15240062). 8. Fibroblast growth factor-8a (FGF8a, 100 ng/mL; R&D Systems). 9. Heparin (Sigma). 10. Ascorbic acid (Sigma). 11. Brain-derived neurotrophic factor (BDNF, Sigma-Aldrich). 12. Corning™ BioCoat™ Poly-D-Lysine/Laminin Multiwell Plate (Fisher Scientific). 13. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea). 14. Refrigerated centrifuge (swing-out rotor with buckets for 50 and 15 mL tubes) (Hettich, Germany). 15. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 16. Filter cap cell culture flasks (300, 175, 75, and 25 cm2 (TPP, Switzerland)).

3 3.1

Methods Cell Culture

1. Maintain the iPSCs on feeder cells layer containing Dulbecco’s modified Eagle’s medium (DMEM)/F-12 supplemented with 20% knockout serum, 0.1 mM 2-mercaptoethanol, 1% nonessential amino acids (NEAAs), 2 mM L-glutamine, 1% penicillin-streptomycin, and 5 ng/mL bFGF (see Note 2). 2. Aspirate the medium containing the cells and rinse the cells with 2 mL per well of PBS. 3. Dissociate cells by 500 μL 50% TrypLE™ Select and transfer the cells to LM511-E8-coated dishes. 4. Incubate the plate at 37 °C and 5% CO2 atmosphere for 3 min and tenderly move the plate in a few speedy level and vertical movements to scatter the cells equally across the cell culture surface. 5. Aspirate the medium again till the whole TrypLE™ Select remove. Then add 1 mL of iPSC medium supplemented with 10 μM Y27632. 6. Dissociate the iPSCs by scraping the cell colonies.

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7. Prepare a mixture of 25 μL of 0.4% Trypan Blue solution and 20 μL of cell media in a micro-centrifuge tube. 8. Gently tap the body of the tube containing the iPSCs (see Note 3). 9. Add 5 μL of cells microcentrifuge tube.

to

the

previously

prepared

10. Put the cover glass on hemacytometer and transfer 11–12 μL of the cell suspension between them. 11. Record each parameter in accordance with the Standard Operating Procedures (SOP) (see Note 4). 12. To count the cells by NucleoCounter® NC-100™, pipette the cell suspension up and down and withdraw small volume of supernatant (about 100 μL) to a new micro-centrifuge tube. Prepare the sample to be counted by the NucleoCounter® NC-100™ according to the manufacturer’s instruction. 13. Replate cells at a density of 1.5 × 105 cells per 6-well plate with 1.5 mL of StemFit medium containing 10 μM of the ROCK inhibitor Y-27632 and supplemented with 0.25 μg/cm2 iMatrix-511 (see Note 5). 14. Passage the cells 3 times to remove the feeder cells. 15. The iPSC medium be replaced by fresh medium within 24–48 h after passage and then change the medium every 2 days. 3.2 Generation of Embryoid Body (EB)

1. After aspirating the cells by TrypLE™ Select and removing feeder cells, culture the 1 × 106 cells per 2 mL EB medium per well of Aggrewell plates containing 80% DMEM/F12, 20%, knockout serum, 1% L-glutamine, 1% nonessential amino acids, 1% penicillin-streptomycin, 0.2% 2-mercaptoethanol (50 mM) supplemented with 10 μM of the ROCK inhibitor (Y-27632) according to the manufacturer’s instruction. 2. Incubate the plates for 24 h at 37 °C with 5% CO2 atmosphere. 3. By using a serological pipette, aspirate the aggregated EBs (see Note 6). 4. Dissociate the cells by a cell scraper from the surface of the culture dish. 5. Transfer cells to a sterile 15 mL conical tube. 6. Wash the culture medium with PBS. Then, transfer the PBS along with the remaining cells on the bottom of the plate to the 15 mL conical tube. 7. Centrifuge the tube for 5 min at 200 × g to pellet the cells. Then, aspirate the supernatant without disturbing the cell pellet and dispose it.

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8. In a 24-well plate with 0.1% gelatin coating, culture EBs in 2 mL of EB medium and keep them for 14–21 days. Media exchanges should be done every other day. 9. In a 96-well plate with 0.1% gelatin coating, culture the EBs in DMEM media containing 10% fetal bovine serum and 1% penicillin-streptomycin for 14 days. The medium should be replaced every 2 days (see Note 7). 10. Every 2 days, change the culture medium of the plates containing the cells. To this end, place the plate under the hood for 5 min in a fixed place so that the cells settle completely at the bottom of the plate. Then, remove the supernatant by a pipette and replace it with fresh culture medium. The size of EBs increases over time. 11. EBs can be arbitrarily differentiated into three germ layers (see Note 8). 3.3 Differentiation of iPSCs into Midbrain Dopaminergic Neurons (mDMA)

1. Culture EBs in neural induction medium. After 4 days, when the medium changed, incubate the plates for 6 days. 2. Enrich the culture medium with 100 ng/mL fibroblast growth factor-8a (FGF8a), 5 mg/mL heparin, 200 mM ascorbic acid, and 20 ng/mL brain-derived neurotrophic factor (BDNF). Then store the cells for 1–2 weeks (see Note 9). 3. Passage the progenitor cells and transfer the cells to Corning™ BioCoat™ Poly-D-Lysine/Laminin multiwell plate in final differentiation medium containing DMEM/F12 supplemented with 2 mM L-glutamine, N2 supplement, 20 mg/mL BDNF, 20 mg/mL glial-derived neurotrophic factor (GDNF), 0.5 mM N6, 29-O-dibutyryladenosine 39, 59 cyclic monophosphate sodium salt (dCAMP), 1 mg/mL laminin, and 1% [v/v] antibiotic/antimycotic. 4. After 14–21 days, progenitor cells convert into mature neural cells (see Note 10). Figure 1 shows all steps of producing EBs and dopaminergic neurons from iPSCs for Parkinson’s disease.

4

Notes 1. It is necessary to have a clean cell culture laboratory with laminar flow in order to follow aseptic practices throughout the procedure. Furthermore, it is critical to confirm that the iPS cell lines you are employing do not release live viruses or are not mycoplasma-contaminated [27]. 2. TrypLE™ Select is classified as cell dissociation enzymes of mammalian cells, which can be used in cell culture due to having functional characteristics similar to trypsin. Also, TrypLE™ Select contains no human- or animal-derived

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Fig. 1 All steps of producing EBs and dopaminergic neurons from iPSCs for Parkinson’s disease

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components. Moreover, unlike trypsin, it has room-temperature stability for at least 6 months with no inactivation required [28]. 3. Taping to the body of the iPSCs tube can help establish a uniform suspension in concentration and appearance [29]. 4. A standard operating procedure (SOP) is a bunch of composed rules or guidelines to help consummation of routine operations by staffs, intended to increment execution, further develop productivity, and guarantee quality through foundational homogenization, which can play a key role in promoting a successful quality system [30]. 5. Pre-coated iMatrix plates are not required for routine iPSC passage [31]. 6. Not pipetting hastily. Rinse the pipet with the liquid and release it gently to avoid foaming. 7. Culturing the cells in 0.1% gelatin-coated 96-well plate can help cells to differentiate spontaneously [32]. 8. The cell markers of germ layers can be analyzed by genome integrity analysis, immunostaining, and flow cytometry assays [32]. 9. The time of storage depends on appearance of rosette-like structures [33]. 10. In order to accurately check the function of the produced dopaminergic nerve cells, the cells produced should be examined by RT-PCR, immunohistochemical assay, dopamine release assay, electrophysiological analysis, and cell sorting [34]. References 1. Magrinelli F, Picelli A, Tocco P et al (2016) Pathophysiology of motor dysfunction in Parkinson’s disease as the rationale for drug treatment and rehabilitation. Parkinson’s Dis 2016: 1 2. DeMaagd G, Philip A (2015) Parkinson’s disease and its management: part 1: disease entity, risk factors, pathophysiology, clinical presentation, and diagnosis. P & T 40(8):504–532 3. Larijani B, Goodarzi P, Payab M et al (2019) The design and application of an appropriate Parkinson’s disease animal model in regenerative medicine. Cell Biol Transl Med 13:89–105 4. DeMaagd G, Philip A (2015) Parkinson’s disease and its management: part 1: disease entity, risk factors, pathophysiology, clinical presentation, and diagnosis. Pharm Ther 40(8):504

5. Naoi M, Maruyama W (1999) Cell death of dopamine neurons in aging and Parkinson’s disease. Mech Ageing Dev 111(2–3):175–188 6. Trist BG, Hare DJ, Double KL (2019) Oxidative stress in the aging substantia nigra and the etiology of Parkinson’s disease. Aging Cell 18(6):e13031 7. Go¨kc¸al E, Gu¨r VE, Selvitop R et al (2017) Motor and non-motor symptoms in Parkinson’s disease: effects on quality of life. Arch Neuropsychiatry 54(2):143 8. Poewe W (2008) Non-motor symptoms in Parkinson’s disease. Eur J Neurol 15:14–20 9. Kouli A, Torsney KM, Kuan W-L (2018) Parkinson’s disease: etiology, neuropathology, and pathogenesis. Exon Publications, pp 3–26

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10. Erro R, Vitale C, Amboni M et al (2013) The heterogeneity of early Parkinson’s disease: a cluster analysis on newly diagnosed untreated patients. PLoS One 8(8):e70244 11. Brugger F, Erro R, Balint B et al (2015) Why is there motor deterioration in Parkinson’s disease during systemic infections-a hypothetical view. NPJ Parkinson’s Dis 1(1):1–5 12. Zahoor I, Shafi A, Haq E (2018) Pharmacological treatment of Parkinson’s disease. Exon Publications, pp 129–144 13. Witt K, Kalbe E, Erasmi R et al (2017) Nonpharmacological treatment procedures for Parkinson’s disease. Nervenarzt 88(4):383–390 14. Larijani B, Hamidpour SK, Tayanloo-Beik A et al (2021) An overview of zebrafish modeling methods in drug discovery and development 15. Lertxundi U, Herna´ndez R, DomingoEchaburu S et al (2016) Pharmacotherapeutic challenges in Parkinson’s disease inpatients. IntechOpen 16. Martı´nez-Morales PL, Liste I (2012) Stem cells as in vitro model of Parkinson’s disease. Stem Cells Int 2012:1 17. Arjmand B, Roudsari PP, Alavi-Moghadam S et al (2021) Potential for stem cell-based therapy in the road of treatment for neurological disorders secondary to COVID-19. In: Regenerative engineering and translational medicine, pp. 1–15 18. Goodarzi P, Aghayan HR, Larijani B et al (2015) Stem cell-based approach for the treatment of Parkinson’s disease. Med J Islam Repub Iran 29:168 19. Li H, Jiang H, Zhang B et al (2018) Modeling Parkinson’s disease using patient-specific induced pluripotent stem cells. J Parkinsons Dis 8(4):479–493 20. Byers B, Lee H-l, Reijo Pera R (2012) Modeling Parkinson’s disease using induced pluripotent stem cells. Curr Neurol Neurosci Rep 12(3):237–242 21. Ke M, Chong C-M, Su H (2019) Using induced pluripotent stem cells for modeling Parkinson’s disease. World J Stem Cells 11(9): 634 22. Valadez-Barba V, Cota-Coronado A, Herna´ndez-Pe´rez O et al (2020) iPSC for modeling

neurodegenerative disorders. Regen Ther 15: 332–339 23. Larijani B, Foroughi-Heravani N, Alaei S et al (2021) Opportunities and challenges in stem cell aging. Cell Biol Transl Med 13:143–175 24. Goodarzi P, Aghayan HR, Soleimani M et al (2014) Stem cell therapy for treatment of epilepsy. Acta Med Iranica 52:651–655 25. Nicholson MW, Ting C-Y, Chan DZ et al (2022) Utility of iPSC-derived cells for disease modeling, drug development, and cell therapy. Cell 11(11):1853 26. Ye L, Swingen C, Zhang J (2013) Induced pluripotent stem cells and their potential for basic and clinical sciences. Curr Cardiol Rev 9(1):63–72 27. Chatterjee I, Li F, Kohler EE et al (2015) Induced pluripotent stem (iPS) cell culture methods and induction of differentiation into endothelial cells. In: Induced pluripotent stem (iPS) cells. Springer, pp. 311–327 28. Scientific TF (2015) Cell culture basics handbook. Gibco 29. Marchenko S, Flanagan L (2007) Counting human neural stem cells. JoVE 7:e262 30. Akyar I (2012) Standard operating procedures (what are they good for?). In: Latest research into quality control, pp. 367–391 31. Yamaguchi A, Ishikawa K-i, Akamatsu W (2021) Methods to induce small-scale differentiation of iPS cells into dopaminergic neurons and to detect disease phenotypes. Springer 32. Ma J, Guo R, Song Y et al (2019) Generation and characterization of a human induced pluripotent stem cell (iPSC) line (HEBHMUi001-A) from a sporadic Parkinson’s disease patient. Stem Cell Res 36:101417 33. Hartfield EM, Yamasaki-Mann M, Ribeiro Fernandes HJ et al (2014) Physiological characterisation of human iPS-derived dopaminergic neurons. PLoS One 9(2):e87388 34. Doi D, Samata B, Katsukawa M et al (2014) Isolation of human induced pluripotent stem cell-derived dopaminergic progenitors by cell sorting for successful transplantation. Stem Cell Reports 2(3):337–350

Methods in Molecular Biology (2023) 2736: 115–125 DOI 10.1007/7651_2022_468 © Springer Science+Business Media, LLC 2023 Published online: 15 December 2022

Standard Operating Procedure for Production of Mouse Brown Adipose Tissue-Derived Mesenchymal Stem Cells Babak Arjmand, Mostafa Rezaei-Tavirani, Sepideh Alavi-Moghadam, Akram Tayanloo-Beik, Mahdi Gholami, Shayesteh Kokabi-Hamidpour, Rasta Arjmand, Ahmad Rezazadeh-Mafi, Fereshteh Mohamadi-jahani, and Bagher Larijani Abstract Over the past years, stem cell technology was heralded as a significant breakthrough of the century in scrutinizing the intricacies of human body biology and discovering different therapeutic approaches. Recently, adipose tissue, as a suitable source of harvesting mesenchymal stem cells, has attracted the attention of many researchers in the field of regenerative medicine. Adipose tissue-derived mesenchymal stem cells can self-renew and differentiate into different types of cells such as adipocytes, chondrocytes, and osteoblasts. Adipose tissue, especially brown type, is considered an attractive cell source for various therapeutic purposes, such as restoring damaged tissue or fighting against diseases such as obesity. The growth of importance of stem cell applications in regenerative medicine has highlighted the need to seek appropriate mesenchymal stem cells sources. Recently, in the light of many efforts in the field of regenerative medicine, mice have gained increasing interest as a suitable source of adipose tissue for the extraction of mesenchymal stem cells, which can be used in the preclinical investigations in order to aid in the treatment of many human diseases. Key words Brown adipose tissue, Isolation, Mesenchymal stem cells, Mouse models, Standard operating procedure

Abbreviations BAT CD DMEM-LG DMSO ESCs FBS FSCs iPSCs KX MSCs PBS PVP-I

Brown adipose tissue Cluster of differentiation Dulbecco Modified Eagle medium-low glucose Dimethyl sulfoxide Embryonic stem cells Fetal bovine serum Fetal stem cells Induced pluripotent stem cells Ketamine-xylazine Mesenchymal stem cells Phosphate-buffered saline Povidone–iodine

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SSEA-4 SVF TMEM26 UCP1 WAT

1

Stage-specific embryonic antigen-4 Stromal vascular fraction Transmembrane protein 26 Uncoupling protein 1 White adipose tissue

Introduction Cell-based clinical trial uses a variety of cells, particularly stem cells or their derivatives, to enable injured, sick, or dysfunctional tissue respond to repair. Stem cells refer to primitive cells that are capable of either self-renew or differentiating into specialized cell types [1]. They can be classified based on origins into four groups including, embryonic stem cells (ESCs), induced pluripotent stem cells (iPSCs), fetal stem cells (FSCs), and adult stem cells. Investigations carried out on different categories of stem cells show that since the extraction and therapeutic uses of ESCs and FSCs are facing many ethical and methodological challenges, the use of alternative cellular sources is preferred. Accordingly, adult stem cells were recognized as one of the appropriate alternative sources [2]. Scientifically, the stability, accessibility, and safety of stem cells are among the most important criteria for selecting an efficient stem cell in regenerative medicine [3]. Herein, mesenchymal stem cells (MSCs), a subtype of adult stem cells, were suggested as suitable types of stem cells due to prominent characteristics such as selfrenewal, multi-potency with immunosuppressive potential, simple harvesting method, and low level of ethical issues [4, 5]. Hereupon, they have raised a big interest in regenerative medicine investigations [5–8]. MSCs originate from various tissues, such as bone marrow, adipose tissue, umbilical cord blood, and Wharton’s jelly [9–11]. However, employing some of these cell sources, e.g., bone marrow and Wharton’s jelly-derived stem cells, is challenging (based on invasiveness, a lack of adequate cell supply, the potential for infection to occur during harvest, or the lack of standard protocols in extraction) [12, 13]. Hence, attention has turned toward adipose tissue due to the use of simple and minimal invasive isolation procedures, providing an abundance of tissue sources and showing significant therapeutic promise in different fields of cell therapy and regenerative medicine [14], like reversing the obesity epidemic [15], treating female stress urinary incontinence [16], improving the nerve regeneration in peripheral nerve injuries [17], and enhancing wound healing [10, 18]. Histologically, adipose tissue is classified as white adipose tissue (WAT) and brown adipose tissue (BAT). WAT is found in almost every organ of the body, has small amounts of mitochondria, and mainly stores extra energy received from food in the form of triglycerides. Therefore,

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increasing the percentage of WAT can be a warning of the onset of many diseases, such as obesity and diabetes. In contrast, BAT is found only in some parts of the body and is rich in mitochondria. It is also worth mentioning that large amounts of uncoupling protein 1 (UCP1) are expressed in the mitochondria of BAT, which plays an important role in the thermogenesis process. Energy consumption, which is necessary to maintain a steady temperature, is inextricably tied to thermogenesis [19]. Since the BAT activity correlated positively with body metabolism, it is expected that the stem cells derived from the BAT are significantly more active than the white type [20]. Moreover, research indicates that BAT-derived stem cells are not only capable of in vitro multi-lineage differentiation can also be transformed into BAT cells with the same characteristics [21]. Because preclinical studies in animal models are critical for evaluating efficacy and safety prior to all types of clinical trials, it may be essential to conduct preclinical studies in appropriate animal models with a body structure similar to the human body, particularly mice, before using BAT-derived MSCs at the clinical level. In the context of animal studies, mouse models have great potential to advance the field of regenerative medicine as suitable animal models due to their outstanding properties. For example, a comparison of the mouse and human genomes has revealed that there is a high degree of genetic similarity between these two organisms. In addition, mouse models are affordable and practical research tools. The small size, availability of background knowledge on genetic and molecular traits, and ability to develop transgenic models of mice can pave the way to uncovering the complexity of diseases and discovering new therapeutic approaches [22]. On the other hand, the importance of using autologous cells in treatment cannot be ignored. Therefore, the use of autologous BAT MSCs seems to be very effective for mice in preclinical studies. Accordingly, it is very important to investigate how to isolate BAT-derived MSCs from mouse. Therefore, the present protocol provides standard operating procedure for production of mouse BAT-derived MSCs.

2

Materials

2.1 Mouse Brown Adipose Tissue Procurement

1. 70% ethanol (Kimia Alcohol Zanjan, Iran). 2. Six-week-old male C57BL/6 donor mice. 3. Ketamine–xylazine (KX) solution. 4. Povidone–iodine (PVP-I). 5. Razor. 6. Sterile scalpel, scissor, and forceps. 7. Sterile conical tubes 50 mL (SPL, Korea).

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8. Phosphate-buffered saline (PBS; Biowest, France). 9. Cool box with ice. 2.2 Harvesting, Brown Adipose Tissue, Mesenchymal Stem Cells Manufacturing, and Banking

1. 70% ethanol (Kimia Alcohol Zanjan, Iran). 2. Phosphate-buffered saline (PBS; Biowest, France). 3. Collagenase CLSAFA/AF (Worthington, USA) (1 mg/mL). 4. Dulbecco Modified Eagle medium-low glucose (DMEM-LG; Biowest, France). 5. Fetal bovine serum (FBS; ATOCEL, Austria). 6. TrypLE Express (Invitrogen, USA) (see Note 1). 7. Trypan blue solution 0.4% (Invitrogen, USA). 8. CryoSure-dimethyl sulfoxide (DMSO)(Sigma,USA). 9. StemPro™ Adipogenesis Differentiation Kit (Thermo Fisher Scientific, USA). 10. StemPro™ Osteogenesis Differentiation Kit (Thermo Fisher Scientific, USA). 11. Biological safety cabinet (Esco, Singapore). 12. Inverted microscope with phase contrast (Nikon, Japan). 13. Weighing balance (Sartorius, Germany). 14. Refrigerated centrifuge (swing-out rotor with buckets for 50 and 15 mL tubes) (Hettich, Germany). 15. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 16. Ultralow temperature freezer (New Brunswick Eppendorf, USA). 17. Liquid nitrogen tank (Statebourne Cryogenics, UK). 18. Sterile conical tubes 15 and 50 mL (SPL, Korea). 19. Sterile scalpel and forceps. 20. Sterile tissue culture plate (SPL, Korea). 21. Filter cap cell culture flasks (75 and 25 cm2) (SPL, Korea). 22. Hemocytometer and cover glass. 23. 0.2 μm sterile syringe filter (SPL, Korea). 24. 100 μm cell strainer (SPL, Korea). 25. Cryovial 2 mL (Nest, Germany). 26. Mr. Frosty freezing container (Nalgene™, Thermo Fisher Scientific, USA).

Standard Operating Procedure for Production of Mouse Brown Adipose Tissue. . .

3

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3.1 Brown Adipose Tissue Procurement

1. To extract inter-scapular BAT (Fig. 1), sterilize the surgical table with 70% ethanol. 2. Anesthetize the mouse intraperitoneally with a ketamine and xylazine solution (0.1 mL/20 g) (see Note 2). 3. Position it so that its back is facing up.

Pre-surgical measures Intraperitoneal injection of Ketaminexylazine

Shaving hair from the neck to below the scapulae

Disinfection of the surgical site with povidone iodine

BAT extraction Closing the incision by sutures

Extracting adipose tissue from the site

Making a small incision along the dorsal midline

Post-surgical measures Being under supervision of trained staff and considering a proper diet

Fig. 1 Isolation of brown adipose tissue from mouse. Isolation of adipose tissue from mouse can be divided into three stages: pre-surgery, surgery, and post-surgery stages. In the pre-surgery stage, the mouse is first anesthetized by intraperitoneal injection of ketamine–xylazine compounds. After anesthetization, the neck of the mice is shaved down to the below the mouse scapulae. The surgical site is disinfected by povidone–iodine. During the surgical stage, a small incision is made in the site between the two scapulae along the dorsal midline. Adipose tissue, which is a mixture of brown and white adipose tissue, is extracted from this site. The incision site is closed by some stitches. In the post-surgery phase, the mouse goes through its recuperation period under the supervision of an expert staff member who is skilled in working with animals, to accomplish a full recuperation. Abbreviation: BAT brown adipose tissue

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4. Use a razor to remove skin hairs as closely as possible to the skin. 5. Pull the skin from the shoulder blades of mouse over its head. 6. Disinfect the skin with PVP-I. 7. Cut open from the neck all the way along the back. 8. Right below the skin, between the shoulders, is where the BAT is located (inter-scapular). Raise the dark red tissue, and make very small cuts all around it to delicately separate it from the body (BAT can be seen as two lobes) (see Note 3). 9. Verify the consistency and color. 10. Place it in the tube containing 50 mL of PBS. 11. Keep in cool box with ice to transfer to the cell culture laboratory. 3.2 Isolation and Culture of Brown Adipose TissueDerived MSCs

1. Move the tube containing tissue from the cool box to the refrigerator, and keep it there until processing time at 4 C. 2. Turn on the biosafety cabinet before starting tissue processing. 3. Wipe the cabinet with 70% ethanol, and allow to ventilation air for 15 min. 4. Spray 70% ethanol on the outside of the tube containing tissue, and transfer it to biosafety cabinet. 5. Open the tube, under the biosafety cabinet, and aseptically transfer the tissue to a sterile culture plate. 6. Record the weight of the tissue. 7. Rinse it thoroughly with PBS. 8. Put it on another sterile culture plate with sterile forceps. 9. Cut it into very small pieces using a sterile scalpel and forceps. 10. Weigh type IV collagenase, and dissolve it in PBS (1 mg/mL) (for each mg of tissue, 1 mg of collagenase is utilized). 11. Use collagenase/ PBS solution to digest tissue fragments at 37 °C for 180 min. 12. Add equal volume of cooled PBS to digested tissue. 13. After that, centrifuge it (400 g for 10 min) to obtain the stromal vascular fraction (SVF). 14. Subsequently, seed the SVF into 25 cm2 culture flasks containing DMEM-LG, supplemented with 10% FBS. 15. Keep flasks at 37 °C, 5% CO2, and 95% humidity in a CO2 incubator. After 48 h of incubation, refresh the culture media to eliminate non-adherent cells and check the cellular morphology (Fig. 2) (see Note 4).

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Fig. 2 Microscopic appearance of brown adipose tissue-derived MSCs at primary culture (6th day)

16. When cells reach approximately 80–90% confluence, remove the media and wash them with pre-warmed PBS. 17. Then, add TrypLE™ Express (0.5–1 mL for 25 cm2 flask, and for other flasks, adjust the volume according to their surface) for cell dissociation. 18. Incubate the flask in the CO2 incubator (5–10 min), and check it for cell detachment. 19. Following the completion of cell dissociation, add equal volume of PBS, and transfer the cell suspension to 15 mL conical tube. 20. Centrifuge at 300 g for 5 min. 21. Carefully remove the supernatant, and resuspend the cell pellet in the appropriate volume of complete culture medium. 22. Utilize the hemocytometer and trypan blue 0.4% solution (1: 1) to count the MSCs and assess the viability of the cells. 23. Subculture the MSCS in the new flasks. 24. Place the flasks in the CO2 incubator (37 °C, 5% CO2, humidified), and carry with the cell expansion process until there are enough MSCs produced to create a master cell bank (usually 2–3 subcultures). The process is shown in Fig. 3.

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A

Extracting the tissue

Washing PBS

Segmenting

Digesting

Rinsing PBS

Collagenase 37°C 180 min

⫻2 Isolating SVF

Counting

Seeding

Incubation 37°C 5% CO2 95% humidity 48 h

400g 10 min

Refreshing culture media Centrifuging

Subculturing

Checking cell morphology

Resuspending

B Storing in freezer

C

Storing in Nitrogen tank

differentiation Fixing Osteogenic and lineage staining

Adipogenic lineage

Flow Cytometry

Fig. 3 Procedure for production of mouse brown adipose tissue-derived mesenchymal stem cells. After harvesting the brown adipose tissue from inter-scapular region of the mouse, it is washed by phosphatebuffered saline (PBS) and then cut into smaller pieces. Collagenase is used to digest the tissue at 37 °C for 180 min. The stromal vascular fraction (SVF) is isolated using centrifugation. Then, cells are counted, and their

Standard Operating Procedure for Production of Mouse Brown Adipose Tissue. . .

3.3 Characterization of Brown Adipose Tissue-Derived MSCs

3.4 Cryopreservation of Brown Adipose Tissue-Derived MSCs

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1. Utilize flow cytometry to examine the immunophenotypes (CD markers) (see Note 5). 2. Examine the ability of cells to differentiate in vitro into adipocytes (adipogenic differentiation) and osteoblasts (osteogenic differentiation), and confirm this by staining with Oil Red O and Alizarin Red (see Note 6). With 80–90% confluence, freezing for cell banking can be accomplished. 1. Harvest the MSCs using TrypLE™ Express (as described in Subheading 3.2). 2. Check cell count and viability by using hemocytometer and trypan blue 0.4% solution (1:1). 3. Make the freezing media by adding 10% DMSO to complete culture medium. 4. Suspend the MSCs in 1.8 mL freezing media, and divide them into 2 mL cryovials (about 10 × 106 cells/vial). 5. Put the cryovials in a Mr. Frosty freezing container after sealing them (see Note 7). 6. Put the container into an ultralow temperature freezer (-80 ° C), and leave it there for at least the next day. 7. After that, take out the cryovials and place them in a liquid nitrogen tank’s vapor phase.

4

Notes 1. The recombinant enzyme TrypLE Express is used to dissociate a variety of adherent mammalian cells and is free of animal origin. It is a direct substitute for trypsin and cleaves peptide bonds on the C-terminal sides of lysine and arginine. TrypLE Express is superior to trypsin and other dissociation agents because it is gentle on cells, stable at room temperature, and simple to use [23].

ä Fig. 3 (continued) viability is examined. The SVF were seeded into flasks containing low-glucose Dulbecco Modified Eagle medium supplemented with 10% fetal bovine serum (FBS) and kept at 37 °C, 5% CO2, and 95% humidity in a CO2 incubator. The culture media is refreshed. Cell morphology is then examined to ensure that the process is done correctly. Then, they should be subcultured or passaged. In order to harvest brown adipose tissue (BAT)-derived mesenchymal stem cells (MSCS), enzymatic separation is performed on the sample, and then it is centrifuged and transferred to cryovials. Cryovials are stored first in the freezer and then in nitrogen tanks. In order to characterize of BAT-derived MSCs, cells are differentiated into osteogenic and adipogenic lineage. Cells are fixed with paraformaldehyde (PFA). Then adipogenic and osteogenic linage are stained by Oil Red O and Alizarin Red S, respectively. The expression of specific markers is examined by flow cytometry technique. Abbreviations: PBS phosphate-buffered saline; SVF stromal vascular fraction

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2. The three Rs—replacement, reduction, and refinement— should be applied to every animal experiment. The greatest standards for animal housing maintenance, including environmental enrichment, are required. It is crucial to stress that conducting high-quality research should be totally consistent with and beneficial for maintaining high standards of animal care and welfare [24–27]. 3. The BAT is covered in a thin layer of WAT, which should be carefully removed. 4. The adherent population in isolation is displayed long cells that looked like fibroblasts. When adipose tissue cells are first isolated, they often have an un-uniform shape; however, as culture time goes on, the cells start to take on a more regular, fibroblast-like appearance. Aging causes BAT to change morphologically. BAT in appearance actually is brown. Unlike white adipocytes, brown adipocytes have many organelles and tiny lipid vacuoles [19]. 5. Transmembrane protein 26 (TMEM26), CD90, CD166, CD44, stage-specific embryonic antigen-4 (SSEA-4), CD73, CD105, and CD137 are all expressed by MSCs generated from BAT. The hematopoietic markers CD34, CD45, and human leukocyte antigen DR are also negative for them. On the other hand, they lack CD31 and had a population of CD146 that was only marginally positive [21]. 6. MSCs are always thought to be a desirable source for differentiating into cells from mesodermal origin, such as osteoblasts, adipocytes, and chondrocytes, according to their predominately mesodermal origin derived nature [28]. 7. For freezing process: Rapid cell freezing techniques are required, and a sterile cryovial must have components that are non-pyrogenic, non-cytotoxic, DNA-free, DNase-free, and RNase-free [29, 30]. References 1. Heidari-Keshel S, Rahimi A, Rezaei-Tavirani M et al (2019) Genomics, proteomics, and metabolomics for stem cells monitoring in regenerative medicine. In: Genomics, proteomics, and metabolomics, pp. 51–66. Springer 2. Falahzadeh K, Jalalvand M, Alavi-Moghadam S et al (2019) Trying to reveal the mysteries of stem cells using “omics” strategies. In: Genomics, proteomics, and metabolomics, pp. 1–50. Springer 3. Goodarzi P, Alavi-Moghadam S, Payab M et al (2019) Metabolomics analysis of mesenchymal stem cells. Int J Mol Cell Med 8(Suppl1):30

4. Arjmand B, Sarvari M, Alavi-Moghadam S et al (2020) Prospect of stem cell therapy and regenerative medicine in osteoporosis. Front Endocrinol 11:430 5. Gilany K, Masroor MJ, Minai-Tehrani A et al (2019) Metabolic profiling of the mesenchymal stem cells’ secretome. In: Genomics, proteomics, and metabolomics, pp. 67–81. Springer 6. Abedi M, Alavi-Moghadam S, Payab M et al (2020) Mesenchymal stem cell as a novel approach to systemic sclerosis; current status and future perspectives. Cell Regen 9(1):1–19

Standard Operating Procedure for Production of Mouse Brown Adipose Tissue. . . 7. Arjmand B, Goodarzi P, Aghayan HR et al (2019) Co-transplantation of human fetal mesenchymal and hematopoietic stem cells in type 1 diabetic mice model. Front Endocrinol 10: 761 8. Parhizkar Roudsari P, Alavi-Moghadam S, Payab M et al (2020) Auxiliary role of mesenchymal stem cells as regenerative medicine soldiers to attenuate inflammatory processes of severe acute respiratory infections caused by COVID-19. Cell Tissue Bank 21(3):405–425 9. Arjmand B, Ranjbaran N, Khatami F et al (2019) Different gene expression profile of mesenchymal stem cells from various sources. In: Genomics, proteomics, and metabolomics, pp. 83–96. Springer 10. Goodarzi P, Alavi-Moghadam S, Sarvari M et al (2018) Adipose tissue-derived stromal cells for wound healing. In: Cell biology and translational medicine, volume 4, pp. 133–149. Springer 11. Larijani B, Aghayan H, Goodarzi P et al (2015) Clinical grade human adipose tissue-derived mesenchymal stem cell banking. Acta Med Iran: 540–546 12. Kim D-W, Staples M, Shinozuka K et al (2013) Wharton’s jelly-derived mesenchymal stem cells: phenotypic characterization and optimizing their therapeutic potential for clinical applications. Int J Mol Sci 14(6):11692–11712 13. Schneider S, Unger M, Van Griensven M et al (2017) Adipose-derived mesenchymal stem cells from liposuction and resected fat are feasible sources for regenerative medicine. Eur J Med Res 22(1):1–11 14. Accession Bunnell BA (2021) Adipose tissuederived mesenchymal stem cells, vol. 10, p. 3433. MDPI 15. Jaber H, Issa K, Eid A et al (2021) The therapeutic effects of adipose-derived mesenchymal stem cells on obesity and its associated diseases in diet-induced obese mice. Sci Rep 11(1): 6291–6291 16. Arjmand B, Safavi M, Heidari R et al (2017) Concomitant transurethral and transvaginalperiurethral injection of autologous adipose derived stem cells for treatment of female stress urinary incontinence: a phase one clinical trial. Acta Med Iran: 368–374 17. Xaki S, Fathi A, Ariana M et al (2021) Effects of adipose-derived mesenchymal stem cells and human amniotic membrane on sciatic nerve repair in rats. Arch Neurosci 8(3) 18. Aghayan HR, Hosseini MS, Gholami M et al (2022) Mesenchymal stem cells’ seeded amniotic membrane as a tissue-engineered dressing

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for wound healing. Drug Deliv Transl Res 12(3):538–549 19. Tsuji W, Rubin JP, Marra KG (2014) Adiposederived stem cells: implications in tissue regeneration. World J Stem Cells 6(3):312–321. https://doi.org/10.4252/wjsc.v6.i3.312 20. Babahajian A, Shamseddin J (2018) Human adipose derived stem cell from white or brown fat; which one work better? J Med Physiol 3(1): 5 21. Silva FJ, Holt DJ, Vargas V et al (2014) Metabolically active human brown adipose tissue derived stem cells. Stem Cells 32(2):572–581 22. Nematizadeh M, Payab M, Gholami M et al (2020) Preclinical studies for development of biomedical products. In: Biomedical product development: bench to bedside, pp. 49–60. Springer 23. Laboshop (2022). https://laboshop.ae/prod uct/gibcotm-trypletm-express-enzyme-1xphenol-red-100-ml 24. Goodarzi P, Payab M, Alavi-Moghadam S et al (2019) Development and validation of Alzheimer’s disease animal model for the purpose of regenerative medicine. Cell Tissue Bank 20(2): 141–151 25. Guille´n J, Steckler T (2020) Good research practice: lessons from animal care and use. In: Bespalov A, Michel MC, Steckler T (eds) Good research practice in non-clinical pharmacology and biomedicine. Springer International Publishing, Cham, pp 367–382 26. Hubrecht RC, Carter E (2019) The 3Rs and humane experimental technique: implementing change. Animals (Basel) 9(10). https:// doi.org/10.3390/ani9100754 27. Larijani B, Goodarzi P, Payab M et al (2019) The design and application of an appropriate Parkinson’s disease animal model in regenerative medicine. Cell Biol Transl Med 13:89–105 28. Robert AW, Marcon BH, Dallagiovanna B et al (2020) Adipogenesis, osteogenesis, and chondrogenesis of human mesenchymal stem/stromal cells: a comparative transcriptome approach. Front Cell Dev Biol 8:561 29. Aghayan HR, Payab M, Mohamadi-Jahani F et al (2021) GMP-compliant production of human placenta-derived mesenchymal stem cells. Methods Mol Biol 2286:213–225. https://doi.org/10.1007/7651_2020_282 30. Larijani B, Aghayan H-R, Goodarzi P et al (2014) GMP-grade human fetal liver-derived mesenchymal stem cells for clinical transplantation. In: Stem cells and good manufacturing practices, pp. 123–136. Springer

Methods in Molecular Biology (2023) 2736: 127–137 DOI 10.1007/7651_2022_475 © Springer Science+Business Media, LLC 2023 Published online: 12 February 2023

Development and Validation of Type 2 Diabetic Zebrafish Model for Cell-Based Treatments Babak Arjmand, Sepideh Alavi-Moghadam, Shayesteh Kokabi-Hamidpour, Rasta Arjmand, Mostafa Rezaei-Tavirani, Bagher Larijani, Parisa Goodarzi, Neda Mehrdad, and Mohsen Rajaeinejad Abstract Diabetes mellitus can be categorized as one of the prolonged metabolic disorders that are associated with inappropriately elevated blood glucose levels. Among the subgroups of this disease, type 2 diabetes accounts for the most patients. Although pharmaceutical and non-pharmaceutical treatments have been employed to control the progression of the disease, as with any treatment approach, both therapeutic approaches are associated with side effects and challenges. Nowadays, the emergence of treatment methods based on stem cells has attracted the attention of researchers in order to treat diabetes fundamentally and provide a long-term solution. Since there are still blind spots regarding the positive and negative effects of these types of treatments, animal studies can give researchers a detailed insight into the effects of stem cellbased treatments. Recently, zebrafish has been proposed as a valuable animal model due to its outstanding genetic and physiological characteristics in biomedical studies including diabetes. Hereupon, in this protocol, the development and validation of type 2 diabetic zebrafish model for cell-based treatments have been explained. Key words Cell therapy, Diabetes mellitus, Modeling, Stem cell, T2DM, Zebrafish

Abbreviations DM FBS HF PBS RAS T1DM T2DM WHO

Diabetes mellitus Fetal bovine serum Heart failure Phosphate-buffered saline Recirculating aquaculture system Type 1 diabetes mellitus Type 2 diabetes mellitus World Health Organization

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Introduction Diabetes mellitus (DM) can be defined as a chronic metabolic disorder characterized by abnormal glucose homeostasis and impaired response to blood sugar levels resulting in elevated levels of glucose concentration in the bloodstream [1]. According to the estimates of the World Health Organization (WHO), it is expected that the number of patients with DM rises in the coming decades, which can have a great economic and health burden on individuals and societies [2]. Hence, early diagnosis and prognosis of DM play a key role in controlling the process of the disease throughout the world. Accordingly, many steps have been taken to discover prognostic biomarkers for DM, in order to apply the most effective interventions for patients [3, 4]. DM is most commonly subdivided into type 1 (T1DM) and type 2 diabetes mellitus (T2DM) [5]. Observations indicate that the risk of developing T2DM is higher and more prevalent among individuals, which can be associated with known complications such as heart failure (HF), peripheral sensory neuropathy, retinopathy, and kidney injuries [6]. Pathologically, two factors including dysfunction of pancreatic beta cells in insulin secretion and peripheral insulin resistance are involved in the occurrence of T2DM [7]. In recent years, a combination of lifestyle changes and the administration of oral and injectable drugs are suggested to patients for the treatment of T2DM. However, these treatment methods can be associated with limitations and challenges for patients. For instance, the risk of foot ulcers and the exacerbation of diabetes-related cardiovascular diseases following exercise, or the side effects of medications, can be included in the category of limitations associated with T2DM treatment approaches. Hence, the discovery of novel approaches with higher therapeutic potential is highlighted in relation to diabetes [8]. In recent years, stem cell-based treatment has received a great deal of attention as one of the candidate approaches in the treatment of diabetes [9–11]. One of the remarkable features of stem cell-based treatments is that these methods can lead to the production of functional β-cells that secrete insulin and solve the problem of regular insulin injections and administration of related drugs for patients. Hence, stem cell-based therapies can hold promise for a long-lasting or permanent solution to diabetes [12]. However, the clinical effectiveness of stem cell-based therapies and their possible side effects are not completely clear and require further studies [9]. Over the past years, preclinical studies and investigations of the effects of stem cells on animal models have been developed [13, 14] in order to address potential challenges and increase researchers’ insight into the effects of such therapeutic approaches in an in vivo biological platform [15]. In this context, zebrafish has been proposed as a suitable model for regenerative

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medicine studies related to endocrine diseases including DM [16]. The presence of orthologous genes between zebrafish and humans, genetic manipulability, and some outstanding physical characteristics like high physiological homology with some organs of the human body and the ability to regenerate some organs [17], transparent body, and a higher number of offspring per lifetime are among the prominent features which have made the zebrafish a valuable model in biomedical studies [18]. Therefore, in this protocol, the development of type 2 Diabetic zebrafish model has been investigated and the validation of a developed model for cell-based therapies has been explained in detail.

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Materials

2.1 Preparing Animal Model 2.1.1 Fish Maintenance and Induction of Type 2 Diabetes

1. Zebrafish (Danio rerio) adult wild type of both sexes (4–6 months old). 2. Recirculating aquaculture systems (RAS) (see Note 1). 3. Commercial flakes containing 48% protein, 8% fat, and 2% fiber (TetraMin™, NC, USA). 4. Live brine shrimp. 5. Glucose (Nuclear™) (55.5 mM (w/v)).

2.2 Monitoring of Diabetes for Validation of Type 2 Diabetic Zebrafish Model

1. 0.04% Tricaine MS-222 (tricaine methane sulphonate) (Sigma, Germany).

2.2.1 Measurement of Blood Glucose Level

4. Sterile scissors.

2. Glucometer strip (AUVON, USA). 3. Tissue paper. 5. Sterile tray or glass. 6. Heparinized 100 μL microcapillary tube (Sarstedt, USA). 7. Heparinized 40 mm microhematocrit tube (StatSpin, USA).

2.2.2 Gene Expression Analysis by Quantitative Real-Time RT-PCR (RTqPCR)

1. Phosphate-buffered saline (PBS) (Biowest, USA). 2. Fetal bovine serum (FBS) (Biowest, USA). 3. 1-mL syringe (AVA, Iran). 4. 30-μm nylon filter (Falcon® Cell Strainer, Corning, Durham, NC, USA). 5. 15 mL falcon (SPL, Korea). 6. RNase-free Microfuge Tubes (SPL, Korea). 7. TRIzol™ reagent (Invitrogen, Carlsbad, CA, USA). 8. Chloroform (Sigma, USA). 9. Isopropanol (Sigma, USA).

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10. DNase-/RNase-free or DEPC-treated water (see Note 2). 11. cDNA Synthesis Kit (Qiagen, Valencia, CA). 12. LightCycler® 480 SYBR Green I Master (Roche Life Sciences, Germany) (see Note 3). 13. RNase Zap (Ambion Inc., Austin, TX). 14. 75% Ethanol, prepared with RNase-free water in a wash bottle. 15. DNase/RNase -free tubes, 2 mL (e.g., Safe-Lock Eppendorf; Fisher Scientific, Pittsburgh, PA). 16. Liquid nitrogen or dry ice/ethanol bath for snap-freezing after excision (see Note 4). 17. DNase/RNase free-50 mL Falcon tubes (SPL, Korea). 18. LC 480 Multiwell Plate 96 (Roche Diagnostics). 19. Forward and reverse PCR primers at 100 μM each. 20. NanoDrop spectrophotometer (Thermo Fisher Scientific, USA). 21. LightCycler® 96 System (Roche Life Sciences, Germany). 22. Pipettes (Eppendorf, Germany). 23. DNase-/RNase-free Pipette Tips (Eppendorf, Germany). 24. Vortex (Stuart, England). 25. Refrigerated Centrifuge (swing-out rotor with buckets for 50 and 15 mL tubes) (Hettich, Germany). 26. Microcentrifuges (Labnet, USA).

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Methods

3.1 Preparing Animal Model 3.1.1 Fish Maintenance

All protocols must be approved by the animal care committee and follow the international guidelines of veterinary medicine along with the guides on the care and use of fish in research and animal welfare considerations (see Notes 5–7). As shown in Fig. 1, fish are maintained as follows: 1. Maintain animals in a recirculating aquaculture system in a cycle of 14 h light: 10 h darkness (at 28 °C). 2. Feed them three times a day with commercial flakes supplemented with live brine shrimp.

3.1.2 Induction of Type 2 Diabetes

As shown in Fig. 1, type 2 diabetes is induced as follows: 1. Place zebrafish in aquaria containing glucose solution which is diluted in water (55.5 mM (w/v)) and maintain at room temperature for 14 days (as already mentioned in the previous part, the feeding schedule and general maintenance measures should be followed.)

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Fig. 1 Principles of housing and husbandry of normal and diabetic zebrafish. All principles related to animal transportation (if using animals raised in other laboratories) must be observed. If zebrafish grown in other laboratories are used, quarantine systems must be used and all principles should be observed. The standard number of adult zebrafish in each aquarium is 4–10 fish. The standard number of embryos is a maximum of 100/35 mL in a 9 cm Petri dish. The standard number of 5–10 dpf larvae is equal to 250/l. in a 9 cm Petri dish. Zebrafish should be raised with a combination of live feeds and processed dry feeds. Keep zebrafish in a recirculating aquaculture system in a cycle of 14 h light: and 10 h darkness. In order to preserve the natural behavior of zebrafish shoaling, it is recommended to keep them in groups. All standards related to chlorine, ammonia, nitrite, nitrate, conductivity, general water hardness, and water pH should be evaluated and monitored frequently. Embryos should be kept at a temperature of 28.5 ± 0.5 °C to 120 hpf. Larvae and adults should be kept at 24–29 °C. Experiments with time limits can be conducted at temperatures ranging from 15 to 39 °C. Attempt to prevent sibling mating from causing repeated inbreeding. All information related to zebrafish should be recorded. The principles of the zebrafish nomenclature committee should be considered. Use sterilized equipment to prevent cross-contamination and co-infections between humans and animals. Filter systems, regulation of water chemistry system, UV light, temperature, and light regulation systems should be monitored [19]

2. To prevent infection with opportunistic microorganisms, change the glucose solution three times a week (see Note 8). 3. Monitor the signs of stress, such as trouble swimming or excessive gill movement (see Note 9).

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3.2 Monitoring of Diabetes for Validation of Type 2 Diabetic Zebrafish Model 3.2.1 Measurement of Blood Glucose Level

Fish must fast for 12 h before any blood measurement procedures begin and spend 15 min in an aquarium filled with water devoid of glucose in order to prevent glucometer strip contamination. 1. For measurement of the blood glucose level, first, use 0.04% Tricaine MS-222 (tricaine methanesulphonate) at 28.5 °C in order to anesthetize the fish (when fish present a lack of motor coordination, slow breathing, and no responsiveness to exterior stimuli, they are deemed to be anesthetized). 2. Then, remove the zebrafish from solution. 3. Pat them dry using tissue paper and place them on a glass surface or sterile tray. 4. After that, decapitate by using scissors to cleanly cut through the pectoral girdle and collect entire blood (the cut is immediately anterior to the articulation of the pectoral fin with the girdle, and severed the heart). 5. Apply glucometer strip directly to the cardiac blood to measure the blood glucose level (see Note 10). 6. Collect whole blood sample by holding either a heparinized 100 μL microcapillary tube adjacent to the severed heart or a heparinized 40 mm microhematocrit tube for repeated measurement of a sample.

3.2.2 Gene Expression Analysis by Quantitative Real-Time RT-PCR (RTqPCR)

1. Allow zebrafish male and female mating. 2. 48-h post-fertilization of zebrafish, wash embryos three times with 0·9 × PBS, and after each wash allow settling by gravity. 3. Remove 0·9 × PBS and immerse in 0·9 × PBS containing 2% FBS. 4. Crush them with the plunger of a 1-mL syringe and pass through a 30-μm nylon filter. 5. Wash the obtained cells with 0·9 × PBS containing 2% FBS. 6. Pellet the cells by centrifugation at 200 × g for 5 min at room temperature. 7. Then resuspended in 0·9 × PBS containing 2% FBS. 8. For total RNA extraction, use RNaseZap as a surface cleaning agent to eliminate RNases immediately and then lyze the cells using TRIzol™ reagent by pipetting up and down several times (if you can’t homogenize your samples immediately, snap freeze them and keep them at -70 °C). 9. To separate nucleoprotein complexes, incubate the homogenized sample for 5 min at room temperature. 10. For each volume of TRIzol reagent, add 0.2 volumes of chloroform.

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11. After tightly capping the tubes, vigorously vortex the samples for 15 s. 12. Incubate samples at room temperature for 5 min. 13. Centrifuge the samples for 15 min at 4 °C with a maximum force of 12,000 g (three phases of separation will occur in the combination) (see Note 11). 14. To precipitate the RNA from the aqueous phase, mix 0.5 mL of isopropanol with 1 mL of TRIzol. 15. Samples should be centrifuged at a maximum of 12,000 g for 10 min at 4 °C after being incubated for 10 min at room temperature (on the side or bottom of the tube, the RNA precipitate will form a pellet that can be challenging to perceive with the naked eye). 16. The RNA pellet should be washed once with 75% ethanol after the supernatant has been removed. 17. Vortex the samples together, then centrifuge them for 5 min at 4 °C at no more than 8000 g (to get rid of any remaining ethanol, repeat the washing process described). 18. Vacuum or air-dry the RNA pellet for 5–10 min. 19. By repeatedly passing the solution through a pipette tip, dissolve the RNA in the DNase/RNase free or DEPC-treated water. 20. Quantify the total extracted RNA by NanoDrop spectrophotometer (see Note 12). 21. Synthesize the cDNA using cDNA Synthesis Kit from 1 μg of total RNA. 22. Perform quantitative PCR using LightCycler® 480 SYBR Green I Master (to detect double-strand cDNA synthesis). 23. Done reactions in a volume of 25 μL using 12.5 μL of diluted cDNA (1:50 for reference genes) by LightCycler® 96 System (see Note 13). 24. Analyze the efficiency per sample using the best real-time PCR analysis software. Figure 2 shows all steps for validation of type 2 diabetic Zebrafish Model.

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Notes 1. RAS are commonly used in home aquariums and fish farms when water exchange is limited and biofiltration is required to avoid ammonia toxicity [20, 21].

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Fig. 2 All steps for validation of type 2 diabetic zebrafish model

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2. In molecular biology workflows where RNA is very sensitive and can be rapidly degraded by RNases, RNase-free (contains no RNA-degrading enzyme) reagents and products are required to prevent RNase contamination. Also, for preventing DNase contamination, which has a significant impact on genetic analysis, DNase-free reagents and products that do not contain DNase (an enzyme that breaks down DNA) are necessary [22, 23]. 3. The LightCycler® 480 SYBR Green I Master is a PCR hot start reaction mix with a single component. For PCR product detection and characterization, it includes FastStart Taq DNA Polymerase and DNA double-strand-specific SYBR Green I dye. Because the mix is given as an all-in-one master reagent, reaction preparation is as simple as adding template DNA and primers. The mix is suitable for high-throughput applications in 96- or 384-well plates on the LightCycler® 480 Instrument and may be used with different forms of DNA (e.g., genomic DNA or cDNA) [24]. 4. Snap freezing (flash freezing) is a process in which the samples are reduced very quickly to a temperature below -70 °C using dry ice or liquid nitrogen [25]. 5. The animal care committee is an integral part of animal care. It uses programs to prepare informed and ethical decisions about the appropriateness of animals’ involvement in research, teaching, or pilot projects, manage the facility’s animal care, and ensure high animal standards and welfare [26, 27]. 6. The international guidelines of veterinary medicine are useful in malpractice lawsuits because they facilitate decision-making and reduce mistakes. In reality, veterinarians frequently embrace such recommendations, which they utilize to improve treatment, reduce stress and ambiguity, and justify their procedures to customers [28]. 7. This is a vital issue while maintaining any animal in captivity, particularly in order to satisfy welfare requirements. To reach the minimum welfare criteria, we must examine five critical factors: (1) Provide the captive animal with a speciesappropriate feed as well as clean drinking water. A decent diet must provide all nutritional requirements to keep the animal in good health. (2) The animal should be given a proper habitat for its demands. This can vary according to the species, and it can relate to indoor and outdoor places, house and cages, as well as size and space. The surroundings should safeguard the animal from the elements, possible predators, and other damage. (3) Any animal in a cage should be permitted to behave normally and have access to an environment that permits this. It is also important to consider whether the animal is a social

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species that typically coexist in groups or pairs. If this is the case, it can have an effect on the animal’s welfare. For socially imprisoned animals, social interaction is essential. The right to a life free from harm, pain, disease, and suffering is something that all animals have. When an animal is in pain, its well-being is in grave danger, and something needs to be done [29]. 8. Opportunistic microorganisms are normally non-pathogenic germs that can become pathogens under specific conditions. They wait until the host’s immune system is weakened before emerging from their prolonged dormancy and attacking [30]. 9. Excessive rises in blood glucose levels may cause indicators of stress, such as difficulties swimming or excessive gill movement, as well as an increase in the percentage of fish mortality [31]. 10. Zebrafish have blood glucose levels (50–75 mg/dL), which is similar to the range (70–120 mg/dL) for humans [31]. Total free glucose levels in zebrafish embryos ranged from 8.2 ± 0.55 mmol L-1 to 19 ± 1.26 mmol L-1 [32]. 11. Three phases of separation will occur in the combination: (1) organic phase in lower, (2) interphase, and (3) RNA present in the upper aqueous phases [33]. 12. The maximum absorbance for nucleic acids is at 260 nm. The purity of DNA and RNA extractions has traditionally been assessed using the ratio of this absorbance maximum to the absorbance at 280 nm. For DNA, a 260/280 ratio of 1.8 is generally regarded as “pure,” whereas for RNA, a ratio of 2.0 is generally regarded as “pure” [34]. 13. Isoforms of the insulin receptor gene (insra-1, insrb-1 and insrb-2) as well as PEPCK are significant genes for zebrafish used in the monitoring of diabetes. After increasing the glucose level, the expression level of insra-1, insrb-1, and insrb-2 will increase and the expression level of PEPCK will decrease [31, 35]. References 1. Kharroubi AT, Darwish HM (2015) Diabetes mellitus: the epidemic of the century. World J Diabetes 6(6):850 2. Goodarzi P, Aghayan HR, Larijani B et al (2015) Stem cell-based approach for the treatment of Parkinson’s disease. Med J Islam Repub Iran 29:168 3. Esmati P, Najjar N, Emamgholipour S et al (2021) Mass spectrometry with derivatization method for concurrent measurement of amino acids and acylcarnitines in plasma of diabetic type 2 patients with diabetic nephropathy. J Diabetes Metab Disord 20(1):591–599

4. Hosseinkhani S, Arjmand B, DilmaghaniMarand A et al (2022) Targeted metabolomics analysis of amino acids and acylcarnitines as risk markers for diabetes by LC–MS/MS technique. Sci Rep 12(1):1–11 5. Sapra A, Bhandari P, Wilhite Hughes A (2021) Diabetes Mellitus (Nursing). 6. Tayanloo-Beik A, Roudsari PP, Rezaei-Tavirani M et al (2021) Diabetes and heart failure: multi-omics approaches. Front Physiol 12: 705424

Development and Validation of Type 2 Diabetic Zebrafish Model for Cell-Based. . . 7. Galicia-Garcia U, Benito-Vicente A, Jebari S et al (2020) Pathophysiology of type 2 diabetes mellitus. Int J Mol Sci 21(17):6275 ˜ alver JJ, Martı´n-Timo´n I, Sevillano8. Marı´n-Pen Collantes C et al (2016) Update on the treatment of type 2 diabetes mellitus. World J Diabetes 7(17):354 9. Ghodsi M, Heshmat R, Amoli M et al (2012) The effect of fetal liver-derived cell suspension allotransplantation on patients with diabetes: first year of follow-up. Acta Med Iranica:541–546 10. Sarvari M, Alavi-Moghadam S, Aghayan HR et al (2021) Stem cells researches and therapies towards endocrine diseases treatment; strategies, challenges, and opportunities. J Diabetes Metab Disord:1–7 11. Larijani B, Goodarzi P, Payab M et al (2019) Metabolomics and cell therapy in diabetes mellitus. Int J Mol Cell Med 8(Suppl1):41 12. Rahim F, Arjmand B, Shirbandi K et al (2018) Stem cell therapy for patients with diabetes: a systematic review and meta-analysis of metabolomics-based risks and benefits. Stem Cell Investig 5:40 13. Song W-J, Shah R, Hussain MA (2010) The use of animal models to study stem cell therapies for diabetes mellitus. ILAR J 51(1):74–81 14. Sakata N, Yoshimatsu G, Tsuchiya H et al (2012) Animal models of diabetes mellitus for islet transplantation. Exp Diabetes Res 2012:1 15. Nagaya M, Hasegawa K, Uchikura A et al (2021) Feasibility of large experimental animal models in testing novel therapeutic strategies for diabetes. World J Diabetes 12(4):306 16. Arjmand B, Tayanloo-Beik A, Foroughi Heravani N et al (2020) Zebrafish for personalized regenerative medicine; a more predictive humanized model of endocrine disease. Front Endocrinol 11:396 17. Larijani B, Hamidpour SK, Tayanloo-Beik A et al (2021) An overview of zebrafish modeling methods in drug discovery and development 18. Tayanloo-Beik A, Kokabi Hamidpour S, Abedi M et al (2022) Zebrafish modeling of autism spectrum disorders, current status and future prospective. Front Psychiatry 13:1514 19. Alestro¨m P, D’Angelo L, Midtlyng PJ et al (2020) Zebrafish: housing and husbandry recommendations. Lab Anim 54(3):213–224 20. Ebeling JM, Timmons MB (2010) Recirculating aquaculture. Cayuga Aqua Ventures, Ithaca 21. Ebeling JM, Timmons MB (2012) Recirculating aquaculture systems. Aquacult Prod Syst 1: 245–277 22. Bettarel Y, Sime-Ngando T, Amblard C et al (2000) A comparison of methods for counting

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viruses in aquatic systems. Appl Environ Microbiol 66(6):2283–2289 23. Demeke T, Jenkins GR (2010) Influence of DNA extraction methods, PCR inhibitors and quantification methods on real-time PCR assay of biotechnology-derived traits. Anal Bioanal Chem 396(6):1977–1990 24. https://lifescience.roche.com/global/en/ products/others/lightcycler-480-sybr-greeni-master-358112.html (2022) 25. Steu S, Baucamp M, von Dach G et al (2008) A procedure for tissue freezing and processing applicable to both intra-operative frozen section diagnosis and tissue banking in surgical pathology. Virchows Arch 452(3):305–312 26. DeHaven WR (2002) Best practices for animal care committees and animal use oversight. ILAR J 43(Suppl_1):S59–S62 27. Steneck NH (1997) Role of the institutional animal care and use committee in monitoring research. Ethics Behav 7(2):173–184 28. Sherman DM (2007) Tending animals in the global village: a guide to international veterinary medicine. John Wiley & Sons 29. Beaver BV, Bayne K (2014) Chapter 4 - Animal welfare assessment considerations. In: Bayne K, Turner PV (eds) Laboratory animal welfare. Academic Press, Boston, pp 29–38. https:// doi.org/10.1016/B978-0-12-385103-1. 00004-X 30. Jurado V, Laiz L, Rodriguez-Nava V et al (2010) Pathogenic and opportunistic microorganisms in caves. Int J Speleol 39(1):2 31. Capiotti KM, Junior RA, Kist LW et al (2014) Persistent impaired glucose metabolism in a zebrafish hyperglycemia model. Comp Biochem Physiol B 171:58–65 32. Singh A, Castillo HA, Brown J et al (2019) High glucose levels affect retinal patterning during zebrafish embryogenesis. Sci Rep 9(1): 4121. https://doi.org/10.1038/s41598019-41009-3 33. Chomczynski P, Wilfinger W, Mackey K Singlestep method of total RNA isolation by Guanidine–Phenol extraction. In: eLS, pp 1 – 6 . h t t p s : // d o i . o r g / 1 0 . 1 0 0 2 / 9780470015902.a0003799.pub3 34. Desjardins RP, Conklin DS (2011) Microvolume quantitation of nucleic acids. Curr Protoc Mol Biol 93(1):A. 3J. 1-A. 3J. 16 35. Elo B, Villano C, Govorko D et al (2007) Larval zebrafish as a model for glucose metabolism: expression of phosphoenolpyruvate carboxykinase as a marker for exposure to antidiabetic compounds. J Mol Endocrinol 38(4): 433–440

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Neuromuscular Junction-on-a-Chip for Amyotrophic Lateral Sclerosis Modeling Sepideh Alavi-Moghadam, Shayesteh Kokabi-Hamidpour, Mostafa Rezaei-Tavirani, Bagher Larijani, Rasta Arjmand, Fakher Rahim, Ahmad Rezazadeh-Mafi, Hossein Adibi, and Babak Arjmand Abstract Amyotrophic lateral sclerosis (ALS) is a progressive and degenerative disorder of the nervous system that can significantly reduce the physical activity of patients at the end stages. As the field of disease pathophysiology has advanced in recent years, studies have looked at the role of neuromuscular junction’s dysfunctionality in ALS. In the past years, various in vitro and in vivo models were developed to scrutinize the underlying mechanisms of the disease and investigate the effects of candidate drugs, but the application of the developed models faced many challenges. Hence, the attentions shifted to cutting-edge technologies such as the organ-on-a-chip, which can mimic the pathophysiology of the disease as a special biological platform using patient-derived cells in the integration of engineering sciences to expand researchers’ perspectives on the disease. In addition, organ-on-a-chip technology can reduce some of the challenges of using other in vitro and in vivo models, which can pave the way for other discoveries and advances in this disease. Key words Amyotrophic lateral sclerosis, Neuromuscular junction, NMJ, Organ on a chip, Spheroid

Abbreviations AA ALS Anti-Anti BDNF DMEM DMEM/F-12 DMSO DPBS FBS GCDR Ham’s F-12 hiPSCs HS Human recombinant IGF-1 KSR

L-ascorbic acid Amyotrophic lateral sclerosis Antibiotic-antimycotic Brain-derived neurotrophic factor Dulbecco’s Modified Eagle Medium Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 Dimethylsulfoxid Dulbecco’s phosphate-buffered saline Fetal bovine serum Gentle cell dissociation reagent Ham’s Nutrient Mixture F-12 Human-induced pluripotent stem cells Horse serum Human recombinant insulin-like growth factor-1 KnockOut serum replacement

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MEFs MEM-α MNs MNPCs NEPs NMJ OOC PLO Pur RA VPA

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Mouse embryonic fibroblasts Minimum essential medium–α Motor neurons Motor neuron progenitor cells Neuroepithelial progenitors Neuromuscular junction Organ-on-a-chip Poly-L-ornithine Purmorphamine Retinoic acid Valproic acid

Introduction The nervous system is a complex biological construct consisting of the central and peripheral nervous systems that direct the vital actions of the body [1]. Nervous system originates from cells known as neural stem cells during development, which are able to form a wide neural network in the post-embryonic stages [2]. In the past years, the number of patients with disorders related to the nervous system has been increasing [3]. In addition, the outbreak of Covid-19 disease in the past years has also been associated with the occurrence of devastating disorders symptoms related to the nervous system in patients [4], which has highlighted the need to seek potential treatment approaches [4, 5]. One of the nervous system-related diseases, which has made researchers for years to discover new approaches with therapeutic potential, is amyotrophic lateral sclerosis (ALS). In broad biological terms, ALS can be defined as a progressive neurodegenerative disorder, which can affect motor nervous system. As a result of the involvement of the motor nervous system, muscles gradually become weak and their function is disturbed. Hence, the progression of disease can eventually lead to paralysis or even death [6]. Clinically, in ALS conditions, degeneration of motor neurons in the brain (upper motor neurons) and spinal cord (lower motor neurons) is considered a pathognomonic phenomenon engaged in the disease process. Gaining a deeper insight into the pathophysiological mechanisms of the disease is tied to three hypotheses, including dying forward (corticomotoneuron hyperexcitabilitymediating degeneration of motor nerve cells toward the spinal cords), dying back (muscle dysfunctionality or degradation of neuromuscular junction (NMJ) led to retrograde degeneration of nerve axons), and the independent degeneration process (separate degeneration of upper motor neurons from lower motor nerves) [7]. However, due to the complexity of the ALS pathophysiology, some studies place more emphasis on the precedence of the dying

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forward hypothesis than the dying back [8], and some of the studies vice versa [9]. On the whole, it should be noted that whether the ALS commence by degeneration of the upper motor neurons or degeneration of the lower motor nerve cells, the NMJ dysfunctionality can be one of the possible processes involved in ALS, which should be considered in the pathological assessments of the disease [10]. Over the past years, various in vitro and in vivo models have been conducted to scrutinize the mechanisms underlying ALS pathogenesis. While the models employed have been successful in increasing awareness and knowledge about ALS, they also have disadvantages. For example, in vitro models could not demonstrate the interaction between different biological organs and systems of an organism in relation to disease [11]. Additionally, the application of animal models involves many challenges, even though they provide a more complex biological platform for disease investigation [12]. Moreover, genetic complexity, the lack of highly sensitive biomarkers, issues and challenges related to trial design, and failure to diagnose the disease in time are among the obstacles in the path of ALS treatment [13]. Accordingly, the introduction of new technologies, such as organ-on-a-chip (OOC), was intended to remove some of the obstacles in the way to scrutinize the pathophysiology of diseases and pave the way to discover more targeted and effective treatment approaches in biomedical studies [14]. Structurally, organ-on-chips are engineered platforms to mimic natural or artificial biological systems in small dimensions, whose structure consists of four basic components, including stem cells which can be also derived from the patient, microfluidic system, and biological stimuli to mimic the biological environment of the human body, and finally sensors to track and comprehensively monitor modeling and biological response to chemicals and experimental drugs in vitro. Due to its ability to mimic neurodegenerative diseases including ALS, OOC has received a lot of attention in recent years, potentially providing a new approach to understanding ALS and discovering new treatments [15]. Hereupon, this protocol first deals with the formation of the two main components of the NMJ on the chips, i.e., motor neurons (MNs) and skeletal muscle cells, and then explains how to seed the cells on a NMJ chip.

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Materials

2.1 Motor Neuron Spheroids Generation

1. Human-induced pluripotent stem cells (hiPSCs) (Royan Institute, Iran). 2. CF1 Mouse Embryonic Fibroblasts, irradiated (MEFs; Gibco™, A34180).

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3. Dispase (Gibco™, 17105041). 4. Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12; Gibco™, 11320033). 5. Neurobasal™ medium (Gibco™, 21103049). 6. N2 (Invitrogen). 7. B27 (Invitrogen). 8. Ascorbic acid (Sigma-Aldrich). 9. GlutaMAX (Gibco™, 35050061). 10. Penicillin-streptomycin (Invitrogen, 15140122). 11. CHIR99021 (Stem Cell Technologies). 12. DMH-1 (Sigma-Aldrich). 13. SB431542 (Stem Cell Technologies). 14. Retinoic acid (RA; Sigma-Aldrich). 15. Purmorphamine (Pur; Stem Cell Technologies). 16. Valproic Acid (VPA; Stem Cell Technologies). 17. Cryovial 2 mL (Nest, Germany). 18. Fetal Bovine Serum (FBS) Biopharm—EDQM certified (Biowest, USA). 19. Dimethylsulfoxid (DMSO; Thermo Fisher Scientific). 20. Liquid nitrogen tank (Statebourne Cryogenics, UK). 21. Water bath (Memmert, Germany). 22. 70% ethanol (kimia alcohol zanjan, Iran). 23. Sterile conical tubes 50 mL (SPL, Korea). 24. Antibiotic-Antimycotic (Anti-Anti, Thermo Fisher Scientific, 15240-062). 25. Refrigerated centrifuge (swing-out rotor with buckets for 50 and 15 mL tubes) (Hettich, Germany). 26. T-25 Flask (Thermo Fisher Scientific). 27. Poly-L-ornithine (PLO; Sigma, P3655). 28. Laminin (Thermo Fisher Scientific, 23017015). 29. L-Ascorbic acid (AA; Sigma-Aldrich, A5960). 30. Y-27632 (ROCK inhibitor; Sigma-Aldrich). 31. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 32. Accutase (Thermo Fisher Scientific, A1110501). 33. Compound E (Stem Cell Technologies, 73954). 34. Brain-derived neurotrophic factor (BDNF, Sigma-Aldrich). 35. NucleoCassette™ (Chemometec, Denmark).

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36. NucleoCounter® NC-100™ (Chemometec, Denmark). 37. 6-well plates (SPL, Korea). 38. 96-well Clear Round Bottom Ultra-Low Attachment Microplate (Corning™ 7007). 39. Centrifuge with adapter (Eppendorf, 5810).

for

culture

well

plates

40. Cell Scraper (Falcon, Cat. no. 353085). 41. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea). 42. Microcentrifuge tube (SPL, Korea). 43. Biological safety cabinet (Esco, Singapore). 2.2 Skeletal Muscle Cells Formation and Injection into an NMJ Chip

1. 4% Pluronic F-127 (Sigma-Aldrich). 2. Distilled water. 3. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 4. Human iPSC-derived skeletal myoblasts (Cellular Dynamic International, cat. no. SKM-301-020-001-PT). 5. T-75 flasks (Thermo Fisher Scientific). 6. Dulbecco’s phosphate-buffered saline (DPBS; Thermo Fisher Scientific). 7. Fibronectin (Sigma-Aldrich). 8. Minimum essential medium–α (MEM-α; Thermo Fisher Scientific). 9. 8-Bromo-cyclic AMP (Stem Cell Technologies). 10. CHIR99021 (Stem Cell Technologies). 11. Dorsomorphin (Stem Cell Technologies). 12. KnockOut Serum Replacement (KSR; Life Technologies, cat. no. 10828-010). 13. Dulbecco’s Modified Eagle Medium (DMEM; Gibco™). 14. Horse serum (HS; Invitrogen, cat. no. 16050130). 15. Human recombinant insulin-like growth factor-1 (Human recombinant IGF-1; Stem Cell Technologies). 16. Penicillin-streptomycin (Invitrogen, 15140122). 17. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea). 18. Gel-loading pipette tips. 19. Chip (Emulate, USA).

2.2.1 Coating and Injection of the MN Spheroid into an NMJ Chip

1. Sterile conical tubes 15 mL (SPL, Korea). 2. Ice. 3. Micropipette (Eppendorf, Germany).

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4. 10-μL pipette tip (Eppendorf, Germany). 5. 200-μL pipette tip (Eppendorf, Germany). 6. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea). 7. Tissue culture dishes, T75. 8. Gel-loading pipette tips. 9. Ham’s Nutrient Mixture F-12 (Ham’s F-12; Gibco™). 10. Reconstitution buffer (Nitta Gelatin). 11. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany).

3

Methods

3.1 Motor Neuron Spheroids Generation

1. Culture-induced pluripotent stem cell (iPSC) lines on irradiated mouse embryonic fibroblasts (MEFs) as described in the standard protocol at WiCell Feeder Based MEF Pluripotent Stem Cell Protocols [16]. 2. Dissociate iPSCs prepared in step 1 with 1 mg/mL dispase and split cells at a 1:6 ratio on irradiated MEFs-coated plates prepared in the previous step. 3. One day later, replace the media with Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12) and Neurobasal medium at 1:1, supplemented with 0.5 × N2, 0.5 × B27, 0.1 mM ascorbic acid, 1 × GlutaMAX and 1 × penicillin-streptomycin, 3 μM CHIR99021, 2 μM DMH-1, and 2 μM SB431542 and maintain them for 6 days. Media exchanges should be done every other day. 4. After 6 days, iPSCs differentiate into neuroepithelial progenitors (NEPs). 5. Dissociate NEPs with 1 mg/mL dispase and split cells at a 1:6 ratio on medium containing DMEM/F12 and Neurobasal medium at 1:1, supplemented with 0.5 × N2, 0.5 × B27, 0.1 mM ascorbic acid, 1 × GlutaMAX and 1 × penicillin-streptomycin, 1 μM CHIR99021, 2 μM DMH-1, 2 μM SB431542, 0.1 μM retinoic acid (RA), and 0.5 μM purmorphamine (Pur) and maintain them for 6 days. Media exchanges should be done every other day. 6. After 6 days, the NEPs differentiate into motor neuron progenitor cells (MNPCs). 7. Dissociate MNPCs with 1 mg/mL dispase and split cells at a 1: 6 ratio on medium DMEM/F12 and neurobasal medium at 1: 1, supplemented with 0.5 × N2, 0.5 × B27, 0.1 mM ascorbic acid, 1 × GlutaMAX and 1 × penicillin-streptomycin, 3 μM

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CHIR99021, 2 μM DMH-1, 2 μM SB431542, 0.1 μM RA, 0.5 μM Pur, and 0.5 mM valproic acid (VPA) once a week. 8. Transfer the cells to the cryovials containing DMEM/F12, 10% fetal bovine serum (FBS), and 10% dimethylsulfoxid (DMSO) so that each cryovial contains ~3–5 million cells. 9. Freeze the cryovials of the cells in liquid nitrogen tank (see Note 1). 10. Thaw cryovial of MNPCs in a 37 °C water bath. Decontaminate the cryovials by spraying and wiping the outside of the cryovials with 70% ethanol. 11. Transfer and culture the cells to a 15 mL conical tube containing 4 mL of DMEM/F-12 and 1× Antibiotic-Antimycotic (Anti-Anti). 12. Centrifuge the tubes at 1200 rpm for 3 min. 13. Aspirate the supernatant and work with pellet. 14. Transfer the cells to a T25 poly-L-ornithine (PLO)/laminincoated flask with 5 mL of MNPCs expansion medium containing 1:1 neurobasal medium: DMEM/F12, 1× Anti-Anti, 0.5× N2, 0.5× B-27, 0.5× GlutaMAX supplement, 100 μM AA, 3 μM CHIR-99021, 2 μM DMH-1, 2 μM SB431542, 0.1 μM RA, 0.5 μM Pur, 0.5 μM VPA, and 10 μM ROCK inhibitor. 15. Incubate the cells at 37 °C and 5% CO2 for 24 h. 16. Change the medium to MNPCs expansion without ROCK inhibitor. Media exchanges should be done every other day for 1 week. 17. Passage the cells and subculture the cells into two new T-25 PLO/laminin-coated flasks (see Note 2). 18. Transfer the cells into a PLO/laminin-coated T-25 flask, split them, and resuspend them in MNPCs expansion medium containing 10 μM ROCK inhibitor as previously described in step 14. 19. Reach the volume to 5 mL. 20. Incubate the cells at 37 °C and 5% CO2 for 24 h. 21. Change the medium to MNPCs expansion without ROCK inhibitor. Media exchanges should be done every other day for 1 week. 22. Add 2 mL of Accutase to medium to split the cells. Then, incubate the cells at 37 °C and 5% CO2 for 4–6 min. 23. Transfer the cells into 1 mL of induction and maturation medium containing 1:1 Neurobasal: DMEM/F12, 1× AntiAnti, 0.5× N-2, 0.5× B-27, 0.5× GlutaMAX supplement,

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100 μM AA, 0.5 μM RA, 0.1 μM Pur, 0.1 μM Compound E, and 10 ng/mL brain-derived neurotrophic factor (BDNF). 24. Count the cells by NucleoCounter® NC-100™. Prepare the sample to be counted by the NucleoCounter® NC-100™ according to the manufacturer’s instruction. 25. Replate cells into 96-well Clear Round Bottom Ultra-Low Attachment Microplate containing induction and maturation medium (5000 cells in 100 μL of medium) (see Note 3). 26. Centrifuge the cells at 1200 rpm for 5 min with special adapters. 27. Incubate the cells at 37 °C and 5% CO2. 28. MN spheroids will be formed. The MN spheroids should consist of at least 80% neuronal cells. 29. Every 2 weeks, add 50 μL of MN induction and maturation media to each well of the 96 U-bottom ultra-low attachment plate. Performing this step greatly increases the lifespan and durability of spheroids. Figure 1 shows all steps for the generation of motor neuron spheroids. 3.2 IPSC-Derived Skeletal Myoblasts Preparation and Injection into an NMJ Chip

1. Maintain the Human iPSC-derived skeletal myoblasts on fibronectin-coated T75 tissue culture dishes containing 10 μg/mL minimum essential medium–α (MEM-α) supplemented with 1 mM 8-bromo-cyclic AMP, 2 μM CHIR99021, 1 μM dorsomorphin, and 5% knockout serum replacement (KSR) for 1 day. 2. Fill gel injection port 1 and compartment 1 with 4% Pluronic F-127. 3. Incubate the microfluidic chip at 37 °C for 10 min. 4. Remove 4% Pluronic F-127. Wash the device with distilled water and DPBS and dry it (see Note 4). 5. Inject 200 μL of culture medium into the right medium reservoir with skeletal myocytes differentiation medium containing DMEM supplemented with 2% horse serum, 50 ng/mL human recombinant insulin-like growth factor-1 (IGF-1), and 1% penicillin-streptomycin. 6. After 24 h, skeletal muscle fibers form between pillar structures. 7. Skeletal myocyte differentiation medium should be injected to the reservoir up to day 14.

3.3 Coating and Injection of the Motor Neuron Spheroids into an NMJ Chip

1. The injection of MN spheroids starts 14 days after muscle cell culture and differentiation. Transfer an MN spheroid to a 15 mL conical tube on ice or on an ice block and mix the cells with cell culture medium through micropipette aspiration using a 200-μL pipette tip (see Note 5).

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Fig. 1 Generation of motor neuron spheroids

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2. Transfer the MN spheroid to a 15 mL conical tube containing 10 μL of Ham’s F-12 10× (Ham’s Nutrient Mixture F-12) using a 10-μL pipette tip. 3. Add 80 μL of collagen cell matrix type 1A to the mixture in the tube and mix them by thoroughly and gently pipetting up and down. 4. Add 10 μL of the reconstitution buffer to the mixture prepared in the previous step. 5. By using a 10-μL pipette tip, transfer the MN spheroid into the left compartment via gel injection port no. 2 with 50 μL of the gel mixture (see Note 6). 6. Incubate the chip at 37 °C for 10 min. 7. Add the maturation culture medium of nerve and muscle cells, respectively, in the reservoirs where the cells are present. 8. Incubate the device at 37 °C in a 5% CO2 for 2 weeks. 9. Media exchanges should be done every day for the first week and every 12 h in the second week. 10. It is suggested that Schwann cells, as progenitors or mature cells, are added to create a myelin sheath for MNs. In some embodiments, the Schwann cells are derived from cells of a patient with ALS. 11. Cells are ready for staining and subsequent processing. Figure 2 shows all steps for the preparation of iPSC-derived skeletal myoblasts and motor neuron spheroids and the injection of them into an NMJ chip.

4

Notes 1. Based on the cell shape and density, MNPCs can be kept in freezer up to passage 3 [17]. 2. One flask is used to hold the MNPCs stock and the other flask is used to produce MN spheroids. The MNPCs in two mentioned flasks will be dissociated by gentle cell dissociation reagent (GCDR) and Accutase, respectively [17]. 3. Note that the evaporation of the culture medium is much higher in the outer wells than in the inner wells. Therefore, use 200 μL of 1× PBS instead of seeding cells in outer wells [17]. 4. In case of washing with DPBS, the device should be incubated for 10 min at 37 °C [18].

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Fig. 2 Preparation of iPSC-derived skeletal myoblasts and motor neuron spheroids and the injection of them into an NMJ chip

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5. The use of ice prevents sudden polymerization of the gel. Pay attention to bring the reagents to the temperature of the mixture in the tube on the ice before use [18]. 6. Remove excess collagen from both compartments [18]. References 1. Ai J, Kiasat-Dolatabadi A, Ebrahimi-Barough S et al (2014) Polymeric scaffolds in neural tissue engineering: a review. Arch Neurosci 1(1): 15–20 2. Rahim F, Arjmand B (2017) Stem cell clinical trials for multiple sclerosis: the past, present and future. In: Neurological Regeneration. Springer, pp. 159–172 3. Moghaddam SA, Yousefi B, Sanooghi D et al (2017) Differentiation potential of human CD133 positive hematopoietic stem cells into motor neuron-like cells, in vitro. J Chem Neuroanat 86:35–40 4. Arjmand B, Roudsari PP, Alavi-Moghadam S et al (2021) Potential for stem cell-based therapy in the road of treatment for neurological disorders secondary to COVID-19. In: Regenerative engineering and translational medicine, pp. 1–15 5. Derakhshanrad N, Saberi H, Meybodi KT et al (2015) Case report: combination therapy with mesenchymal stem cells and granulocytecolony stimulating factor in a case of spinal cord injury. Basic Clin Neurosci 6(4):299 6. Masrori P, Van Damme P (2020) Amyotrophic lateral sclerosis: a clinical review. Eur J Neurol 27(10):1918–1929 7. van den Bos MA, Geevasinga N, Higashihara M et al (2019) Pathophysiology and diagnosis of ALS: insights from advances in neurophysiological techniques. Int J Mol Sci 20(11):2818 8. Eisen A (2021) The dying forward hypothesis of ALS: tracing its history. Brain Sci 11(3):300 9. Pandya VA, Patani R (2020) Decoding the relationship between ageing and amyotrophic lateral sclerosis: a cellular perspective. Brain 143(4):1057–1072

10. Bose P, Tremblay E, Maois C et al (2019) The novel small molecule TRVA242 stabilizes neuromuscular junction defects in multiple animal models of amyotrophic lateral sclerosis. Neurotherapeutics 16(4):1149–1166 11. Gois AM, Mendonc¸a DM, Freire MAM et al (2020) In vitro and in vivo models of amyotrophic lateral sclerosis: an updated overview. Brain Res Bull 159:32–43 12. Bonifacino T, Zerbo RA, Balbi M et al (2021) Nearly 30 years of animal models to study amyotrophic lateral sclerosis: a historical overview and future perspectives. Int J Mol Sci 22(22):12236 13. Katyal N, Govindarajan R (2017) Shortcomings in the current amyotrophic lateral sclerosis trials and potential solutions for improvement. Front Neurol 8:521 14. Leung CM, De Haan P, Ronaldson-Bouchard K et al (2022) A guide to the organ-on-a-chip. Nat Rev Methods Primers 2(1):1–29 15. Arjmand B, Kokabi Hamidpour S, Rabbani Z et al (2022) Organ on a chip: a novel in vitro biomimetic strategy in amyotrophic lateral sclerosis (ALS) modeling. Front Neurol 12:2541 16. https://www.wicell.org/ (2022) 17. Castellanos-Montiel MJ, Chaineau M, FrancoFlores AK et al (2022) An optimized workflow to generate and characterize iPSC-derived motor neuron (MN) spheroids. bioRxiv 18. Osaki T, Uzel SG, Kamm RD (2020) On-chip 3D neuromuscular model for drug screening and precision medicine in neuromuscular disease. Nat Protoc 15(2):421–449

Methods in Molecular Biology (2023) 2736: 151–161 DOI 10.1007/7651_2023_497 © Springer Science+Business Media, LLC 2023 Published online: 11 July 2023

Primary Human Leukemia Stem Cell (LSC) Isolation and Characterization Neslihan Meric¸ and Fatih Kocabas¸ Abstract Leukemia stem cells (LSC) are thought to be the basis of leukemia progression since they are highly resistant to conventional chemotherapy. LSC isolation is critical in experimental studies, drug development, and application. Due to their likely hematopoietic stem cell (HSC) origin, LSCs have surface antigens that are similar to HSC. Surface markers such as CD34, CD123, CD133, and CD33 have been used extensively to assess LSCs. LSCs could be separated from other cells using magnetic selection (MS) or flow cytometry selection (FCS) methods using these markers. Understanding the role of LSCs in cancer progression and how to therapeutically target them in vitro and in vivo is critical for the development of LSC-targeting drug candidates. In this chapter, we set out to describe the primary human LSC purification and characterization processes used on patient samples with leukemia and lymphoma. Key words Flow cytometry, Hematopoietic stem cells, Leukemia, Leukemia stem cells, Magnetic selection, Surface markers

1

Introduction Leukemia is a hematological malignant disease that is a lifethreatening condition for patients carrying this disease [1]. Acute and chronic cancers are classified as myeloid or lymphoid in origin. The mechanisms of acute and chronic leukemias differ, as do the therapeutic options. LSCs are distinguished by their quiet behavior, juvenile phenotype, particular metabolic properties, and dependency on certain signaling pathways [2]. Besides, LSCs have selfrenewal, drug resistance, and the ability to initiate leukemia when transferred to NOD/SCID mice [3, 4]. LSCs and HSCs share the same bone marrow (BM) niche [5] and both exhibit CD34+ CD38- staning. This supports the concept that LSCs are derived from HSCs and then undergo a sequence of malignant changes [6, 7]. The most noticeable marker of LCSs is the interleukin-3 receptor alpha chain (CD123), which is not abundantly expressed in CD34+/CD38- hematopoietic cells [8–10]. Furthermore, studies support the presence of LSCs in the CD34+/CD38population [3].

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While leukemia stem cells are well understood in AML, their existence and relevance in ALL are less clear [11]. Lapidot and colleagues discovered human AML-IC in immunodeficient SCID mice [12] in vivo [13] in 1994. Later research demonstrated that the NOD/SCID model was more successful [14]. LSCs, also known as leukemia-initiating cells (LIC) from time to time, are a subpopulation of leukemia cells that are distinguishable from other leukemia cells by having stem cell features [15]. Ng et al. conducted a study in which four AML patients were identified as CD34+CD38- phenotype and implanted into mice. LICs cells were found in transfected mice [16]. It was discovered that patient LSCs and LSCs extracted from transplanted xenografts were similar [17]. LSCs are seen in CD34+CD38- cells and a limited number of CD34- cells, but not in significant numbers of CD34+ cells [3]. Recent investigations have established that the prevalence of CD34+/CD38- LSCs correlates with MRD levels after chemotherapy, disease response, and poor survival [18]. Treatment of AML patients differs in the presence of LSCs, and resistance to treatment rises [17, 18]. In a larger gene analysis involving 1047 AML patients, the expression of fifty-two genes representing LSCs and non-LSCs was evaluated. It was discovered that patients with high-score LSCs had poor OS (overall survival) and DFS (diseasefree survival) [19]. Although xenograft transplantation tests verified the presence of LCSs in CD34+/CD38-cells, Taussig et al. discovered the presence of LSCs in samples from NPM-mutated AMLs with low CD34 expression [20, 21]. In some samples, LSCs were only found in CD34- cells, whereas in others, they were found in both CD34+ and CD34- cells, highlighting the heterogeneity of LSC phenotyping. Additionally, it has been shown that LSCs’ phenotype has changed and that their presence alters how CD34 and CD38 are expressed [17]. Different ratios of AML subtypes were found in LSCs, and it was found that the ratio of LSCs directly correlated with high-risk AML [19, 22].

2 2.1

Materials Instruments

Cytoflex S Flow Cytometry (Beckman, Cat. No # B47903, USA), S3E Cell Sorter (Life Science Research-Bio-Rad, Cat.No # 1451008), FACSAriaIII (BD, Cat. No # 23-11539-00), Hettich® ROTINA 38/38R Centrifuge (Sigma, Cat. No # Z720119), Centrifuge (Labogene, Cat. No # 416), BD Magnetic Bead System (EasySep™ Human CD34 Positive Selection Kit II, Cat.No # 17856).

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2.2 Bone Marrow and Blood Samples

BM and PB samples are taken from adult ALL and AML patients at initial diagnosis or at relapse. K2EDTA tubes are used to collect patient samples. Samples should be kept at 4 °C and processed as soon as possible.

2.3 Materials for Separating Mononuclear Cells (MNCs) from Bone Marrow-Peripheral Blood

BM-PB mononuclear cells (MNCs) are isolated by using Ficoll gradient centrifuge technique. Ficoll-Paque HistopaqueTM (Sigma, Cat No #10831). Plastic spray-coated K2EDTA Tube (BD, Cat No # 367835). Dulbecco’s phosphate buffer saline (DPBS) (Thermofisher Scientific, Cat No # 14190144). Hemocytometer (Neubauer counting chamber). BD FACS™ Lysing Solution (BD, Cat No # 349202). 12× 75 mm polystyrene round-bottom tube.

2.4 Separation Kits for LSC

EasySep Human CD34 Positive Selection Kit II (Stem Cell Technologies, Catalog #17856). EasySep Human FITC Positive Selection Kit (Stem Cell Technologies, Catalog # 17662). Apart from these, it is explained for which samples the selection kits are suitable (see Notes 1 and 2).

2.4.1 Recommended Medium

PBS containing 2% fetal bovine serum and 1 mM EDTA. Note that recommended medium does not contain Ca2+ and Mg2+.

2.5

Anti-human CD34 PE (Biolegend, Cat No # 343505). Anti-human CD33 FITC (Biolegend, Cat No # 303304). Anti-human CD123 FITC (Biolegend, Cat No # 306013). Anti-human CD133 APC (eBioscience, Cat No # 12-0349-42).

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Markers for LSCs

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3.1 Patient Bone Marrow and Blood Samples

After the patient’s consent and ethical approval from local ethics committees, bone marrow aspirates should be obtained from patients diagnosed with AL following appropriate immunophenotyping and other diagnostic tests that are performed in the clinic. LSC separation should be planned after initial quantification of the CD34+ cell number and blast rate. In patients diagnosed with AL, LSC separation is recommended with patient samples with 50% CD34+ cells or 40–50% blast (Fig. 1).

3.2 Ficoll Gradient Method for MNC Isolation

1. Dilute 5 mL of patient blood (up to 15 mL) in 1:1 ratio with Dulbecco’s phosphate-buffered saline (DPBS) in a 50 mL falcon tube and mix by gently inverting (Fig. 2).

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2. Place 5 mL Ficoll-Paque separation solution into another 50 mL falcon tube. 3. Overlap the Ficoll-Paque separation solution with the bone marrow/PBS mixture with care. 4. Centrifuge the tube at 3000 × g for 30 min at room temperature without brake. 5. Transfer carefully the cloudy interphase mononuclear cell layer to the new 50 mL falcon tube. The majority of these cells are

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lymphocytes, monocytes, thrombocytes, hematopoietic stem, and progenitor cells. 6. Wash MNCs with 3× DPBS and gently mix by inverting. The cells then need to be centrifuged for 5 min at 1800 rpm before the supernatant is removed. 7. Use a hemocytometer to count the cells after they have been suspended in 10 mL of DPBS. For the control group, apply the same processing to healthy samples. 3.3 Removal of Erythrocytes with Lysis Solution

If erythrocytes are still present in the detached MNCs (red color), erythrocytes should be removed with a lysis solution. A high erythrocyte ratio may cause difficulties in staining of MNCs with LSC markers for LSC separation. 1. Centrifuge cells suspended with DPBS at 1800 rpm for 5 min and then remove the supernatant. 2. Suspend in 500 μL of DPBS. 3. Add 5 mL of BD FACS™ Lysing Solution to the cell suspension and mix well. 4. Incubate at room temperature for 5 min. 5. Proceed with the sorting process after QC pass. 6. Regardless of the sorting process, firstly, samples are acquired in the S3E Cell Sorter device. To this end: (a) Open the dot plot with forward scatter (FSC) area and side scatter (SSC) area parameters (Fig. 3). (b) Generate an FSC area – FL2 area dot plot. Here, the FL2 area represents the CD34+ marker. (c) Take the R1 gate to exclude debris from the plot with FSC/SSC and cover all cells (see Fig. 3a). (d) Go to the FSC/FL1 plot and take the R2 gate from inside the R1 gate (see Fig. 3b). 7. Quantify initially UNS sample by inserting the tube into the device (the loading stage is brought to the run position). 8. Adjust detectors according to the LSC marker used. Select the cycle mode and determine target cell number, parameters, and threshold values. We recommend accusation and sorting of hematopoietic stem cell samples at +4 °C. 9. Mark the CD34 negative cell population based on the UNS sample. 10. Do acquisition of the CD34+ stained tube with the same voltage and instrument settings.

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Fig. 3 CD34+ LSC gating strategy on S3E Cell Sorter (a) Gating from the entire leukocyte population in the FSC/SSC plot (= size/granularity) excluding debris. (b) Gating on CD34+LSCs as a marker for CD34+activity, in fluorescence channel FL1

11. After the acquisition of the CD34+ stained sample, observe the CD34+ cell group above the gate from which the negative cell group was taken (see Fig. 3b). 12. Make necessary adjustments in the S3eTM Cell Sorter device to start the sorting process. 13. After the desired cell group door is taken, right-click on the door and determine the sort direction. 14. Click on the sort logic button and determine the collection vessel. Generally, a 5 mL collection tube is used. 15. Select the sort mode (purity, enrich, or single) and event limit (100.000). Purity mode is preferred in CD34+ cell sorting. 16. Determine the volume of the sort collection tube (0.50 mL) and place the collection tube (12 × 75 mm polystyrene round bottom tube containing 0.50 mL RPMI medium) in the first or second compartment of the collection channel. 17. Close the sort chamber door and press on the start sorting button on the control panel. Then, sort statistics screen that appears and sorting starts. 3.3.1 Immunophenotyping After Sorting

Before sorting, MNC cells are stained with LSC markers. After sorting, LSCs are stained with LSC markers and compared before and after (see Figs. 4–6). It is recommended to use other LSC markers together with CD34+ marker when performing LSC characterization (see Note 3).

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To separate CD34+ cells from other samples, including freshly mobilized peripheral blood, bone marrow mononuclear cells, previously frozen cord blood mononuclear cells, human embryonic stem (ES), and induced pluripotent stem (iPS) cell cultures, use the “EasySep Human CD34 Positive Selection Kit II.” 1. Isolated LSCs can be used for cell culture in vitro, xenotransplantation in vivo, and molecular analysis. We describe the use of the freshly mobilized peripheral blood and bone marrow mononuclear cells in this protocol (EasySep Human Cord Blood CD34+ Selection Kit II (Stem Cell Technologies, Cat No#17896) 2. Isolate MNCs with Ficoll gradient centrifugation method and lyse erythrocytes as described in Subheadings 3.2. and 3.3. 3. Resuspend cells with medium according to cell concentrations. (We recommended PBS containing 2% fetal bovine serum and 1 mM EDTA). For 1 mL, add medium to make up to 10 mL. Then, the cell suspension needs to be pipetted up and down 2 or 3 times. 9. Place the tube containing the cell suspension on the magnet and incubate for 10 min at RT.

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10. Remove the supernatant in the tube and discard at once with a 10 mL serological pipette. Then, isolate the cells remaining in the tube. Then, carefully remove the tube from the magnet. 11. If the cell suspension is ≤1 mL, add medium to make up to 3 mL. If the cell suspension is >1 mL, add medium to make up to 10 mL. Then, the cell suspension needs to be pipetted up and down 2 or 3 times. 12. Place the tube containing the cell suspension on the magnet and incubate for 5 min at room temperate. 13. Remove the supernatant in the tube and discard at once with a 10 mL serological pipette. Remove carefully the tube from the magnet. 14. Repeat steps 10–12. 15. Remove the tube from the magnet. Suspend cells in the medium. Centrifuge at 300 × g for 10 min at brake low. Then, remove the supernatant. 16. Resuspend the pellet with the desired medium. Isolated LSCs are now ready to use.

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3.3.3 Immunophenotyping After MS

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Before MS, MNC cells are stained with LSC markers. After MS, LSCs are stained with LSC markers and compared before and after (see Figs. 4–6). When CD34+ isolation is performed using the MS method, it is observed that CD34+ cells are isolated together with other LSC cells.

Notes 1. “EasySep Human Cord Blood CD34+ Selection Kit II (Stem Cell Technologies, Cat No#17896)” is used to obtain high-purity CD34+ from a fresh whole umbilical cord blood sample. 2. Use the complete kit for “EasySep Human Whole Blood CD34+ Cells (Stem Cell Technologies, Catalog #15086)” to isolate CD34+ cells from fresh blood or buffy coat. 3. While performing immunophenotyping with a flow cytometer, we added CD34 antibodies next to LSC markers. Since CD34+ cells are the main identifying marker of LSCs, it is recommended to evaluate their positivity with other LSC markers.

Acknowledgments This study was supported by funds provided by Gilead Sciences International Hematology and Oncology program. NM and FK were supported by “Gilead ile Hayat Bulan Fikirler.” References 1. Juliusson G, Hough R (2016) Leukemia. pp 87–100 2. Polak A, Bialopiotrowicz E, Krzymieniewska B et al (2020) SYK inhibition targets acute myeloid leukemia stem cells by blocking their oxidative metabolism. Cell Death Dis 11:956 3. Hanekamp D, Cloos J, Schuurhuis GJ (2017) Leukemic stem cells: identification and clinical application. Int J Hematol 105(5):549–557 4. Gupta PB, Chaffer CL, Weinberg RA (2009) Cancer stem cells: Mirage or reality? Nat Med 15(9):1010–1012 5. Vormoor J, Lapidot T, Pflumio F et al (1994) Immature human cord blood progenitors engraft and proliferate to high levels in severe combined immunodeficient mice. Blood 83(9):2489–2497 6. Weissman I (2005) Stem cell research: Paths to cancer therapies and regenerative medicine. J Am Med Assoc 294(11):1359–1366

7. Passegue´ E, Wagers AJ, Giuriato S et al (2005) Global analysis of proliferation and cell cycle gene expression in the regulation of hematopoietic stem and progenitor cell fates. J Exp Med 202(11):1599–1611 8. Okada Y, Feng Q, Lin Y et al (2005) hDOT1L links histone methylation to leukemogenesis. Cell 121(2):167–178 9. Wang J, Iwasaki H, Krivtsov A et al (2005) Conditional MLL-CBP targets GMP and models therapy-related myeloproliferative disease. EMBO J 24(2):368–381 10. Jordan CT, Upchurch D, Szilvassy SJ et al (2000) The interleukin-3 receptor alpha chain is a unique marker for human acute myelogenous leukemia stems cells. Leukemia 14:1777– 1784 11. Bernt KM, Armstrong SA (2009) Leukemia stem cells and human acute lymphoblastic leukemia. Semin Hematol 46(1):33–38

Primary Human LSC Isolation and Characterization 12. Shultz LD, Ishikawa F, Greiner DL (2007) Humanized mice in translational biomedical research. Nat Rev Immunol 7:118–130 13. Lapidot T, Sirard C, Vormoor J et al (1994) A cell initiating human acute myeloid leukaemia after transplantation into SCID mice. Nature 367(6464):645 14. Bonnet D, Dick JE (1997) Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat Med 3(7):730–737 15. Chung SS, Park CY (2014) Acute myeloid leukemia stem cells-updates and controversies. Cancer Stem Cells:143–160 16. Ng SWK, Mitchell A, Kennedy JA et al (2016) A 17-gene stemness score for rapid determination of risk in acute leukaemia. Nature 540: 433–437 17. Buss EC, Ho AD (2011) Leukemia stem cells. Int J Cancer 129(10):2328–2336 18. Terwijn M, Zeijlemaker W, Kelder A et al (2014) Leukemic stem cell frequency: a strong

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biomarker for clinical outcome in acute myeloid leukemia. PLoS One 9(9):e107587 19. Gentles AJ, Plevritis SK, Page P et al (2010) Association of a leukemic stem cell gene in acute myeloid leukemia. JAMA 304(24): 2706–2715 20. Jin L, Lee EM, Ramshaw HS et al (2009) Monoclonal antibody-mediated targeting of CD123, IL-3 receptor α chain, eliminates human acute myeloid leukemic stem cells. Cell Stem Cell 5(1):31–42 21. Lechman ER, Gentner B, Ng SWK et al (2016) MiR-126 regulates distinct self-renewal outcomes in normal and malignant hematopoietic stem cells. Cancer Cell 29(2):214–228 22. Taussig DC, Vargaftig J, Miraki-Moud F et al (2010) Leukemia-initiating cells from some acute myeloid leukemia patients with mutated nucleophosmin reside in the CD34- fraction. Blood 115(10):1976–1984

Methods in Molecular Biology (2023) 2736: 163–176 DOI 10.1007/7651_2022_467 © Springer Science+Business Media, LLC 2023 Published online: 15 December 2022

GMP-Compliant Mesenchymal Stem Cell-Derived Exosomes for Cell-Free Therapy in Cancer Babak Arjmand, Sepideh Alavi-Moghadam, Mostafa Rezaei-Tavirani, Shayesteh Kokabi-Hamidpour, Rasta Arjmand, Kambiz Gilany, Mohsen Rajaeinejad, Fakher Rahim, Nazli Namazi, and Bagher Larijani Abstract Cancer is categorized as one of the life-threatening disease in the world, which has recently been associated with a significant increase in the incidence and prevalence rate. Hence, the discovery of effective approaches for prevention, early diagnosis, and effective treatment for cancer has been prioritized by oncology researchers. In recent decades, mesenchymal stem cells show great potential to advance the field of regenerative medicine and oncology research due to representing prominent characteristics. Recently, studies indicate that mesenchymal stem cells can play an important role by secreting extracellular vesicles like exosomes in modulating the biological functions of target cells through paracrine regulation. Indeed, the exosomes derived from mesenchymal stem cells can represent the same therapeutic potential as parent cells with fewer side effects. Therefore, it can be demonstrated that exosomes can be a suitable drug delivery candidate in regenerative medicine and targeted therapy. It is also noteworthy that as the use of exosome therapy becomes more common in clinical studies, the importance of improving basic criteria such as safety, efficiency, and quality of stem cell products will also be highlighted. Based on this concept, the good manufacturing practice principles were put forward to examine the standard of cell products from different qualitative and quantitative aspects to progress the cell therapy. In other words, the principles of good manufacturing practice should be observed not only in the extraction and isolation of stem cells but also in the extraction of products related to stem cells such as exosomes in the field of treatment. Key words Exosomes, Extracellular vesicles, Mesenchymal stem cell, Mesenchymal stem cell-derived exosomes, Regenerative medicine

Abbreviations BSA CCM CD CTRM DMEM-LG ECL EDQM ELISA

Bovine serum albumin Cell culture conditioned media Cluster of differentiation Cell therapy and regenerative medicine Dulbecco Modified Eagle medium-low glucose Enhanced chemiluminescence European Directorate for the Quality of Medicine & Healthcare Enzyme-linked immunosorbent assay

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EV FBS GMP hPLMSCs HRP MNCs MSC-EXO MSCs PBS PCR PVDF QA RNA RT SDS-PAGE SOP TBST TEM TSEs

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Extracellular vesicle Fetal bovine serum Good manufacturing practice Human placenta-derived mesenchymal stem cells Horseradish peroxidase Mononuclear cells Mesenchymal stem cell-derived exosome Mesenchymal stem cells Phosphate-buffered saline Polymerase chain reaction Polyvinylidene difluoride Quality assurance Ribonucleic acid Room temperature Sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) Standard operating procedure Tris-buffered saline with 0.1% Tween® 20 detergent Transmission electron microscope Transmissible spongiform encephalopathies

Introduction Cancer is the name given to a wide range of diseases which can affect any part of the body due to uncontrolled abnormal cell division and growth. Since the cancer can be associated with the onset of dangerous disease state, it has been classified as one of dangerous and life-threatening disease worldwide, which has also been accompanied by the rise of the number of patients in recent years. Therefore, the increase in the incidence and prevalence rate of cancer has been an alarm for researchers to search for effective diagnosis and therapeutic approaches to eradicate or curb the upcoming rise in cancer incidence rate as quickly and effectively as possible [1, 2]. Over the past years, the application of various types of stem cells, like mesenchymal stem cells (MSCs), has become an interesting and promising scientific topic in the field of oncology research [3]. In the field of biological sciences, MSCs are defined as a subset of non-hematopoietic and multipotent stromal cells that originate from different tissues such as bone marrow, adipose tissue, dental pulp, etc. [4, 5]. Functionally, these types of adult stem cells are endowed with outstanding features, such as self-renewal, multilineage differentiation, hypo immunogenic function, immunomodulation, and tissue regeneration, which have made MSCs top-used cells with high therapeutic potential in different areas of cell therapy and regenerative medicine (CTRM) realm [4–6], like chronic autoimmune diseases [7], spinal cord injuries [8], diabetes [9, 10], and wound healing [11, 12]. Over the past years, studies

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have revealed that MSCs also have paracrine properties, which have a great contribution to modulating biological processes such as angiogenesis, apoptosis, mitosis, and inflammation by releasing soluble factors like exosomes [13]. Biologically, exosomes are a subtype of extracellular vesicles, which participate in the regulation of intercellular communications [14]. These nanoscale extracellular vesicles are released by different cell types, present in almost all body fluids, and responsible for transferring different cargos of biological macromolecules including ribonucleic acid (RNA)s, lipids, and proteins with the intention to impact on target cell’s function. They can, therefore, affect the biological processes of target cells as mediators, which is why many scientists have become interested in exploring and exploiting the function of these vesicles in different fields of medicine. In addition to the widespread presence of these vesicles in human body fluids and their biological cargos, exosomes represent some surface markers (e.g., CD9, CD63, and CD81), which can be identified and analyzed by employing different biochemistry and molecular biology techniques. Accordingly, exosomes not only can have a crucial role in the regulation of other cells’ fate but also pave the way for accurate diagnosis of various human diseases [15]. In recent years, the concept of MSC-derived exosomes has been met with enthusiasm by researchers in the regenerative medicine field due to representing both characteristics of exosomes and MSCs by these extracellular vesicles. Indeed, MSC-derived exosomes are biocompatible and don’t provoke an immune response due to their acellular structure. Additionally, these cells can resist or evade chemical or enzymatic destruction and immune system surveillance. Therefore, they can have a long-circulating half-life. Also, they are easily able to penetrate into the blood–brain barrier due to their small size. Moreover, in combination with nanoparticles, they can not only have the same therapeutic effects as MSCs but also can have fewer side effects compared to the parent cells. Accordingly, it can be concluded that the outstanding features of MSC-derived exosomes have caused them to be considered a suitable candidate therapeutic strategy in medical studies (Fig. 1) [18], which has been proven by studies established for the treatment of diseases, such as wound healing [19], cardiac regeneration [20], liver [21], and kidney disease [22]. It is worth mentioning that the rise of MSCs application, as well as the use of the exosome therapy approach in the field of regenerative medicine and targeted drug delivery, highlights the need to increase and enhance good product criteria such as safety, efficiency, and quality of products based on stem cells [23]. Therefore, the requirement to consider good manufacturing practice (GMP) guidelines becomes prominent in this field, which can help to pave the way for the development of potential therapies from basic science to clinical applications [24–26]. Hereupon, according to the advantages and outstanding features of MSCs

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Fig. 1 The application of exosomes in medicine. Exosomes can play an important role both diagnosis and treatment of cancer. Exosomes naturally have a great contribution to progress various biological processes. However, in the cancerous state, cells use exosomes to advance their aggressive goals. Since exosomes are present in almost all body fluids, they can be used as biomarkers. Therefore, different types of cancers can be diagnosed by analysis of exosomes in body fluids [16]. In order to treat cancer, therapeutic agents can be loaded on exosomes. Hence, exosomes can also act as a drug carrier to target cancer cells and prevent the development and progression of tumors [17]

and the important role of exosomes derived from MSCs in regenerative medicine and oncology research, this paper will investigate the method of isolation of exosomes from MSCs based on the principles of GMP.

2 2.1

Materials Cell Culture

1. Human placenta-derived mesenchymal stem cells (hPLMSCs). 2. 70% ethanol (Kimia Alcohol Zanjan, Iran). 3. Phosphate-buffered saline (PBS; Biowest, France). 4. Dulbecco Modified Eagle medium-low glucose (DMEM-LG; Biowest, France). 5. Fetal bovine serum (FBS) Biopharm—EDQM certified (Biowest, USA) (see Note 1). 6. TrypLE™ Select (Thermo Fisher Scientific, USA) (see Note 2). 7. Sterile serological pipettes 5, 10, and 25 mL (SPL, Korea).

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8. Sterile conical tubes 15 and 50 mL (SPL, Korea). 9. Filter cap cell culture flasks (300, 175, 75, and 25 cm2 (TPP, Switzerland)). 10. Clean room (GMP) facility (see Note 3). 11. Biological safety cabinet (Esco, Singapore) (see Note 4). 12. Inverted microscope with phase-contrast (Nikon, Japan). 13. Hemocytometer and cover glass. 14. Trypan blue solution 0.4% (Invitrogen, USA). 15. NucleoCassette™ (Chemometec, Denmark). 16. NucleoCounter® NC-100™ (Chemometec, Denmark). 17. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 2.2 Cell Culture Conditioned Media (CCM) Preparation

1. Clean room (GMP) facility. 2. Biological safety cabinet (Esco, Singapore). 3. Sterile Pasteur pipettes 3 mL (Sigma, USA). 4. Cultured (hPLMSCs) at relatively high density (104 cells/ cm2). 5. Serum-free medium for hMSCs (StemPro MSC SFM)(Gibco, USA). 6. Sterile conical tubes 15 and 50 mL (SPL, Korea). 7. Refrigerated centrifuge (swing-out rotor with buckets for 50 and 15 mL tubes) (Hettich, Germany). 8. CO2 incubator (set at 5% CO2, 37 °C, and 95% relative humidity) (Memmert, Germany). 9. Refrigerated centrifuge (swing-out rotor with buckets for 50 and 15 mL tubes) (Hettich, Germany). 10. 0.2 μm sterile syringe filter (Corning, USA). 11. Ultralow temperature freezer (New Brunswick Eppendorf, USA).

2.3 Isolation of Mesenchymal Stem Cell-Derived Exosomes (MSC-Exo)

1. Clean room (GMP) facility. 2. Biological safety cabinet (Esco, Singapore). 3. Sterile Pasteur pipettes 3 mL (Sigma, USA). 4. MSC conditioned media. 5. Ultra-centrifuge (Eppendorf, Germany). 6. Sterile conical tubes 15 and 50 mL (SPL, Korea). 7. Phosphate-buffered saline (PBS; Biowest, France). 8. Ultralow temperature freezer (New Brunswick Eppendorf, USA).

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2.4 Identification of Mesenchymal Stem Cell-Derived Exosomes (MSC-Exo)

1. MSC-Exo suspension.

2.4.1 Scanning Electron Microscopy Photography by Transmission Electron Microscope (TEM)

5. Filter paper.

2.4.2 Western Blot

1. MSC-Exo suspension.

2. Pipette tips: crystalline, yellow, and blue (TPP, Switzerland). 3. Glutaraldehyde solution of 0.1% (Merck, Germany). 4. Carbon-coated TEM grid (Sigma, USA). 6. Uaranyl acetatecetate (Sigma, USA). 7. Transmission electron microscope (TEM; JEM-1400 Plus, JEOL Ltd., Tokyo, Japan).

2. Pipette tips: crystalline, yellow, and blue (TPP, Switzerland). 3. RIPA buffer (2×) (Sigma, USA). 4. Sodium dodecyl–sulfate polyacrylamide gel electrophoresis (SDS-PAGE) protein loading buffer (5×). 5. BCA assay kit (Beyotime, P0010). 6. 0.22 μm polyvinylidene difluoride (PVDF) membranes. 7. Blocking buffer (5% skimmed milk in TBST). 8. Primary antibodies (1:1000) CD9 (1:200; sc-59140, Santa Cruz Biotechnology, TX, USA), CD63 (1:200; sc-5275, Santa Cruz Biotechnology), or CD81 (1:400; sc-166029, Santa Cruz Biotechnology)). 9. Tris-buffered saline with 0.1% Tween® 20 detergent (TBST). 10. Goat anti-mouse horseradish peroxidase (HRP) secondary antibody (1:3000). 11. Enhanced chemiluminescence (ECL) Select Western Blotting Detection Reagent (GE Healthcare UK Ltd., Buckinghamshire, UK). 12. Image Quant LAS 4000 mini (GE Healthcare). 2.4.3 Enzyme-Linked Immunosorbent Assay (ELISA)

1. MSC-Exo suspension. 2. Pipette tips: crystalline, yellow, and blue (TPP, Switzerland). 3. Microcentrifuge tubes. 4. Refrigerated microcentrifuge (Eppendorf, Germany). 5. Polyclonal antibodies directed against extracellular vesicle (EV)/exosome antigens (capture antibodies) (e.g., anti-CD9 (cluster of differentiation 9) antibody clone H110; anti-CD81 (cluster of differentiation 81) antibody clone H-121 and clone PA5-79003). 6. Monoclonal antibodies against EV/exosome antigens (detection antibodies) non-interfering with polyclonal antibodies

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[e.g., anti-CD63 (cluster of differentiation 63) antibody clone H5C6]. 7. Deionized water. 8. Carbonate buffer (pH 9.6). 9. Blocking buffer (0.5% bovine serum albumin (BSA) in PBS). 10. PBST (PBS + 0.1%Tween20). 11. Phosphate-buffered saline (PBS; Biowest, France). 12. 96-well plate with highly charged polystyrene surface suitable for antibodies attachment (SPL, Korea). 13. Micro-titer plate shaker (Scilogex SCI, China). 14. Vortex (Eppendorf, Germany). 15. ELISA sealing film or parafilm (Sigma, USA). 16. Stat Fax 4200- Microplate Reader (Vesta Tajhiz Part, Iran). 17. Blue POD substrate(Sigma, USA). 18. Humidified chamber or incubator at 37 °C (Memmert, Germany).

3 3.1

Methods Cell Culture

1. It is necessary to carry out each of these processes in the cleanroom and under the biological safety cabinet. 2. The hPLMSCs must be isolated from human placenta based on our pervious protocol [25]. 3. Each 10–15 × 106 mononuclear cells (MNCs) must be seeded into a T75 culture flask. 4. As a complete culture media, use DMEM-LG supplemented with 10% FBS Biopharm. 5. Put the culture flasks in an incubator (37 °C, 5% CO2, and 95% humidity). 6. Check the flasks for contamination and visiting spindle-shaped PLMSCs after 24-h (see Notes 5 and 6) (Fig. 2). 7. Every 3 days, take the media out and replace it with new cultural media. 8. Remove the media and wash it with pre-warmed PBS when PLMSCs are between 80% and 90% confluent. 9. Next, add TrypLE™ Select (2–3 mL for a flask measuring 75 cm2, and adjust the volume for other flasks based on their surface) to facilitate cell dissociation. 10. Check the flasks for cell detachment after 5–10 min of CO2 incubator incubation.

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Fig. 2 Microscopic appearance of PLMSCs at primary culture (×10)

11. Add an equivalent volume of PBS after the cell dissociation is complete; then transfer the cell suspension to a 50 mL conical tube(s). 12. Centrifuge at 300 g for 5 min. 13. Resuspend the cell pellet in the appropriate volume of complete culture media after carefully discarding the supernatant. 14. Use the NucleoCounter equipment to determine the number of viable cells. 15. In order to double-check, utilize a hemocytometer and trypan blue 0.4% solution (1:1) to count PLMSCs, and assess the viability of the cells. 16. Record each parameter in accordance with the standard operating procedure (SOP) (see Note 7). 17. Subculture the PLMSCS at a density of 104 cells/cm2 in the new cell culture flasks. 18. Place the flasks in to the CO2 incubator (37 C, 5% CO2, humidified), and conduct the cell expansion process until enough PLMSCs are produced to continue the research (usually 3–6 subcultures). 19. Utilize the right techniques to characterize and identify MSCs (see Note 8). 3.2 Cell Culture Conditioned Media (CCM) Preparation

1. It is necessary to carry out each of these processes in the cleanroom and under the biological safety cabinet. 2. Replace the media with serum-free DMEM when PLMSCs are between 80% and 90% confluent and culture them for 48-h. 3. Then, collect the cell supernatants.

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4. To remove cell debris and apoptotic bodies, centrifuge consecutively at 300 g for 10 min and 10,000 g for 20 min at 4 °C, and carefully remove the supernatant. 5. Finally, filter the supernatant through a 0.22 μm filter sterilizer (the collected medium was defined as hPLMSC cultured conditioned medium (hPLMSC-CM). 6. Store in - 80 °C freezer for further analysis. 3.3 Isolation of Mesenchymal Stem Cell-Derived Exosomes (MSC-Exo)

1. Ultracentrifuge hPLMSC-CM at 100,000 g for 70 min at 4 °C. 2. Rinse precipitate with PBS. 3. By performing a second round of ultracentrifugation at 100,000 g for 70 min at 4 °C, high purity exosomes can be obtained. 4. Resuspend the pelleted (MSC-Exo) and store it at -80 °C for further analysis.

3.4 Identification of Mesenchymal Stem Cell-Derived Exosomes (MSC-Exo) 3.4.1 Scanning Electron Microscopy Photography by Transmission Electron Microscope (TEM) 3.4.2 Western Blot

1. Pipette a drop of MSC-Exo suspension (10 μL) onto a carboncoated grid and allow to adsorb to the grid for 1 min. 2. Use filter paper to remove the excess exosomes. 3. For negative staining, 1% of uranyl acetate (10 μL) add dropwise to the grid for 1 min, and allow to dry for 10 min at room temperature (RT). 4. Use high-resolution transmission electron (HRTEM) at 80–120 kv for TEM examination.

microscope

1. Dilute MSC-Exo at a ratio of 1:1 with protein loading buffer (2×). 2. Separate in a 10–20% gradient SDS-PAGE gel. 3. Transfer proteins to PVDF membranes and block with blocking buffer. 4. Incubate the membranes for an overnight with the primary antibody CD9, CD63, or CD81 at 4 °C. 5. Then, wash them with TBST three times. 6. Incubate the membranes for an hour with goat anti-mouse HRP secondary antibody (1:3000) at RT. 7. Detect the bands using ECL Select Western Blotting Detection Reagent (GE Healthcare UK Ltd., Buckinghamshire, UK), and capture images using an Image Quant LAS 4000 mini (GE Healthcare).

3.4.3 Enzyme-Linked Immunosorbent Assay (ELISA)

1. Coat 4 μg/mL polyclonal antibodies (capture antibodies) to the plate (96-well plate) in a volume of 100 L/well of carbonate buffer.

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2. Shake the plate for 30 min and then place the plate overnight at 4 °C. 3. With 300 μL of PBS, wash the plate three times. 4. Add 100 μL blocking buffer and then incubate for 1-h at RT. 5. Wash the plate three times with PBS. 6. Coat the plate with 50 μL MSC-Exo. 7. Cover the plate with parafilm, and place it on shaker for 15–20 min, and incubate it at 37 °C overnight. 8. Wash three times with 300 μL of PBST and once with PBS. 9. Add 4 μg/mL of monoclonal detection primary antibodies, and then incubate for 1 h at 37 °C. 10. Wash three times with 300 μL of PBST and once with PBS. 11. Add 100 μL secondary anti-mouse-HRP (dilution 1:50,000), and then incubate for 1 h at 37 °C. 12. Wash three times with PBST. 13. Add 50 μL POD and incubate in dark 15 min at RT. 14. To complete the reaction, add 50 μL of 4 N H2SO4. 15. Use a microplate spectrophotometer reader, to determine the optical density at 450 nm. Figure 3 shows all these steps schematically.

4

Notes 1. Regulatory agencies around the world are extremely concerned about the possibility of prions being transmitted through the use of therapeutic biologicals. FBS, which is still frequently used in the pharmaceutical and biotechnology sectors as a supplement to cell culture media in the expansion of stem cells for regenerative medicine actions, needs to pass rigorous inspection. In others words, utilizing a qualified FBS (with complete traceability) is actually recommended for GMP-grade cellular product development. The certified FBS free of transmissible spongiform encephalopathies (TSEs) are listed on the European Directorate for the Quality of Medicine & Healthcare (EDQM) website (which is continually updated). FBS Biopharm (from Biowest Company) has been utilized for serum supplementation in this protocol. The company has recommended using this EDQM-certified FBS for biopharmaceutical manufacturing and other uses that need for the greatest level of product quality and documentation [26–28].

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Fig. 3 Procedure for extracting exosomes from mesenchymal stem cells. In step one, the mesenchymal stem cells (MSCs) are derived from human placenta. Cells are seeded into flasks. The flasks are put in the incubator. The sample is centrifuged and re-suspended. After several steps, the cells are counted by hemocytometer.

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2. Clinical applications of regenerative medicine need the use of xeno-free workflows. Herein, a variety of adherent mammalian cells can be separated using the recombinant enzyme TrypLETM Select, which is made in accordance with GMP guidelines and is free of animal products [25, 29]. 3. For executing GMP-based and clinical-grade production, clean room facilities, which are enclosed areas with precisely controlled temperatures, humidity levels, air pressures, and air particle concentrations, are essential [30–32]. 4. The most common use of a biological safety cabinet, also known as a biosafety cabinet, is for managing pathogenic biological samples or for applications that demand a sterile work area [33]. 5. The earliest feasible microbial contamination detection is required. The type of the microbe will determine the detection techniques. They include biological testing, polymerase chain reaction (PCR), chemical or fluorescent staining, optical microscopy, turbidimetry, pH measurements, or a simple visual assessment. Typically, optical microscopy can be used to identify bacteria and fungi. By appearing turbid or spotty, the contaminated cultures can be seen by the naked eye as early as over the weekend due to their rapid development rate. Consequently, testing kits can be used to identify these microorganisms. Mycoplasma cannot be seen by the naked eye or even by optical microscopy in cell cultures. Therefore, these organisms can remain undetected for extended periods of time and are discovered utilizing specific assays [34–36]. 6. MSCs appeared as elongated and aligned spindle-shaped or fibroblast-like morphologies when they are seeded on smooth surfaces [37]. 7. An organization’s routine or repeated action is documented by a set of written instructions called a SOP. A successful quality system must include the development and use of SOPs because they give people the knowledge they need to do their jobs well ä Fig. 3 (continued) Cells are subcultured and put in an incubator. In step two, the sample is centrifuged to remove cell debris and apoptotic bodies after collecting the cell supernatants. The supernatant is filtered and stored in freezer. In step three, the sample is isolated by ultracentrifuge and rinsed with PBS. The exosomes are obtained by the second round of ultracentrifugation. The sample is resuspended and stored in freezer. In step four, the electron microscopy photography by transmission electron microscope (TEM), Western blotting, and enzyme-linked immunosorbent assay (ELISA) are used to identify the of MSC-Exo. In the final step, the MSC-Exo are characterized by applying karyotyping, flow cytometry, and staining techniques. Abbreviations: CCM cell culture conditioned media; ELISA enzyme-linked immunosorbent assay; hPLMSCs human placentaderived mesenchymal stem cells; MSC-Exo mesenchymal stem cell-derived exosomes; TEM transmission electron microscope

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and enable consistency in the quality and integrity of a product or end-result. In other words, the operational components of an organization that would be governed by a work plan or a Quality Assurance (QA) Project Plan are described in SOPs in both technical and fundamental programmatic operational terms [38]. 8. For characterization, we can check the immunophenotypes (CD markers) using flow cytometry (MSCs can express cell surface markers such as CD44, CD105, and CD90) and examine a cell’s ability to differentiate in vitro into osteoblasts, adipocytes, and chondrocytes, checking adhesion to culture flasks, and we can also check the cellular karyotype [25]. References 1. Arjmand B, Hamidpour SK, Alavi-Moghadam S et al (2022) Molecular docking as a therapeutic approach for targeting cancer stem cell metabolic processes. Front Pharmacol 13 2. Arjmand B, Hamidpour SK, Tayanloo-Beik A et al (2022) Machine learning: a new prospect in multi-omics data analysis of cancer. Front Genet 13:824451 3. Rahim F, Arjmand B, Larijani B et al (2018) Stem cells treatment to combat Cancer and genetic disease: from stem cell therapy to gene-editing correction. In: Stem cells for cancer and genetic disease treatment, pp. 29–59. Springer 4. Arjmand B, Goodarzi P, Aghayan HR et al (2019) Co-transplantation of human fetal mesenchymal and hematopoietic stem cells in type 1 diabetic mice model. Front Endocrinol 10: 761 5. Larijani B, Aghayan H, Goodarzi P et al (2015) Clinical grade human adipose tissue-derived mesenchymal stem cell banking. Acta Med Iran 53:540–546 6. Gilany K, Masroor MJ, Minai-Tehrani A et al (2019). Metabolic profiling of the mesenchymal stem cells’ secretome. In: Genomics, proteomics, and metabolomics, pp. 67–81. Springer 7. Abedi M, Alavi-Moghadam S, Payab M et al (2020) Mesenchymal stem cell as a novel approach to systemic sclerosis; current status and future perspectives. Cell Regen 9(1):1–19 8. Derakhshanrad N, Saberi H, Meybodi KT et al (2015) Case report: combination therapy with mesenchymal stem cells and granulocytecolony stimulating factor in a case of spinal cord injury. Basic Clin Neurosci 6(4):299 9. Larijani B, Arjmand B, Ahmadbeigi N et al (2015) A simple and cost-effective method for

isolation and expansion of human fetal pancreas derived mesenchymal stem cells. Arch Iran Med 18(11) 10. Madani S, Setudeh A, Aghayan HR et al (2021) Placenta derived mesenchymal stem cells transplantation in type 1 diabetes: preliminary report of phase 1 clinical trial. J Diabetes Metab Disord 20(2):1179–1189 11. Aghayan HR, Hosseini MS, Gholami M et al (2022) Mesenchymal stem cells’ seeded amniotic membrane as a tissue-engineered dressing for wound healing. Drug Deliv Transl Res 12(3):538–549 12. Goodarzi P, Alavi-Moghadam A, Sarvari M et al (2018) Adipose tissue-derived stromal cells for wound healing. In: Cell biology and translational medicine, volume 4, pp. 133–149. Springer 13. Pankajakshan D, Agrawal DK (2014) Mesenchymal stem cell paracrine factors in vascular repair and regeneration. J Biomed Technol Res 1(1):9 14. Goodarzi P, Alavi-Moghadam S, Payab M et al (2019) Metabolomics analysis of mesenchymal stem cells. Int J Mol Cell Med 8(Suppl1):30 15. Goodarzi P, Larijani B, Alavi-Moghadam S et al (2018) Mesenchymal stem cells-derived exosomes for wound regeneration. Cell Biol Transl Med 4:119–131 16. Soung YH, Ford S, Zhang V et al (2017) Exosomes in cancer diagnostics. Cancers 9(1):8 17. Chen L, Wang L, Zhu L et al (2022) Exosomes as drug carriers in anti-cancer therapy. Front Cell Dev Biol 10:728616 18. Shen M, Chen T (2021) Mesenchymal stem cell-derived exosomes and their potential agents in hematological diseases. Oxid Med Cell Longev 2021:4539453

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19. Ros¸ca AM, T¸ut¸uianu R, Titorencu ID (2018) Mesenchymal stromal cells derived exosomes as tools for chronic wound healing therapy. Romanian J Morphol Embryol 59(3):655–662 20. Sun S-J, Wei R, Li F et al (2021) Mesenchymal stromal cell-derived exosomes in cardiac regeneration and repair. Stem Cell Reports 16(7): 1662–1673 21. Shokravi S, Borisov V, Zaman BA et al (2022) Mesenchymal stromal cells (MSCs) and their exosome in acute liver failure (ALF): a comprehensive review. Stem Cell Res Ther 13(1):1–21 22. Cao Q, Huang C, Chen X-M et al (2022) Mesenchymal stem cell-derived exosomes: toward cell-free therapeutic strategies in chronic kidney disease. Front Med 9:23 23. Chen Y-S, Lin E-Y, Chiou T-W et al (2020) Exosomes in clinical trial and their production in compliance with good manufacturing practice. Tzu-Chi Med J 32(2):113 24. Aghayan H-R, Goodarzi P, Arjmand B (2014) GMP-compliant human adipose tissue-derived mesenchymal stem cells for cellular therapy. In: Stem cells and good manufacturing practices, pp. 93–107. Springer 25. Aghayan HR, Payab M, Mohamadi-Jahani F et al (2020). GMP-compliant production of human placenta-derived mesenchymal stem cells. In: Stem cells and good manufacturing practices, pp. 213–225. Springer 26. Larijani B, Aghayan H-R, Goodarzi P et al (2014) GMP-grade human fetal liver-derived mesenchymal stem cells for clinical transplantation. In: Stem cells and good manufacturing practices, pp. 123–136. Springer 27. Ebrahimi-Barough S, Ai J, Payab M et al (2020) Standard operating procedure for the good manufacturing practice-compliant production of human endometrial stem cells for multiple sclerosis. In: Stem cells and good manufacturing practices, pp. 199–212. Springer 28. International B (2022). https://www. biophar minter national.com/view/con siderations-for-us-fetal-bovine-ser umsourcing

29. Arjmand B, Goodarzi P, Alavi-Moghadam S et al (2020) GMP-compliant human schwann cell manufacturing for clinical application. In: Stem cells and good manufacturing practices, pp. 227–235. Springer 30. Aghayan HR, Arjmand B, Burger SR (2016) GMP facilities for clinical cell therapy product manufacturing: a brief review of requirements and design considerations. Perinatal TissueDerived Stem Cells: 215–227 31. Arabi M, Mohamadi-Jahani F, AlaviMoghadam S et al (2020) Standards and regulatory frameworks (for cell-and tissue-based products). In: Biomedical product development: bench to bedside, pp. 89–97. Springer 32. Arjmand B, Alavi-Moghadam S, Payab M et al (2020) GMP-compliant adenoviral vectors for gene therapy. In: Stem cells and good manufacturing practices, pp. 237–250. Springer 33. Pawar SD, Khare AB, Keng SS (2021) Selection and application of biological safety cabinets in diagnostic and research laboratories with special emphasis on COVID-19. Rev Sci Instr 92(8):081401. https://doi.org/10. 1063/5.0047716 34. Blajchman M (2000) Reducing the risk of bacterial contamination of cellular blood components. Dev Biol 102:183–193 35. Langdon SP (2004) Cell culture contamination. Cancer Cell Culture 88:309–317 36. Lincoln CK, Gabridge MG (1998) Cell culture contamination: sources, consequences, prevention, and elimination. Methods Cell Biol 57: 49–65 37. Long EG, Buluk M, Gallagher MB et al (2019) Human mesenchymal stem cell morphology, migration, and differentiation on micro and nano-textured titanium. Bioact Mater 4:249– 255. https://doi.org/10.1016/j.bioactmat. 2019.08.001 38. Akyar I (2012) Standard operating procedures (what are they good for?). In: Latest research into quality control, pp. 367–391

Methods in Molecular Biology (2023) 2736: 177–192 DOI 10.1007/7651_2023_482 © Springer Science+Business Media, LLC 2023 Published online: 28 May 2023

Establishing Brain Tumor Stem Cell Culture from Patient Brain Tumors and Imaging Analysis of Patient-Derived Xenografts Elham Mahmoudian and Arezu Jahani-Asl Abstract Brain tumor stem cells (BTSCs) have been isolated from different types of brain tumors including glioblastoma. Although BTSCs share common characteristics with neural stem cells (NSCs), such as capacity to self-renew and undergo long-term proliferation, they have tumor-propagating capabilities. A small population of BTSC can give rise to secondary tumor when transplanted into severe immunodeficient (SCID) mice. The histological and cytological features, as well as genetic heterogeneity of the xenografted tumors in mice, closely resemble those of primary tumors in patients. Patient-derived xenografts (PDX), therefore, provide a clinically relevant model to study brain tumors. Here, we describe our protocol for establishing BTSC cultures following surgical excision of human brain tumors and the procedures to conduct PDX studies in SCID mice. We also provide our detailed step-by-step protocol on in vivo imaging system (IVIS) of the PDX tumors as a noninvasive method to trace the cells and tumor volume. Key words Bioluminescence, Brain tumors stem cells, Glioblastoma, In vivo imaging system, Magnetic resonance imaging, Optical imaging, Patient-derived xenografts, Reporter gene

1

Introduction Glioblastoma (GB) is the most malignant primary brain tumor in adults. The present standard of care includes surgical removal of the tumors followed by treatment with temozolomide (TMZ) and ionizing radiation (IR) therapy. Despite intense efforts, the median survival rate for GB patients remains ~18 months following diagnosis [1, 2]. GB tumors exhibit cellular heterogeneity, comprising both undifferentiated and differentiated cells, and contain a rare subpopulation of tumorigenic stem cells, termed brain tumor stem cells (BTSC) [2–8]. The identification of BTSC within GB tumors nearly two decades ago has transformed our understanding of GB pathogenesis [9–13]. Similar to neural stem cells (NSC), BTSCs have the capacity to either undergo long-term proliferation or

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differentiate into multiple lineages [14–16]. Proteins involved in the regulation of NSC self-renewal, such as transcription factors FoxG1, Sox2, Oct4, polycomb repressor Bmi1, and RNA-binding protein Musashi-1, are also expressed in the BTSCs [17–22]. Neurodevelopmental factors such as Olig2, Oct3, and Sall2 are also known to play important roles in BTSC maintenance [20]. Furthermore, BTSCs are highly proliferative but at the same time are resistant to DNA-damaging chemotherapy and radiation therapy [23, 24], raising a possibility that during the treatment of GB tumors, BTSC may generate a cellular hierarchy that contributes to the acquisition of drug resistance [25]. Due to their ability to differentiate into multiple lineages, BTSCs can generate all the cellular subpopulations within a tumor, suggesting that they can recapitulate the functional heterogeneity of the tumor. The transplantation of a limited number of BTSC into severe immunodeficient (SCID) mice can generate tumors that resemble those observed in human patients, exhibiting similar histological/cytological features and genetic heterogeneity [9–11, 26–28]. Patientderived stem cell xenografts (PDX), therefore, provide a clinically relevant model for investigating underlying molecular mechanisms of disease pathogenesis or conducting pharmacological screening for a patient-tailored therapeutic approach. In this protocol, we provide detailed method for establishing BTSC cultures from the patient tumors and analysis of secondary tumors in SCID mice via imaging platforms, focusing on in vivo imaging system (IVIS) as a simple and noninvasive method that allows real-time monitoring of the tumors in the brain from initiation to progression while analyzing different readouts including tumorigenesis, response to treatment, or analysis of animal survival [29–31]. IVIS is a reporter gene-based technology. The firefly luciferase (FLuc) gene is the most commonly used reporter that catalyzes D-luciferin oxidation to produce photons. The photons can be detected in vivo by whole-body bioluminescence imaging (BLI) [32, 33]. Due to its low background and high sensitivity, BLI can detect minor events. Several factors influence bioluminescence, including the distribution of D-luciferin, availability of co-factors, depth of signal, and absorption by tissues [29]. The luciferase– luciferin system doesn’t need external light excitation, and in presence of the D-luciferin, FLuc turnover occurs rapidly, which allows real-time, sensitive, quantitative imaging of FLuc expression in living subjects. This method also allows imaging of up to five animals in one run of the experimental duration (~20 min) and thus is more affordable than the alternative methods such as small animal magnetic resonance imaging (MRI) [30, 32].

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Materials 1. CB17.Cg-PrkdcscidLyst bg-J/Crl (Fox Chase SCID mouse strain #236). 2. Luciferase-expressing brain tumor stem cells (BTSCs).

2.1 Tools and Equipment

1. Software Living Image® 4.7.3 Software (October 2018). 2. IVIS® Spectrum Imaging System. 3. 1.5 mL Eppendorf safe-lock tubes. 4. Transfer pipettes. 5. 1 mL syringe. 6. Weigh scale. 7. Timer. 8. Cell counter. 9. Centrifuge. 10. Ice pack. 11. Hood. 12. Incubators. 13. Gloves. 14. A set of pipette. 15. Tips. 16. Large surgical scissors (Fine Science Tools, #14200-21, or similar). 17. Fine surgical scissors (Fine Science Tools, #14084-08, or similar). 18. Fine forceps (Fine Science Tools, #11251-35, or similar). 19. Spatula. 20. Petri dishes 100 mm, sterile. 21. 15 mL polypropylene conical centrifuge tubes. 22. 50 mL polypropylene conical centrifuge tubes. 23. 1.5 mL Eppendorf safe-lock tubes. 24. 0.2 μm sterile syringe filter. 25. 40 μm sterile cell strainer. 26. 10 mL syringe allergy syringe tray 1/2 mL 27 G REF 305535. 27. Hamilton syringe 10 μL gastight #1701. 28. Stereotaxic instruments anesthesia setups.

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Reagents

1. Isoflurane (ISO) 100% Inhalation Vapor, Liquid. 2. D-Luciferin (4,5-dihydro-2-(6-hydroxy-2-benzothiazolyl)-4thiazolecarboxylic acid potassium salt KC11H7N2O3S2), Monosodium Salt, (Pierce™ D-Luciferin, Monosodium Salt, cat. # 88291), molecular mass = 318.42 g/mol. 3. Dulbecco’s phosphate-buffered saline (D-PBS). 4. NeuroCult NS-A proliferation kit (Stem Cell Technologies, cat. # 05751). 5. Heparin solution (2 mg/mL) (Stem Cell Technologies, cat. # 7980). 6. Human EGF (20 ng/mL). 7. Human FGF (10 ng/mL). 8. Collagenase (Sigma C6885). 9. Kynurenic acid (Sigma K3375). 10. DNase I (Sigma 11284932001). 11. Accumax solution Innovative Cell Technologies. 12. Penicillin (100 U/mL)/streptomycin (100 mg/mL). 13. 70% Ethanol (ETOH). 14. 0.4% Trypan blue. 15. Optixcare® Eye Lube. 16. Carprofen. 17. Saline (0.9% NaCl). 18. Xylocaine 2% Jelly (Topical Anesthetic Lubricant).

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Methods Initial Setup

3.1 Establishing BTSC Cultures from Human Patients 3.1.1 Making Media 3.1.2 BTSC Isolation Following Surgical Excision of the Tumors

Brain tumor stem cell (BTSC) media is prepared prior to use as follow: Neurocult NS-A proliferation kit supplemented with 2 μg EGF/100 mL of media, 1 μg of FGF2/100 mL of media, and 100 μL of 0.2% heparin/100 mL of media. 1. Place a piece of brain tumor tissue in a sterile petri dish and wash with sterile 1× PBS containing penicillin and streptomycin. 2. Transfer the tissue to another sterile petri dish and mince the tissue with a sterile scalpel into small pieces. 3. For tissue banking, snap freeze a piece of tissue in liquid nitrogen and store it at -80 °C.

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4. For small and soft tissue, do a mechanical disruption using a scalpel and then pipette up and down in sterile 1× PBS to break up the tissue. If the tissue size is large or highly infiltrated, enzymatic digestion may be required (see Note 1). 5. Spin tissue at 50 g for 5 min. 6. If the sample is highly vascularized, a hypotonic lysis step is required (see Note 2). 7. Place a 70 μm cell strainer on a 50 mL falcon tube and pass the suspension through the strainer. 8. Wash the strainer with an additional 10 mL of 1× PBS passing it through the cell strainer. 9. Spin the solution from step 8 at 50 g for 5 min. 10. Remove the supernatant and add 1–10 mL of BTSC media depending on the size of the pellet (see Note 3). 11. Pipette up and down 5–10× to resuspend the pellet. 12. Count cells and plate between 200,000 and 1 million cells in a T25 flask in 6 mL of BTSC media (see Note 4). 13. On the third day following initial plating, add 4 mL of freshly made media to each flask. 14. On the seventh day, passage the cells by collecting the content of the flask and spinning in a 15 mL tube at 50 g for 5 min. Add 5 mL of fresh BTSC media to resuspend the pellet and add it back to the flask. 15. Feed the flask every 3–4 days until the spheres are large enough (usually 200–300 μm in diameter), and then proceed with passaging by resuspending the spheres in a 15 mL tube and spinning at 200 g for 10 min. 16. Remove the media and resuspend the cells in 1 mL of media by triturating 15–20 times with a p1000 pipette set at 800 μL using low retention tips. 17. Transfer the cells into a flask with 10 mL of fresh BTSC media (see Note 5). 3.2 BTSC Maintenance (Passage, Freezing Down/Thawing) 3.2.1 Passaging Established Cell Lines

1. Once spheres are large enough (~200–300 μm), passage the cells as described in Subheading 3.1. Following triturating 25–30 times with the P1000 using low retention pipette tips, switch to a P200 pipette and triturate another 25–30 times with low retention tips until a homogeneous solution (usually single cell) is obtained. 2. Add the cells to a new flask with fresh BTSC media for experimental use or freeze down the cells as described in Subheading 3.2.2.

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3.2.2 Freezing BTSCs

1. Collect cells in a 15 or 50 mL tube depending on the number of cells in the solutions being collected and spin cells at 200 g for 10 min (see Note 6). 2. Remove the supernatant and resuspend the pellet with 1 mL of 10% DMSO-BTSC media solution. 3. Transfer the solution into a cryovial. 4. Place cryovials in a cryo-freezing container at -80 °C overnight. 5. Transfer the cryovials to liquid nitrogen tank for long-term storage.

3.2.3 Thawing Frozen BTSCs

1. Remove a cryovial containing BTSCs from liquid nitrogen tank and place it in a 37 °C water bath to thaw the cells quickly. 2. Add the content to 9 mL of BTSC media in a 15 mL tube and mix gently. 3. Spin cells at 200 g for 10 min. 4. Aspirate the media and resuspend the pellet in 10 mL of fresh media and plate it in a flask.

3.3 Tumor Induction and Mice Preparation 3.3.1 Preparation of Luciferase-Expressing BTSC Cell Line for Injection

1. Warm the BTSC medium in a water bath at 37 °C for 10 min. 2. Using a 10 mL serological pipette, collect the cell solution contained in the flask at 37 °C. 3. Transfer the cells to a 15 mL polypropylene conical tube. 4. Centrifuge for 10 min at 200 g at room temperature (22–25 °C) and discard the media by aspiration (see Note 7). 5. Add 200 μL of Accumax solution and mix gently using a P200 micropipette and incubate at 37 °C for 10 min in the incubator or in a water bath. 6. Remove the cell-Accumax-containing tubes from the incubator and triturate gently 10 times using a P200 micropipette and low retention filtered long pipette tips to homogenize. 7. Add 800 μL of BTSC plating media to the 200 μL Accumaxcell suspension. 8. Count the number of viable cells using an exclusion dye such as trypan blue. 9. Prepare a cell suspension of 3 × 105 cells/mL in BTSC medium and homogenize by pipetting up and down using a P1000 micropipette. 10. Centrifuge the cell suspension at 200 g for 5 min at 4 °C. 11. Discard the media.

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Fig. 1 Schematic representation of BTSCs preparation followed by xenografting into the mouse brain. After employing Accuamx to single cell the spheres, and exclusion of dead cells, the cells are resuspended in 1× D-PBS. 3 × 105 luciferase-expressing BTSCs are stereotactically implanted into the right striata of the 8-week-old SCID mice. (Created using Biorender.com)

12. Add 4 μL of D-PBS to the pellet and homogenize by pipetting up and down using a P20 micropipette. 13. Keep the cells on the ice before injection (Fig. 1). 3.3.2 Animal Preparation (Tumor Induction)

1. All experiments on animals should be conducted under institutional guidelines. The experiments in this protocol were approved by the uOttawa Animal Care Committee. 8-weekold male CB17.Cg-PrkdcscidLyst bg-J/Crl (Fox Chase SCID mouse strain #236) was used for this protocol. Housing room temperature and relative humidity were adjusted to 22.0 ± 2.0 °C and 55.0 ± 10.0%, respectively. The light/dark cycle was adjusted to 12 h lights-on and 12 h lights-off. Autoclaved water and irradiated food pellets were given ad libitum. 2. 3 × 105 luciferase-expressing BTSCs were stereotactically implanted into the right striata with the following coordinates: medial-lateral (ML) 0.8 mm, anterior-posterior (AP) 1 mm dorsal to bregma, and dorsal-ventral (DV) 2.5 mm. 3. Carprofen was administered for 2 days after injection at 20 mg/kg every 24 h. 4. Seven and twenty-one days post-injection, mice were subjected to bioluminescence imaging to trace the injected cells and tumor initiation.

3.3.3 D Luciferin Preparation for Injection

1. Thaw D luciferin (either potassium or sodium salt) at room temperature. 2. Dissolve D luciferin in D-PBS to a final concentration of 15 mg/mL (see Note 8). 3. Put the cage of the animal under the hood and weigh each mouse.

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4. Inject luciferin using a syringe. The adequate amount of luciferin is based on the weight of each animal. The standard dosage for injection is 150 mg/kg injection (see Note 9). 3.4

Performing IVIS

1. IVIS spectrum instrument (PerkinElmer, MA, USA) was used and Bioluminescent imaging (BLI) was quantified using Living Image 2.0 software (PerkinElmer, MA, USA) in the present protocol. 2. The IVIS scanner is connected to a resource of an anesthetic agent and a chamber to sedate mice (see Note 10). 3. Prior to use, check the level of isoflurane in the vaporizer to ensure that the isoflurane reservoir is replenished to the level indicated by the upper triangle mark whenever a decrease in volume is observed (see Note 11). 4. Check the scavenging system by weighing the charcoal canisters and recording the weight on the canisters (see Note 12). 5. Attach canisters to filter waste gas from the induction chamber and IVIS scanner (see Note 12). 6. Turn on the scavenger or evacuation pump on the IVIS machine to initiate the removal of excess gas from the IVIS device. 7. The initialization of the imaging system is imperative before its usage. Upon initiation of the Living Image software, the “Initialize” button on the software’s control panel should be clicked (refer to initialization of the IVIS spectrum, Subheading 3.6.2). 8. Upon achieving the demand temperature (-90 °C or -105 °C for IVIS Systems cooled by a Cryotiger® unit), the system is initialized. This is evidenced by the temperature box displaying the color green, signifying the readiness of the instrument for operation. 9. Administer D-luciferin intraperitoneally to the mice (refer to D luciferin preparation, Subheading 3.3.3). To ensure the optimal signal intensity of luciferase, a minimum of 15 min is recommended before acquiring images, as this duration corresponds to the approximate stabilization point of the luminescent signal. 10. Place the mice inside the induction chamber 5 min after the administration of D luciferin, turn the vaporizer dial to 3%, and set oxygen flow to 1 L/min (LPM) (see Note 13). 11. Apply sterile eye lubricant to each eye of the mice. 12. Transfer the mice to the IVIS scanner 5–6 min after anesthesia following switching the anesthesia system from the chamber to the IVIS scanner (see Note 14).

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Fig. 2 Workflow diagram in IVIS imaging procedure. Once the weight of the mouse is measured, sufficient luciferin is given, and a chamber box connected to oxygen (O2) and isoflurane (ISO) is used to sedate the mouse and make it ready for IVIS imaging. (Created using Biorender.com)

13. Initiate the imaging process by clicking on the “acquire” button 15 min after the administration of D-luciferin (Fig. 2). 14. Once the image has been captured, it can be saved into a designated directory (refer to acquire a luminescent image, Subheading 3.6.3). 15. Transfer the mice to the recovery cage. 16. Shut down the anesthesia by turning the isoflurane vaporizer to “0” and setting the oxygen flow meter to 0 LPM. 17. Take them back to the animal room once the mice get full recovery. 3.5 Determining the Kinetic Curve

In general, in most mouse strains, the peak signal time is 15 min, but to determine the peak signal time and plateau phase for each animal model and system, a kinetic study of luciferase can be conducted. A kinetic curve for luciferase activity can be obtained by following these steps: 1. Luciferin should be injected based on the weight of the animals, as per standard protocol. 2. Sedation of the animals prior to injection may prolong the kinetics (i.e., peak expression time of luciferase). The way luciferin is administered may also affect its biodistribution. 3. Wait for 2–3 min after injection.

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4. Sedate the animal with isoflurane and oxygen vapor, which is most commonly used to sedate small animals. 5. Transfer the animals to the IVIS scanner. 6. Take the first image 10 min after luciferin injection on sedated animals. 7. A kinetic curve can be generated by acquiring images at 2 minute intervals for a duration of 20–30 min (see Note 15). 3.6 Analysis Using the Living Image® Software

IVIS Imaging Systems use Living Image software to acquire, view, and analyze images. Other workstations only provide analysis features, while IVIS Spectrum’s PC workstation provides both acquisition and analysis software.

3.6.1 Create an Account

Follow the institutional guidelines which typically request the users to create an account. Click on Living Image under find “Caliper Life Sciences,” which should appear by clicking all programs. Then select a user ID from the drop-down list and enter a password in the dialog box.

3.6.2 Initialization of the IVIS Spectrum

1. After logging in, the control panel interface of the software becomes available. Ensure that the imaging system should be initialized every time the Living Image software is launched. The initialization procedure resets all settings to the default including electronics, controllers, and software variables. Also, the initialization moves all motor-driven components to their default positions. 2. To initialize the IVIS Imaging System, click on Initialize in the control panel. In the IVIS acquisition control panel, the temperature status of the charge-coupled device (CCD) camera is displayed. 3. When the temperature box turns green, it indicates that the temperature has reached -90 °C or - 105 °C for IVIS Systems, and the system has been initialized. The green light indicates that the instrument is ready for operation and image acquisition.

3.6.3 Acquire a Luminescent Image

1. In the main window, the control panel provides the image acquisition functions. 2. To get a Luminescent Image, put a checkmark next to Luminescent and Photograph. 3. Select Auto exposure in the control panel. 4. Field of view (FOV) is the size of the stage area to be imaged. Select an FOV based on Table 1 and the number of animals that you want to put in the device. For example, when imaging 2–3 mice, it is better to use a larger FOV (C or D).

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Table 1 Field of view (FOV) settings – IVIS spectrum FOV setting

FOV (cm)

A

4

B

6.5

Ca

13

D

22.5

a

Position C is the default setting

5. Select a focus option in the control panel. 6. To achieve optimal focus on the top side of an animal, the stage should be set at Z = 0. This is a universal condition for dorsal focusing irrespective of the configuration of the field of view. 7. Once all the setup has been done, click “Acquire” and wait until the imaging is completed (Fig. 3). 8. A dialog box will be popped out to prompt the user to enter the image information (optional) while the imaging process is in progress. The user can confirm the input by clicking “OK.” 9. The system has an autosave function as well. Select Acquisition → Auto-Save on the menu bar. Also, the image can be saved in the desired folder. 10. When the acquisition is finished, an image analysis toolbar will appear automatically by which the image can be modified or add some information on the right or left side of the image. 11. To analyze the images, click on “ROI tools” in the tool palette. 12. Select the ROI style and determine the number of ROIs. Then, modify the position and dimensions of the ROI to cover the area of interest. 13. By double-clicking on the region of interest (ROI), additional options will become accessible, allowing for the creation of an ROI or a set of ROIs designed for one animal. The same size of ROIs can be retained across all animals to acquire comparable data by duplicating the created ROI(s) to all other animals. 3.7 Magnetic Resonance Imaging

MRI was conducted to visualize and verify tumor growth in vivo using a 7T GE/Agilent MRI machine at the University of Ottawa Preclinical Imaging Core Facility. The mouse was subjected to MRI at the end of the assay. 1. A common method for performing an MRI on mice is to use inhalational anesthetic, isoflurane. In a resource of an anesthetic agent and connected chamber box, at the beginning to sedate mice, turn on the chamber oxygen to 1 and isoflurane to 2–3% with a flow rate of 0.8–1.0 L/min.

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Fig. 3 Bioluminescence imaging of BTSC-derived xenografts. Mice injected with BTSCs were imaged 7 and 21 days after surgery to trace the luciferaseexpressing BTSC injection. The IVIS image is transformed into pseudocolor images where each numerical value is assigned a specific color. A color table defines the relationship between numerical data and colors in the image. Each pixel has a photon intensity value, with brighter areas corresponding to higher photon detection

2. Transfer the mouse to this chamber and let it be sedated. 3. Once the mouse is anesthetized completely, transfer it to the MRI frame which is connected to the anesthetic agent, and keep it with isoflurane in O2: induction at 3%, maintenance at 1.5% isoflurane.

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Fig. 4 MRI imaging of BTSC-derived xenografts. Multiple MRI images were captured in a sequence from the prefrontal cortex (PFC) at a thickness of 300 μm. A coronal 2D fast spin echo T2-weighted pulse sequence is utilized, with repetition time of 4500 ms, effective echo time of 13 ms, field of view of 3 cm, matrix size of 256 × 256, number of averages of 2, and a scanning duration of 6 min. The mouse has a visible intracranial tumor (the mass in the right portion of the brain) at the site of injection (right panel). No tumor controls are presented in the left panel

4. In this experiment, serial MRI images were taken from the prefrontal cortex (PFC) at 300 μm thickness. A coronal 2D fast spin echo T2-weighted pulse sequence was used with repetition time = 4500 ms, effective echo time = 13 ms, field-ofview = 3 cm, matrix size = 256 × 256, number of averages = 2, and scan time = 6 min (Fig. 4). 5. Following MRI, the mouse needs to be put back in the home cage and be observed until the animal is recovered from anesthesia.

4

Notes 1. If the tumor tissue size is large or highly infiltrated, an optional enzymatic digest can be done with a filter sterilized collagenase enzyme mix that includes 0.2 mg/mL collagenase, 0.2 mg/ mL kynurenic acid, and 0.5 mg/mL DNase I. 2. If the sample is highly vascularized, add 9 mL of sterile water and pipette up and down several times to dissociate the pellet followed by addition of 1 mL of 10× PBS and pipette up and down.

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3. The amount of BTSC media depends on the size of the pellet. Typically aim to have 200,000 cells per mL prior to plating. 4. In cases where an excess of cell debris is observed, changing half of the media after 2 h of culture can be beneficial. This is due to the tendency of viable cells to settle down in the flask, while the cellular debris remains suspended in the media during the first 12 h of culture. 5. The cells may take more than a week to recover and start growing healthy spheres. 6. It is best to freeze BTSC cells as small healthy spheres and at low passage. 7. Aspirate gently or use a serological pipette carefully to avoid losing the pellet. It is advised to leave 200–500 μL of media on top of the pellet. 8. The protocol for dissolving D-luciferin may vary depending on the supplier. 9. As an example, 10 μL of luciferin stock solution is injected per gram of body weight (normally ~200 μL for a 20 g mouse). 10. Isoflurane is the most common anesthetic used for laboratory experiments in mice and for IVIS imaging. 11. Note that isoflurane container should not be overfilled as it can pose a safety hazard and may result in a spill or leak of the anesthetic gas. Also, overfilling the isoflurane container can cause inaccurate readings of the anesthetic levels and lead to improper dosing during procedures. 12. Isoflurane is a commonly used anesthetic that can produce harmful effects if it is not properly scavenged and removed from the breathing system during imaging procedures. The carbon canisters used to scavenge isoflurane works by absorbing the gas into the carbon filter, and over time, the carbon becomes saturated with isoflurane and loses its effectiveness. To ensure the proper removal of isoflurane, the carbon canisters must be weighed before and after each imaging to monitor the amount of isoflurane being captured. If the weight of the canister increases by more than 50 g from its initial weight, it indicates that the carbon is saturated with isoflurane and can no longer effectively capture additional gas. Therefore, the canisters should be weighed before each imaging session to make sure their weight is not over 50 g of the original weight. 13. If you have more than one mouse, start the timer after the last injection. 14. It is not advisable to transfer mice immediately after injecting them with D-luciferin. This is because prolonged exposure to anesthesia can be detrimental to the health of the mice, and it is

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therefore recommended to minimize the duration of anesthesia as much as possible. 15. Note that the best time point for imaging your model is determined by the kinetic curve. The peak signal in most of the models occurs around 15 min after IP or SQ Luciferin injection.

Acknowledgments This work was supported by CIHR grants #183710, #179865, #162198 and The brain tumour charity #497225 to AJ-A. AJ-A holds a Canada Research Chair. All MRI and IVIS images were acquired by the University of Ottawa Preclinical Imaging Core Facility (PCIC, RRID: SCR_021832). We thank members of Jahani-Asl laboratory, in particular Dr. Dianbo Qu for reading and editing the protocol. We thank Preclinical Imaging Core Facility, and animal care and veterinary service members of the university of Ottawa. We thanks Roland Pilgram at the University of Ottawa for conducting the MRI imaging. References 1. Stupp R, Mason WP, van den Bent MJ, Weller M, Fisher B, Taphoorn MJ, Belanger K, Brandes AA, Marosi C, Bogdahn U, Curschmann J, Janzer RC, Ludwin SK, Gorlia T, Allgeier A, Lacombe D, Cairncross JG, Eisenhauer E, Mirimanoff RO (2005) Radiotherapy plus concomitant and adjuvant temozolomide for glioblastoma. N Engl J Med 352:987–996 2. Louis DN, Ohgaki H, Wiestler OD, Cavenee WK, Burger PC, Jouvet A, Scheithauer BW, Kleihues P (2007) The 2007 WHO classification of tumours of the central nervous system. Acta Neuropathol 114:97–109 3. Louis DN (2006) Molecular pathology of malignant gliomas. Annu Rev Pathol 1:97–117 4. Bernstein JJ, Woodard CA (1995) Glioblastoma cells do not intravasate into blood vessels. Neurosurgery 36:124–132. discussion 132 5. Holland EC (2001) Gliomagenesis: genetic alterations and mouse models. Nat Rev Genet 2:120–129 6. Bachoo RM, Maher EA, Ligon KL, Sharpless NE, Chan SS, You MJ, Tang Y, DeFrances J, Stover E, Weissleder R, Rowitch DH, Louis DN, DePinho RA (2002) Epidermal growth factor receptor and Ink4a/Arf: convergent mechanisms governing terminal differentiation

and transformation along the neural stem cell to astrocyte axis. Cancer Cell 1:269–277 7. Uhrbom L, Dai C, Celestino JC, Rosenblum MK, Fuller GN, Holland EC (2002) Ink4a-Arf loss cooperates with KRas activation in astrocytes and neural progenitors to generate glioblastomas of various morphologies depending on activated Akt. Cancer Res 62:5551–5558 8. Bajenaru ML, Hernandez MR, Perry A, Zhu Y, Parada LF, Garbow JR, Gutmann DH (2003) Optic nerve glioma in mice requires astrocyte Nf1 gene inactivation and Nf1 brain heterozygosity. Cancer Res 63:8573–8577 9. Singh S, Dirks PB (2007) Brain tumor stem cells: identification and concepts. Neurosurg Clin N Am 18:31–38, viii 10. Singh SK, Clarke ID, Terasaki M, Bonn VE, Hawkins C, Squire J, Dirks PB (2003) Identification of a cancer stem cell in human brain tumors. Cancer Res 63:5821–5828 11. Singh SK, Hawkins C, Clarke ID, Squire JA, Bayani J, Hide T, Henkelman RM, Cusimano MD, Dirks PB (2004) Identification of human brain tumour initiating cells. Nature 432:396– 401 12. Galli R, Binda E, Orfanelli U, Cipelletti B, Gritti A, De Vitis S, Fiocco R, Foroni C, Dimeco F, Vescovi A (2004) Isolation and

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characterization of tumorigenic, stem-like neural precursors from human glioblastoma. Cancer Res 64:7011–7021 13. Yuan X, Curtin J, Xiong Y, Liu G, Waschsmann-Hogiu S, Farkas DL, Black KL, Yu JS (2004) Isolation of cancer stem cells from adult glioblastoma multiforme. Oncogene 23:9392–9400 14. Park DM, Rich JN (2009) Biology of glioma cancer stem cells. Mol Cells 28:7–12 15. Lathia JD, Mack SC, Mulkearns-Hubert EE, Valentim CL, Rich JN (2015) Cancer stem cells in glioblastoma. Genes Dev 29:1203–1217 16. Chen J, McKay RM, Parada LF (2012) Malignant glioma: lessons from genomics, mouse models, and stem cells. Cell 149:36–47 17. Kaneko Y, Sakakibara S, Imai T, Suzuki A, Nakamura Y, Sawamoto K, Ogawa Y, Toyama Y, Miyata T, Okano H (2000) Musashi1: an evolutionally conserved marker for CNS progenitor cells including neural stem cells. Dev Neurosci 22:139–153 18. Manoranjan B, Wang X, Hallett RM, Venugopal C, Mack SC, McFarlane N, Nolte SM, Scheinemann K, Gunnarsson T, Hassell JA, Taylor MD, Lee C, Triscott J, Foster CM, Dunham C, Hawkins C, Dunn SE, Singh SK (2013) FoxG1 interacts with Bmi1 to regulate self-renewal and tumorigenicity of medulloblastoma stem cells. Stem Cells 31:1266–1277 19. Graham V, Khudyakov J, Ellis P, Pevny L (2003) SOX2 functions to maintain neural progenitor identity. Neuron 39:749–765 20. Suva ML, Rheinbay E, Gillespie SM, Patel AP, Wakimoto H, Rabkin SD, Riggi N, Chi AS, Cahill DP, Nahed BV, Curry WT, Martuza RL, Rivera MN, Rossetti N, Kasif S, Beik S, Kadri S, Tirosh I, Wortman I, Shalek AK, Rozenblatt-Rosen O, Regev A, Louis DN, Bernstein BE (2014) Reconstructing and reprogramming the tumor-propagating potential of glioblastoma stem-like cells. Cell 157: 580–594 21. Fasano CA, Dimos JT, Ivanova NB, Lowry N, Lemischka IR, Temple S (2007) shRNA knockdown of Bmi-1 reveals a critical role for p21-Rb pathway in NSC self-renewal during development. Cell Stem Cell 1:87–99 22. Abdouh M, Facchino S, Chatoo W, Balasingam V, Ferreira J, Bernier G (2009)

BMI1 sustains human glioblastoma multiforme stem cell renewal. J Neurosci 29:8884– 8896 23. Chen J, Li Y, Yu TS, McKay RM, Burns DK, Kernie SG, Parada LF (2012) A restricted cell population propagates glioblastoma growth after chemotherapy. Nature 488:522–526 24. Bao S, Wu Q, McLendon RE, Hao Y, Shi Q, Hjelmeland AB, Dewhirst MW, Bigner DD, Rich JN (2006) Glioma stem cells promote radioresistance by preferential activation of the DNA damage response. Nature 444:756–760 25. Venugopal C, Hallett R, Vora P, Manoranjan B, Mahendram S, Qazi MA, McFarlane N, Subapanditha M, Nolte SM, Singh M, Bakhshinyan D, Garg N, Vijayakumar T, Lach B, Provias JP, Reddy K, Murty NK, Doble BW, Bhatia M, Hassell JA, Singh SK (2015) Pyrvinium targets CD133 in human glioblastoma brain tumor-initiating cells. Clin Cancer Res 21:5324–5337 26. Parada LF, Dirks PB, Wechsler-Reya RJ (2017) Brain tumor stem cells remain in play. J Clin Oncol 35:2428–2431 27. Singh SK, Clarke ID, Hide T, Dirks PB (2004) Cancer stem cells in nervous system tumors. Oncogene 23:7267–7273 28. Vescovi AL, Galli R, Reynolds BA (2006) Brain tumour stem cells. Nat Rev Cancer 6:425–436 29. Berger F, Paulmurugan R, Bhaumik S, Gambhir SS (2008) Uptake kinetics and biodistribution of 14C-D-luciferin--a radiolabeled substrate for the firefly luciferase catalyzed bioluminescence reaction: impact on bioluminescence based reporter gene imaging. Eur J Nucl Med Mol Imaging 35:2275–2285 30. Gheysens O, Gambhir SS (2005) Studying molecular and cellular processes in the intact organism. Prog Drug Res 62:117–150 31. Bhaumik S, Gambhir SS (2002) Optical imaging of Renilla luciferase reporter gene expression in living mice. Proc Natl Acad Sci U S A 99:377–382 32. Contag PR, Olomu IN, Stevenson DK, Contag CH (1998) Bioluminescent indicators in living mammals. Nat Med 4:245–247 33. Weissleder R, Pittet MJ (2008) Imaging in the era of molecular oncology. Nature 452:580– 589

Methods in Molecular Biology (2023) 2736: 193–205 DOI 10.1007/7651_2023_500 © Springer Science+Business Media, LLC 2023 Published online: 01 September 2023

An Optimized Protocol for piggyBac-Induced iPSC Generation from hPBMCs by Automatic Electroporation Pelin Kilic

and Begum Cosar

Abstract Thanks to the Yamanaka transcription factors, pluripotency is recovered in the cell culture dish during in vitro cell reprogramming. Induced pluripotent stem cells (iPSCs) have been introduced to the scientific world as a source of disease models, which are predominantly used in drug discovery and monitoring disease pathophysiology, and as a source of master cell lines for developing cellular therapies. Successfully attaining iPSCs requires careful optimization of many factors, including selection of the transfection method and the appropriate transfection agents; culturing conditions before and after transfection; recovery conditions after transfection, freezing, and thawing; storage conditions; and the choice of cost-effective cell and colony characterization and molecular verification steps. In our optimized procedure, we describe the isolation of peripheral blood mononuclear cells (PBMCs) after blood harvest and their efficient reprogramming into iPSCs using automated electroporator, with piggyBac. Key words Cryopreservation, Electroporation, Induced pluripotent stem cells (iPSCs), Passaging, Peripheral blood mononuclear cells (PBMCs), piggyBac, Transfection

1

Introduction Embryonic stem cells (ESCs) are totipotent cells with self-renewal, differentiation, and colony formation capacities, with the capability to transform into all kinds of tissues and organs [1]. After a series of studies on fibroblasts by Takahashi and Yamanaka [2, 3], four genes were recognized for reprogramming fibroblasts into ESC-like cells. These genes were OCT4, SOX2, KLF4, and c-MYC [4]. Thanks to these transcription factors, pluripotency is recovered during reprogramming. Hence, it is possible to use ESC-like induced pluripotent stem cells (iPSCs) in research and development (R&D) today, which has helped to eliminate ethical concerns [5]. Thanks to their properties, iPSCs have been introduced to the scientific world as an alternative source with a wide range of clinical applications such as drug discovery and monitoring disease pathophysiology, and providing master cell lines for autologous/allogeneic genetically modified cellular therapies [6–8]. Considering these advantages, researchers are still working to improve protocols to produce more efficient and high-purity

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iPSCs. Successfully attaining iPSCs requires careful optimization of many factors, including selection of the transfection method and the appropriate transfection agents [9], culturing conditions before and after transfection [10], recovery conditions after transfection, freezing and thawing [11], storage conditions [12], and the choice of cost-effective cell and colony characterization and molecular verification steps. To date, there have been a number of protocols issued for iPSC generation [1, 13–18]. To our knowledge, there is no previous protocol specifically addressing a combination of automated electroporator-dependent, piggyBac transposon-mediated iPSC generation from human peripheral blood mononuclear cells (hPBMCs). Our optimized procedure comprehensively explains how to dedifferentiate hPBMCs into hiPSCs. Within this scope, we describe the isolation of PBMCs after blood harvest from patients and healthy volunteers, optional cultivation steps until transfection, and efficient reprogramming into iPSCs using an automated electroporator, with the piggyBac transposontransposase system as the vector of choice. Additionally, we provide alternative approaches regarding the establishment of different conditions for both PBMC and iPSC cultivation steps to create the most effective methodology.

2

Materials Filter all the non-sterile solutions with sterile syringe filter before use (see Note 1). Prepare all media and reagents at ambient temperature, and store them according to the manufacturer’s recommendations (see Note 2). Be careful of decontamination and sanitizing procedures before and after working on critical and general surfaces, accordingly. We generally prefer additional sporicidal agents such as 3% v/v hydrogen peroxide to accompany 70% v/v ethanol or isopropyl alcohol. Follow necessary disposal procedures in accordance with universal laboratory biosafety rules for waste materials.

2.1 Peripheral Blood Harvest and Transfer Conditions, PBMC Isolation Preparation, and Culture Conditions

1. Green top blood collection tubes with sodium heparin preservative: Ask the healthcare professional to harvest 5 mL of the donor’s peripheral blood (see Notes 3 and 4). Store at ambient temperature for a maximum of 4 h before cell isolation for best results. 2. Heparinized blood sample (5 mL or 10 mL). 3. Dulbecco’s phosphate buffered saline without calcium and magnesium (DPBS) containing 0.05% fetal bovine serum (FBS) (PBS/FBS): For 5 mL heparinized blood sample, add

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275 μL FBS to enough PBS to make a final volume of 55 mL (see Note 5). 4. Density gradient medium: Prefer a commercially available density gradient of 1.077 g/mL (see Note 6). 5. Basal RPMI 1640 medium: Used for suspending the centrifuged cell pellet in 1 mL for cell count. 6. Stimulated complete mononuclear cell culture medium: For 50 mL of complete medium, add 5 mL FBS, 500 μL penicillin/streptomycin/amphotericin B (100×), 10 μL interleukin 3 (IL-3) human, 10 μL stem cell factor (SCF), 50 μL insulin-like growth factor (IGF-1) human and 5.5 mL of commercially available basal mononuclear cell culture medium supplement to enough commercially available basal mononuclear cell culture medium for a final volume of 50 mL (see Note 7). Store at 4 °C when not in use (see Note 8). 2.2 PBMC Cryopreservation Conditions

1. DPBS: Use for washing the cell pellet after centrifugation. 2. Basal RPMI medium: Use for resuspending the washed cells ready for cryopreservation and also use at the thawing step to resuspend PBMCs. 3. PBMC cryopreservation medium: Cell suspension in basal RPMI 1640 medium, FBS, dimethyl sulfoxide (DMSO) (5: 4:1) (see Note 9).

2.3

Vector Isolation

1. PB-EF1α-OCT4-SOX2-KLF4-MYC-IRES-GFP 4-in-1 iPSC piggyBac Vector: Isolate before use.

Human

2. Commercially available plasmid extraction kit: Use for the extraction of the plasmid from Escherichia coli (E. coli). 3. Agarose gel (1%): Use for plasmid base pair (BP) verification by gel electrophoresis. We prefer a concentration of 1% in relation to the length of the piggyBac transposon. 4. Ethidium bromide: Use to stain the DNA gel and visualize the DNA content of the plasmid (see Note 10). 5. DNA ladder (1 kb): Use as a DNA standard. 2.4

Transfection

1. PBMC cell suspension: Transfect these cells (see Note 11). In our laboratory, we use fresh PBMCs right after cell isolation, cultivated PBMCs, or frozen-thawed PBMCs for transfection. This depends on the planned status and timeline of the study in question. 2. PB-EF1α-OCT4-SOX2-KLF4-MYC-IRES-GFP Human 4-in-1 iPSC piggyBac Vector isolate: Use in the optimal concentration.

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3. Transfection kit: Use with the appropriate automated transfector for accurate transfection. 2.5 Post-transfection Cell Recovery and iPSC Culture Conditions

1. Unstimulated MEM complete medium: For 50 mL of complete medium, add 5 mL FBS, 500 μL penicillin/streptomycin/amphotericin B (100×) to enough basal MEM for a final volume of 50 mL. Use as recovery medium right after transfection (see Note 12). 2. Vitronectin: Use for coating the bottom of the cell culture plate for iPSC adhesion (see Note 13). 3. Commercially available human PSC-specific xeno-free and feeder-free medium: Use for iPSC growth and expansion (see Note 14). 4. DPBS: Use for washing the culture plates in order to discard any unwanted contaminants before passaging.

2.6 iPSC Cryopreservation Conditions and Passaging

1. DPBS: Use for washing the cells after centrifugation. 2. Human PSC-specific xeno-free and feeder-free medium: Use for resuspending the washed cells ready for cryopreservation and also use at the thawing step to resuspend iPSCs (see Note 15). 3. iPSC cryopreservation medium: Cell suspension in human PSC-specific xeno-free and feeder-free medium, dimethyl sulfoxide (DMSO) (9:1) (see Note 16). 4. Commercially available enzyme-free reagent for dissociation and passaging of PSCs: Use for passaging iPSCs (see Note 17). 5. Rho kinase (ROCK) inhibitor (10 mM): Use to block possible cell apoptosis.

3

Methods Carry out all cellular procedures in the cell culture laboratory, under a class II laminar flow biological safety cabinet, with the use of a 5% CO2 incubator, a stainless steel water bath, appropriate centrifuge and storage devices, and disposables and personal protective equipment appropriate for maintaining sterile cell culturing conditions. Vector isolation can be carried out in the microbiology laboratory.

3.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation

1. Add 20 mL of PBS/FBS to 10 mL of blood sample in a 50 mL falcon (2:1). 2. Add the diluted blood sample very slowly onto another 50 ml falcon tube with a 1:1 ratio of density gradient medium. 3. Centrifuge at 750 × g for 20 min (Accel = 0 Decel = 0).

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4. Carefully transfer the buffy coat with a Pasteur pipette into a separate 50 ml falcon. 5. Add enough PBS/FBS to make a final volume of 50 mL. 6. Centrifuge at 350 × g for 10 min (Accel = 0 Decel = 0). 7. Discard the supernatant. 8. Dissolve the pellet in 25 mL of PBS/FBS. 9. Centrifuge at 160 × g for 15 min (Accel = 0 Decel = 0). 10. Discard the supernatant (see Note 18). 11. Add 1 mL of basal medium to resuspend the cells and perform cell count (see Note 19). 3.2 PBMC Cultivation Before Transfection

1. Inoculate the cell suspension onto well plates with stimulated complete mononuclear cell culture medium (see Note 20). 2. Upon reaching the desired confluence (Fig. 1a–c) (see Note 21), centrifuge at 600 × g for 5 min (Accel = 0 Decel = 0). 3. Discard the supernatant. 4. Wash with DPBS. 5. Centrifuge at 600 × g for 5 min (Accel = 0 Decel = 0). 6. Add 1 mL of basal medium to resuspend the cells and perform cell count and viability assay.

3.3 PBMC Cryopreservation and Thawing Before Transfection

1. Transfer the cell suspension into a falcon tube and centrifuge at 300 × g for 10 min. 2. Discard the supernatant. 3. Resuspend the cell pellet in the appropriate volume of basal medium so that the optimal number of cells are preserved per vial. 4. Prepare the cryopreservation medium inside the cryogenic vial (see Note 22). 5. Place the cryogenic vials inside a freezing container and perform interim storage overnight at -86 °C. 6. The next day, transfer the cryogenic vials into a liquid nitrogen (LN) tank for long-term storage. 7. For further processing, remove a cryogenic vial from the LN tank and swiftly thaw (see Note 23). 8. Transfer all of the cell suspension by aspirating dropwise with a small amount of basal medium.

3.4

Vector Isolation

1. Isolate the vector with the use of a commercially available plasmid extraction kit, in accordance with the manufacturer’s protocol.

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Fig. 1 (a) Peripheral blood mononuclear cells (PBMCs) entering cultivation (2 days to transfection) (image under inverted microscope; 10×). (b) Peripheral blood mononuclear cells (PBMCs) being cultivated (reaching confluence 24 h to transfection) (image under inverted microscope; 10×). (c) Cultivated peripheral blood mononuclear cells (PBMCs) (highly confluent at the day of transfection) (image under inverted microscope; 10×)

2. Measure the amount of nucleic acid obtained after isolation by a spectrophotometer. 3. Transfer agarose into gel electrophoresis with ethidium bromide for verification of the transposon quality (see Note 24). Use the DNA ladder as a standard against the isolate sample. 4. Visualize the agarose gel under UV lamp in dark conditions. 3.5 Transfection, Post-transfection Cell Recovery, and iPSC Cultivation

1. Mix the optimal number of cells in its suspension with the transfection kit and guide the piggyBac transposon (see Note 25) into the PBMCs with the use of an automatic electroporator, according to the manufacturer’s protocol. 2. Treat the transfected cells with unstimulated complete medium (see Note 26).

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Fig. 2 (a) Sample image of proper iPSC colony morphology (image under inverted microscope; 10×). (b) Sample image of proper iPSC colony morphology (image under inverted microscope; 10×)

3. Centrifuge the cell culture plates at 600 × g for 10 min (see Note 27). 4. Discard the unstimulated complete medium and replace it with the stimulated complete mononuclear cell culture medium (see Note 28). 5. Centrifuge the cell culture plates at 600 × g for 10 min. 6. Discard the stimulated complete mononuclear cell culture medium and replace it with stimulated complete mononuclear cell culture medium and human PSC-specific xeno-free and feeder-free medium (hybrid medium; 1:1) (see Note 29). 7. Centrifuge the cell culture plates at 600 × g for 10 min. 8. Discard the hybrid medium and replace it with the human PSC-specific xeno-free and feeder-free medium (Fig. 2a, b) (see Note 30). 3.6 iPSC Passaging, Cryopreservation, and Thawing

1. For passaging and cryopreservation, centrifuge the cell culture plates at 600 × g for 10 min. 2. Remove the supernatant (see Note 31). 3. Wash cells with DPBS. 4. Add the commercially available enzyme-free reagent for dissociation and passaging of PSCs. 5. Incubate at ambient temperature for 5–7 min. 6. Discard the commercially available enzyme-free reagent for dissociation and passaging of PSCs. 7. Incubate the remaining colonies at 37 °C for 10 min.

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8. For passaging, inoculate the cells onto new vitronectin-coated cell culture plates (in 1:2 or 1:3 ratio) and add the human PSC-specific xeno-free and feeder-free medium (see Note 32). 9. For cryopreservation, resuspend the cell pellet in appropriate volume of human PSC-specific xeno-free and feeder-free medium so that the optimal number of iPSC colonies are preserved per vial. 10. Prepare the iPSC cryopreservation medium inside the cryogenic vial. 11. Place the cryogenic vials inside a freezing container and perform interim storage overnight at -86 °C. 12. The next day, transfer the cryogenic vials into a liquid nitrogen (LN) tank for long-term storage. 13. For further processing, remove a cryogenic vial from the LN tank and swiftly thaw. 14. Transfer all of the colonies by aspirating dropwise with a small amount of human PSC-specific xeno-free and feeder-free medium. 15. Repeat step 8 for further passaging (see Note 33). 16. Samples for quality control, characterization, and molecular verification can be collected during the course of iPSC generation (see Note 34). We ideally collect samples at days 2, 5, 7, 10, 12, 20, 22, during further passaging and/or subculturing, pre-cryopreservation, and post-thawing (see Note 35).

4

Notes 1. Preferably use the sterile syringe filter of 0.22 μm. If not available, you may also use the 0.45 μm sterile syringe filter, however in this case, filtering each solution twice through it. 2. Before using any media or reagent, pre-warm inside a stainless steel water bath heated at 37 °C. 3. Isolation of PBMCs frequently manifests individual discrepancies regarding cell numbers. Should you suspect a risk of low numbers, you may prefer to collect 10 mL from each donor. 4. The donor must be comprehensively informed of the study in which they will enroll, and the written and signed consent provided in accordance with the relevant ethics committee approval must be received from each donor at all times. 5. We prefer using fresh PBS/FBS solution. 6. In case of shortage in commercially available density gradient, a standard laboratory density gradient such as sterile glycerol with a density of 1.26 g/mL can also be used.

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7. In case of a shortage in commercially available mononuclear cell culture medium, substitute with RPMI 1640 or minimum essential medium (MEM) with the same volumes of cytokines, growth factors, and other supplementary materials. Use 500 μL L-glutamine in cases where glutamine is not readily included in the content of RPMI 1640 or MEM. 8. Complete media of all sorts should be used up within 4–5 weeks for obtaining optimal culturing conditions. 9. Optimally, cryopreserve max. 20 × 106 cells per 1.5 mL cryogenic vial. 10. Safer, commercially available DNA stains can be preferred. 11. In our laboratory setting, we use automated electroporators with which transfection can be initiated with a cell number as low as 1–3 × 106 PBMCs. The minimal number of cells necessary for transfection initiation depends on the device, its transfection kits, and the recommended manufacturer’s protocols. 12. We obtained the best cell recovery results in the MEM condition. However, we have also observed that a similar recovery condition with the use of RPMI 1640 can also be used. 13. It is possible to substitute vitronectin with matrigel. 14. Use the human PSC-specific xeno-free and feeder-free medium as per the steps explained in Notes 26 to 30. 15. Use the ROCK inhibitor at the thawing step. 16. Optimally, cryopreserve 2–4 iPSC colonies per 1.5 mL cryogenic vial. In our laboratory, we generally use DMSO in the cryopreservation step. However, commercially available cryopreservation media for PSCs may be preferred for more efficient freezing and thawing. 17. Should there be a shortage of commercially available enzymefree reagent for dissociation and passaging of PSCs, the use of 0.5 mM ethylenediaminetetraacetic acid as a dissociation reagent is also possible. 18. Repeat steps 8–10, however this time centrifuging at 300 × g for 10 min (Accel = 0 Decel = 0). 19. Cell count is a necessity for monitoring the number of cells to be transfected, as it is a critical step in loading the minimum required number of cells to the automated electroporator. Cell count, and viability where necessary, can be performed by the use of the conventional trypan blue (%0.4) exclusion assay either manually on a hemocytometer or via an automated cell counter.

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20. The number of cells to be inoculated onto a cell culture plate may be estimated according to the manufacturer’s protocol of the preferred cell culture plates. 21. Our optimal culturing period for PBMCs is 2–4 days. 22. Swiftly perform all cryopreservation steps in cold temperature, preferably inside ice, at 4 °C. 23. At all thawing steps, act quickly and carefully to leave a small amount of DMSO inside the cryogenic vial after aspiration of all of the cell suspension. 24. Store the isolated piggyBac transposon at -20 °C until the day of transfection. 25. As per our experience, a piggyBac transposon concentration range of 0.5–3 μg/μL can be chosen for efficient transfection. 26. The optimal post-transfection cell recovery period in our laboratory is 24 h. Before cultivation, coat the cell culture plates with vitronectin, according to the manufacturer’s protocol. 27. As iPSCs are adherent cells, it is necessary to settle them with a mild centrifugation step before any intervention such as medium change, passaging, subculturing, and cryopreservation. 28. The optimal period of cultivation with the stimulated complete mononuclear cell culture medium is 3 days. 29. The optimal period of cultivation with the hybrid medium in our laboratory is 2 days. 30. Replace the hybrid medium with human PSC-specific xenofree and feeder-free medium on the 6th day following transfection. In our experience, the minimum period until initial passaging of iPSCs should be around 15–20 days (Pn). 31. In our laboratory, we do not discard the supernatant, we rather subculture (Nn) for a better yield of iPSC colonies. In our experience, the minimum period until initial subculturing of iPSCs should be no less than 10 days. 32. After each passaging and cryopreservation step, add the ROCK inhibitor and incubate overnight at 37 °C. The next day, centrifuge at 600 × g for 10 min and discard the supernatant. Wash cells with DPBS and add the human PSC-specific xenofree and feeder-free medium. 33. Medium change can be performed on every other day or every 3–4 days. This can be fixed according to monitored morphology changes in the colonies. 34. iPSCs are, in fact, fragile cells that need careful screening and handling. Morphology, intermittent immunoassays, or protein expression assays alone are not sufficient to give identity to iPSC colonies. For instance, expression analysis of the

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Fig. 3 (a) Sample image of distorted iPSC colony morphology (day 21 post-transfection) (image under inverted microscope; 10×). (b) Sample image of scattered iPSC colony morphology (day 26 post-transfection) (image under inverted microscope; 10×)

Fig. 4 (a) Sample image-1 of crater formation on iPSC colony morphology (day 21 post-transfection) (image under inverted microscope; 10×). (b) Sample image-2 of crater formation on iPSC colony morphology (day 22 post-transfection) (image under inverted microscope; 10×)

transcription factors may not necessarily mean that the iPSCs are in good quality. They may even manifest distorted morphology at the early stages of development (Figs. 3a, b and 4a, b). Further analyses for authentic identity, chromosomal shifts, stability factors, etc. may need to be further examined. 35. Any combination of iPSC quality testing such as alkaline phosphatase (AP) staining for rapid screening methods for pluripotent stem cell screening in terms of AP activity, immunofluorescent imaging, quantitative real-time polymerase chain reaction (qRT-PCR), RNA-seq and flow cytometry

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for rapid and efficient isolation, purity, expression of pluripotency markers and further evaluation of how the related genes are expressed, and the study of functional cell properties, G-banded karyotyping assays for detection of chromosomal abnormalities, short tandem repeat (STR) profiling for the authenticity of the iPSC lines in comparison with the parental line and/or donor tissue, and next generation sequencing (NGS) for screening genomic instability and identification of undifferentiated iPSC detection can be performed by proper sampling. It is also crucial to observe if the iPSC line will yield embryoid bodies (EBs) and differentiate into all the three germ layers, which are the ultimate verification parameters of iPSC lineage commitment.

Acknowledgments The creation of this protocol was supported by the project #2200299 Techno-entrepreneurship Technical Support Program (BI˙GG) Technoinvestment Capital Fund Project 1512 of the Sci€ ˙ TAK). entific and Technological Research Council of Turkey (TUBI References 1. Rivera T, Yuanyuan Z, Yuhui N et al (2020) Human-induced pluripotent stem cell culture methods under cGMP conditions. Curr Protoc Stem Cell Biol 54:e117. https://doi.org/10. 1002/cpsc.117 2. Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126(4):663–676. https://doi.org/10. 1016/j.cell.2006.07.024 3. Takahashi K, Tanabe K, Ohnuki M et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131: 861–872. https://doi.org/10.1016/j.cell. 2007.11.019 4. Doss MX, Agapios S (2019) Current challenges of iPSC-based disease modeling and therapeutic implications. Cell 8(5):403. https://doi.org/10.3390/cells8050403 5. Omole AE, Fakoya AOJ (2018) Ten years of progress and promise of induced pluripotent stem cells: historical origins, characteristics, mechanisms, limitations, and potential applications. PeerJ 6:e4370. https://doi.org/10. 7717/peerj.4370. eCollection 2018 6. Behl T, Kaur I, Sehgal A et al (2022) “Cutting the Mustard” with induced pluripotent stem cells: an overview and applications in healthcare

paradigm. Stem Cell Rev Rep 18:2757–2780. https://doi.org/10.1007/s12015-02210390-4 7. Nicholson MW, Ting CY, Chan DZH et al (2022) Utility of iPSC-derived cells for disease modeling, drug development, and cell therapy. Cell 11:1853. https://doi.org/10.3390/ cells11111853 8. Sharma A, Sances S, Workman MJ et al (2020) Multi-lineage human iPSC-derived platforms for disease modeling and drug discovery. Cell Stem Cell 26:309–329. https://doi.org/10. 1016/j.stem.2020.02.011 9. Rao MS, Malik N (2012) Assessing iPSC reprogramming methods for their suitability in translational medicine. J Cell Biochem 113: 3061–3068. https://doi.org/10.1002/jcb. 24183 10. Kuo HH, Gao X, DeKeyser JM et al (2020) Negligible-cost and weekend-free chemically defined human iPSC culture. Stem Cell Reports 14:256–270. https://doi.org/10. 1016/j.stemcr.2019.12.007 11. Uhrig M, Ezquer F, Ezquer M (2022) Improving cell recovery: freezing and thawing optimization of induced pluripotent stem cells. Cell 11(5):799. https://doi.org/10.3390/ cells11050799

piggyBac-Induced iPSC Protocol 12. Steeg R, Mueller SC, Mah N et al (2021) EBiSC best practice: how to ensure optimal generation, qualification, and distribution of iPSC lines. Stem Cell Reports 16(8): 1853–1867. https://doi.org/10.1016/j. stemcr.2021.07.009 13. Woltjen K, H€am€al€ainen R, Kibschull M et al (2011) Transgene-free production of pluripotent stem cells using piggyBac transposons. In: Schwartz P, Wesselschmidt R (eds) Human pluripotent stem cells. Methods in Molecular Biology, vol 767. Humana Press. https://doi. org/10.1007/978-1-61779-201-4_7 14. Yang W, Mills JA, Sullivan S et al (2012) iPSC reprogramming from human peripheral blood using Sendai virus mediated gene transfer. In: StemBook [Internet]. Cambridge, MA: Harvard Stem Cell Institute; 2008-. Available from: https://www.ncbi.nlm.nih. gov/books/NBK143766/. https://doi.org/ 10.3824/stembook.1.73.1

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15. Lorenzo IM, Fleischer A, Bachiller D (2013) Generation of mouse and human ınduced pluripotent stem cells (IPSC) from primary somatic cells. Stem Cell Rev Rep 9:435–450. https://doi.org/10.1007/s12015-0129412-5 16. Kim Y, Rim YA, Yi H et al (2016) The generation of human ınduced pluripotent stem cells from blood cells: an efficient protocol using serial plating of reprogrammed cells by centrifugation. Stem Cells Int:1329459. https:// doi.org/10.1155/2016/1329459 17. Singh A, Verma V, Yadav CB et al (2017) Induced pluripotent stem cell (iPSC) reprogramming protocols. In: Verma V, Singh MP, Kumar M (eds) Stem cells from culture dish to clinic. Nova Science Publishers, Inc 18. Gao L, Wang F, Wang Y et al (2022) A protocol for the generation of patient-specific iPSC lines from peripheral blood mononuclear cells. STAR Protocols 3(3):101530. https://doi. org/10.1016/j.xpro.2022.101530

Methods in Molecular Biology (2023) 2736: 207–223 DOI 10.1007/7651_2023_478 © Springer Science+Business Media, LLC 2023 Published online: 05 May 2023

Signaling Pathways in Trans-differentiation of Mesenchymal Stem Cells: Recent Advances Vaishak Kaviarasan, Dikshita Deka, Darshini Balaji, Surajit Pathak, and Antara Banerjee Abstract Mesenchymal stem cells are a group of multipotent cells that can be induced to differentiate into other cell types. The cells fate is decided by various signaling pathways, growth factors, and transcription factors in differentiation. The proper coordination of these factors will result in cell specification. MSCs are capable of being differentiated into osteogenic, chondrogenic, and adipogenic lineages. Different conditions induces the MSCs into particular phenotypes. The MSC trans-differentiation ensues as a response to environmental factors or due to circumstances that prove to favor trans-differentiation. Depending on the stage at which they are expressed, and the genetic alterations they undergo prior to their expression, transcription factors can accelerate the process of trans-differentiation. Further research has been conducted on the challenging aspect of MSCs being developed into non-mesenchymal lineage. The cells that are differentiated in this way maintain their stability even after being induced in animals. The recent advancements in the transdifferentiation capacities of MSCs on induction with chemicals, growth inducers, improved differentiation mediums, growth factors from plant extracts, and electrical stimulation are discussed in this paper. Signaling pathways have a great effect on MSCs trans-differentiation and they need to be better understood for their applications in therapeutic techniques. So, this paper tends to review the major signaling pathways that play a vital role in the trans-differentiation of MSC. Key words Genetic modification, Induced pluripotent stem cells, Mesenchymal stem cells, Signaling pathways, Therapies, Trans-differentiation

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Introduction Mesenchymal stem cells (MSCs) are a kind of multipotent stromal cells that has the capability to self-renew and differentiate into various other cell types. They are desirable candidates for scientific studies due to their ease of isolation, relatively rapid expansion rates, high migratory capability, and capacity to prevent allogeneic reactions following transplantation [1]. In recent times, they have been derived from a wide range of organs and tissues that include placental tissue, adipose tissue, testicles, dental pulp, menstrual blood, liver, pancreas, spleen, menstrual blood, amniotic fluid, umbilical cord blood, and the lung [2]. They are marked by their spindle-shaped morphological features, their capacity to

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differentiate in vitro into chondrocytes, osteocytes, and adipocytes. According to earlier studies, there isn’t a single unique marker which is used to characterize MSCs from other cells that have comparable fibroblastic properties. As a result, these cells exhibit either positive or negative expressions of several surface antigens, which defines their immunophenotype. MSCs lack hematological and endothelial markers including CD34, CD31, CD14, CD11, and CD45 but exhibit surface antigens like CD105, CD90, CD29, CD73, and CD44 [3]. MSCs can self-renew like other stem cells, but because of their multipotency and phenotype plasticity, it has been suggested that self-renewal of stem cells is less important for the functioning of mesenchymal tissues [4]. Owing to this plasticity and commitment, differentiation may be reversible by reacting to environmental cues, just as they may have retained a greater level of plasticity, thereby allowing them to switch from one lineage to other at a further point. For instance, this flexibility is crucial for the development and remodeling of bones [5]. When a committed cell type that has already started to differentiate along one pathway switches onto a distinct differentiation lineage, this process is known as trans-differentiation [6]. They were discovered to exhibit trans-differentiation plasticity when cultured in a variety of specific ways and when provocative compounds were added. As a result, it was discovered that populations and clonal lines of MSCs displayed ectodermal, mesodermal, and endodermal genes, indicating a much wider plasticity or stemness of MSCs than the predicted one. These observations of MSCs demonstrate their capacity to transdifferentiate into distinct forms of mesodermal, ectodermal, and endodermal cells when exposed to a particular concoction of growth factors and substrates [7]. Numerous pathways and transcriptional factors govern in vitro cell development and differentiation. Cell specification is the resultant of the carefully controlled usage of these elements [8]. This review tends to address the various signaling pathways that are linked with the transdifferentiation of MSCs and also the recent advancements in the respective field.

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Genetic Manipulation of MSC As previously stated, exposure to extrinsic signaling molecules can result in trans-differentiation of MSCs. Genetic modification is a different approach toward using MSCs for regenerative medicine [9]. These modifications can be accomplished through gene delivery, which uses transplanted engineered stem cells as a carrier for the transport of specific targets or by cell reprogramming, which employs knocked-down expression of genes to regulate the differentiation of the stem cell in a particular aspect. Stem cell genetic engineering is a sensitive and challenging process. Due to their

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remarkable tendency for proliferation, they do not make suitable entrant for nonviral episomal plasmid transfection or nucleofection. Mostly stem cells are susceptible to antibiotic selection, making it difficult to maintain transfected stem cells in a stable state. However, since certain studies have indicated that tumor transformation is a significant concern, one should be aware of the drawbacks of viral integration [10]. Bone marrow produced MSCs being investigated as possible carriers for the disease-specific cell and gene therapy. Due to their potential to localize to damage sites, they are promising entrants for the replacement or repair of organ systems and/or the administration of gene therapy products [11, 12]. Lentiviruses are the most effective vectors, according to comparative research comparing various viral transduction techniques of MSCs. Lentiviral gene delivery to rat MSCs has been demonstrated to be superior to other viral or nonviral gene delivery techniques by McMahon et al. [13]. In contrast to other viral and nonviral vectors, lentiviral vectors have been shown to be able to transduce human [14, 15] or mouse MSCs and maintain transgene expression for longer. These investigations also demonstrated that the transduction process does not adversely influence MSCs’ capacity for growth, differentiation, and migration and does not result in exorbitant cell death [16].

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Major Signaling Pathways Involved in MSC

3.1 WNT and βCatenin Pathways

A group of immensely conserved glycoproteins known as WNT proteins play crucial functions in cell growth and differentiation. Triggering of this pathway enhances the MSC trans-differentiation while diminishing the commitment to the adipocytic lineage [17, 18]. There seems to exist about 12 Frizzled receptors and more than 19 Wnt proteins in general. Each of the 19 Wnts plays a distinct role in the different developmental stages of bone production, such as Wnt7b, which is activated throughout osteoblastogenesis, and Wnt10b, which is released in bone marrow. It is the second significant signaling pathway that is engaged in osteoblastogenesis in two ways: canonical Wnt/β-catenin cascade and the non-canonical or -catenin-independent signaling cascade pathway. Non-canonical Wnt signaling pathway does not require β-catenin, but canonical Wnt signaling does [19–21]. In the beginning, the ligands were grouped into canonical and non-canonical groups based on the type of signal that was triggered when they bound to the receptors. Based on the pathophysiological context, certain ligands can activate Wnt signaling in a -catenin-dependent or independent fashion, depending on the kind of activation (canonical or not). For instance, Wnt5a, which was once categorized as a non-canonical signal, can both stimulate and suppress this signaling during the progression of both cancer and the embryo

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[22, 23]. Catenin-dependent Wnt signaling rises the bone mass, within which glycogen synthase kinase-3 is being restrained, and also enhances the proliferation of osteoblasts. Numerous studies indicated that the canonical Wnt signaling pathway boosts MSCs’ potential for self-renewal and reduces apoptosis. LRP5 overexpression, that is a crucial co-receptor that activates canonical Wnt signaling, was known to promote MSC proliferation [24]. There are various -catenin-independent Wnt signaling pathways, each of which is connected to a certain co-receptor or other essential components. One of such, the planar cell polarity (PCP) pathway, is important for tissue polarization and is mostly engaged in epithelial and mesenchymal cells. Wnt-PCP entails at least two complexes that are situated on the proximal and distal membranes of adjacent cells, accordingly [25]. Wnt/Ca2+ seems to be another well-known -catenin-independent pathway that regulates intracellular Ca2+ concentrations [26]. When Wnt binds to the co-receptors FZD and Ror2, non-canonical Wnt signaling is triggered. Several important components, including Rok, Dvl, DAG, Rho, and Ca + 2, activate this signaling pathway. By the phosphorylation of LEF/TCF transcription factors, that are essential components of the canonical signaling cascade, NLK blocks the canonical pathway [27]. This Wnt pathway’s regulation is crucial for MSC proliferation, self-renewal, and differentiation, among other aspects of cell fate determinations. To fully use the regenerative characteristics of MSCs in various scientific disciplines, including bone, lung, and heart biology, this signaling modulation in MSCs is specifically being extensively researched [28]. Recent data have demonstrated that the Wnt/β-catenin inhibitor quercetin limits the multipotent quality and proliferation of MSCs by promoting osteogenesis and inhibiting the differentiation of both adipocytes and chondrocytes [29]. 3.2

Notch Signaling

A key evolutionary mechanism that is conserved from flies to humans is referred to as Notch signaling [30]. Cell-cell contact can govern cell proliferation and/or differentiation into the various lineages by a pathway called Notch signaling pathway. Transmembrane receptor proteins called Notch receptors regulate distinct cell differentiation during embryogenesis and throughout life. It is believed that Notch components exist in MSCs as stemness signaling regulators. These receptors are comprised of transmembrane proteins (TM), proteins with the Notch intracellular domain (NICD), and Notch extracellular domain (NECD) which traverses from the cell that transmits the signal to the cell that acquires it. JAG1, JAG2, DLL3, DLL4, and DLL1 are the ligands for these receptors. Interaction of these ligands to the receptor complex causes NECD to be released from this complex. The complex protein will then translocate to the nucleus where it interacts with the target genes to initiate the classical Notch signaling cascade

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[31]. By inhibiting Runx2 activity, Notch signaling prevents osteoblast differentiation and preserves the bone marrow’s supply of mesenchymal progenitors. Normally, the FGF-dependent Notch signaling system regulates stem zone of spinal cord. Additionally, it increases spinal cord cell proliferation while never governing cell mobility outside the stem region. This signaling pathway assists the MSCs in differentiation of neurons [32]. Numerous studies portrayed that this pathway stimulates neurogenesis, and also under hypoxic conditions, promotes MSC osteogenic development [33]. Notch signaling is stimulated either as a paracrine signal to regulate contacts among different cell types or as a molecular event behind the differentiation of stem cells. Considering Notch engagement in stem cell differentiation, the osteoblast switch is the example. According to research, the Notch inhibitor DAPT may decrease ALP expression in MSCs that are encountering BMP9-dependent formation of osteoblasts, which would then result in lessened osteogenic differentiation both in vivo and in vitro. Meanwhile, DLL-1 treatment of MSCs increases the expression of ALP, OPN, and OCN [34, 35]. Several more investigations have supported Notch’s role in osteoblast differentiation. The lower expression of Notch signaling elements is related to adipocyte differentiation, addressing that Notch participation is lineage-dependent during MSC differentiation, i.e., inactivated for adipogenic differentiation and activated for osteogenic differentiation [36, 37]. 3.3 TGFβ/BMP Superfamily

The TGFβ superfamily is composed of over 30 secreted dimeric polypeptides that perform a critical role in the modulation of several cellular functions including differentiation, proliferation, and embryonic development. TGFβ1, TGFβ2, and TGFβ3 are the three isoforms that are largely conserved and share a group of conserved cysteine residues [38]. The TGF-superfamily of signaling molecules includes a set of proteins known as the BMPs that cause endochondral bone growth [39]. These secreted dimeric ligands have intracellular kinase activity that enables them to attach to cell surface receptors. Specific receptor-regulated SMAD (R-SMAD) proteins are phosphorylated as a result of ligand binding. While SMADs 1, 5, and 8 are phosphorylated by BMP signaling, SMADs 2 and 3 are specific for TGF- receptors. Following R-SMAD phosphorylation, CoSMAD (SMAD4) interacts with R-SMAD to produce an interaction complex that can go to the nucleus and control gene transcription [40]. Through I-SMAD proteins, which work in a negative feedback loop after being upregulated by TGF-β ligands, TGF-β/BMP signaling is controlled. By binding to receptors, preventing RSMAD phosphorylation, competing with R-SMADs and SMAD4 to form complexes, and speeding receptor turnover, I-SMADs can impair signal transduction [41, 42]. BMP signaling also has an impact on the formation of

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embryonic limbs and digits. Targeted BMP2 and BMP4 alterations in the limb mesenchyme stopped osteogenesis [43]. BMP/TGF-β signaling regulates MSC differentiation in vitro, along with its roles in the embryonic development of the mesodermal lineage. The important osteogenic transcription factor Runx2 is expressed as a result of BMPs. Runx2 is induced by BMP signaling through the osteogenic transcription factor Distal-less homolog (DLX5). DLX5 is precisely activated by BMP signaling, but not by other TNF-β members [44]. Runx2 interacts with SMAD 1 and SMAD 5 to perform a critical part in the transcriptional regulation of BMP-induced osteogenesis, even though DLX5 is a crucial modulator of BMP-induced osteogenesis. Runx2-SMAD complexes develop and are directed to particular nuclear matrix-associated locations where they control the target genes expression when BMP signaling is activated [45]. When the TGF-β-/BMP pathway is combined with other signaling pathways, it can have beneficial or adverse effects and finally result in desired biological effects [46]. Terminal differentiation, chondrocyte proliferation, ECM deposition, and mesenchymal condensation are only a few of the phases of chondrogenesis that are all governed by TGF-β [47]. 3.4

FGF Signaling

Fibroblast growth factors (FGFs) contain 23 members produced in almost all different tissues during development, which are crucial for differentiation of osteocytes and chondrocytes [48]. Their ligands are 20–35 kDa and adhere to the respective extracellular ligand-binding domain and also to a largely conserved intracellular signaling region that contains residues of tyrosine and intrinsic kinase [49]. On ligand interaction, FGFs form dimers, which trigger the autophosphorylation of the intrinsic kinase residues and the start of their signaling cascade. The FRS2 protein is then phosphorylated that also promotes the PKC, SAPK/JNK, p38 MAPK, ERK1/2, and PI3K MAPK pathways, bringing the Grb2/SOS complexes to the plasma membrane [50]. FGF ligands and receptors have a major impact on osteoblast differentiation. There is arguing proofs concerning the impacts of FGFs ligands on proliferation of osteoblasts. FGF-2 specifically promotes ALP activity in rat bone marrow precursor cells, while FGF-2, -4, and -8 promote Runx2 production. Matrix mineralization requires FGF-2, -9, and -18, while FGF-9 triggers the formation of the late osteogenic marker osteocalcin [51, 52]. Osteocyte differentiation and proliferation are both regulated by FGFR2, but osteogenic differentiation is predominantly regulated by FGFR1. FGF3 may be a key factor in osteogenesis and regulates chondrocyte proliferation during endochondral ossification [53]. FGFs are essential for chondrogenesis as well. The spatial and temporal control of FGF ligands and receptors is necessary for FGF signaling in chondrogenesis. FGFR3, however, is not produced in the central core of mesenchymal condensations

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where growing chondrocytes are, and as contrast to FGFR3, FGFR2 is expressed earlier in the condensing mesenchyme [54].

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Trans-differentiation Potential of MSC Trans-differentiation is the phenotypic transformation of one differentiated cell type into another. It has been thoroughly investigated and proven as a physiologic characteristic of amphibian species, like during limb regeneration and the transformation of pigmented epithelia into neural retinal cells. Since ASCs from vertebrates have been demonstrated to develop into cell types which do not typically appear in the dwelling organs or tissues, it will also take place in mammals. It has the potential for self-renewal, cloning and tissue differentiation distinguishing stem cells from other cell types [55]. The initial MSCs found more than 40 years ago formed deposits of bone and cartilage in culture, which is today acknowledged as a unique feature of MSCs [56]. Later studies carried out in the 1970s using autologous transplantation of BM-MSCs pellets to the subcapsular area of a rabbit kidney revealed that MSCs can develop in vivo into osteocytes. By establishing that MSC development into a specific phenotype is reliant on environmental factors [57], Caplan built on this valuable support. For instance, proximity to the vasculature is necessary for MSC differentiation into osteoblasts, but not for differentiation into chondrocytes. However, no vasculature is necessary for development into the chondrocyte [58]. In the end, these preliminary findings led the door for research that specified particular circumstances to support MSC differentiation [59]. Although there is an ample proof that show ASCs have the ability to transdifferentiate in mammalian systems, other discoveries, such as cell fusion and cell heterogeneity, have been used to contradict and disprove this idea [60]. In a recent work, it was examined whether differentiated hMSCs might retain their capacity for multi-differentiation in a well-defined cell culture system. Since only newly separated hMSCs were used in the in vitro culture system, it is possible to state that the trans-differentiation event witnessed was a true characteristic of differentiated hMSCs instead of the results of cell fusion. They also disregarded the possibility of contamination of the starting cell population by primitive or progenitor cells by studying a homogeneous population of fully developed osteoblasts [61]. These studies show that completely differentiated cells are more pliable than initially believed and that MSCs with predetermined commitment to a certain mesenchymal cell lineage could transdifferentiate into various cell types in reaction to inductive extracellular cues; such a characteristic supports a refined hierarchy of cell differentiation with trans-differentiation and differentiation processes [62]. Since a huge number of differentiated cells proliferate during the trans-

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differentiation process, it is possible that pre-committed cells undergo genome reprogramming to modify their phenotypes by dedifferentiating into a primitive stem cell phase. It is expected that controlling cell dedifferentiation and trans-differentiation will be crucial for the growth, upkeep, and regeneration of mammalian tissues [63].

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Transcription Factor-Induced Trans-differentiation of MSCs Reports on somatic and ESCs have demonstrated the value of genetically manipulating transcription factors (TFs) expression to promote and sustain differentiation. In particular for MSCs, genetic modification may be used to deliver TFs (alone or in combination with other extrinsic signaling molecules) to accelerate differentiation and broaden the plasticity spectrum of MSC. TFs have been administered to MSCs in a variety of ways, with varying degrees of success. To promote differentiation of MSC toward chondrogenic or osteogenic fate, TF administration was used in the early MSC/TF investigations [64]. In a subsequent study, it was observed that the induced expression of the Smad8 TF and bone morphogenic protein 2 was adequate enough in in vitro and in vivo system to develop neotendon from MSCs [65]. The goal of more difficult TF/MSC research after that was to get MSCs to develop into cell types unrelated to those of the mesenchymal lineage. The elevated levels pancreatic duodenal homeobox 1 (PDX1) transcription factor expression, marked the positive outcomes showing effective production of functional insulinproducing cells were described [66, 67]. Alluringly, our results from a study showed that TF-expressing MSCs might be used as an origin for β-cell replacement therapy using diabetes-related animal models. Notably, in keeping with the above-mentioned PDX1 findings, Ngn1-expressing MSCs were found to function as replacement neural cells in an animal stroke model since their transplanting dramatically improved motor function compared to that of MSCs that had not been genetically changed [68]. Several studies suggested that suppressing a TF, namely the neuronrestrictive silencer factor, could promote MSCs to develop into neural tissue [69, 70]. In conclusion, forced expression of individual TFs in MSCs can facilitate trans-differentiation into endothelial, neuronal, and pancreatic β-cells [66, 68]. Interestingly, transdifferentiated cells continue to display their identity after being excised from growth plates and injected into animals.

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6 TFs-Induced Direct Transformation of MSCs Lineage Versus Procreation of iPS Cells In earlier reports [62, 63], the conversion of MSCs into iPS cells was discussed. The ability of these reprogrammed MSCs to induce differentiation into all three types of germ layer cells is consistent with the pluripotent nature of iPS cells. But as we’ll see below, the information we have makes us believe that the former strategy will be more effective in terms of replacement therapy [71, 72]. MSCs seems to be an appealing candidate for this method of inducing trans-differentiation, especially in the context of the considerably low yield in the generation of iPS cells and data signifying the potential for reprogramming of lineage when TFs are displayed in cells from an origin similar to the target cell type [73–75]. As MSCs express regulatory genes that are involved in the stipulation of a variety of cell activities, fates, and functions [76], this is a result of the complexity of their transcriptome and may explain the flexibility of MSCs. Lineage transformation is a simpler protocol, technically speaking. For example, it’s plausible that going through the ESC-like condition is superfluous if one wants to instruct an MSC to transdifferentiate into a neuron. Instead, it’s likely that altering the balance between particular TFs in the transcriptome could more effectively steer the cell toward the intended neuronal fate than the iPS cell state. Second, there are at least two benefits to the lineage conversion strategy when taking into account potential clinical applications. More significantly, however, the iPS cell simulates as an ESC and is prone to developing teratomas after conversion, while technically merely serving as an emissary in the reprograming process [77].

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Mechanisms Behind Plasticity of Adult Mesenchyme The presence of mesenchymal cells is a crucial difference between the progression of fungi and plants and that of metazoans. For instance, localized cell proliferation and cell hypertrophy result in conservative shape modifications in plants. Contrarily, these cells contain motor proteins and a proteolytic toolbox which allows them to move around the embryo, engage and react to signals from the ECM, and become differentiated into certain components like cartilage, bone, muscle, etc. Moreover, it may be implied that epithelia are descended from mesenchyme because, both evolutionarily and ontogenetically, they come before mesenchyme [78]. The transition of epithelial to mesenchyme, that happens during the formation of vertebrates after gastrulation, is the process by which the primitive embryonic mesenchyme develops from epiblastic cells (EMT). Following this, early mesenchymal cells endure

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mesenchymal-to-epithelial conversions to rearrange and form secondary epithelial components like lateral and paraxial mesoderm [79]. Numerous organs, such as the kidney and heart, develop as a result of these conversions. As per recent experimental data, MSCs are thought to exist in vivo as modified marrow pericytes which cover the endothelium of the marrow sinuses [80]. Likewise, lineage studies showed that the most of perivascular cells and blood vessels are produced by Annexin A5 positive cells that turn into angioblasts [81]. These results agree with reliant studies providing the origin of endothelial cells and pericytes from vascular stem cells [82]. The splanchnic hemangioblast, a shared progenitor for endothelial and hematopoietic lineages, is thought to infiltrate the surrounding splanchnopleura as a result of the splanchnic mesothelium, undergoing an EMT [83]. As a result, there is convincing evidence that mesothelial cells are the source of MSCs at the beginning of their development. This developmental approach is in accordance with modern data indicating how cells from bone marrow and the stromal cells in the growing fetal liver that endure an EMT also help intestinal epithelial tumors [84, 85]. Researchers may therefore be more interested in identifying whether the cells inside adherent populations are true stem cells that preserve the capability to endure mesenchymal to epithelial transitions in vivo instead of focusing on whether MSCs display general developmental plasticity (like the potency to differentiate into various types of epithelial cells) [86] as shown in Fig. 1.

Fig. 1 Diagram illustrating the interactions between MSCs and cancer cells in the context of MSC-based therapies

Signaling Pathways in Trans-differentiation of Mesenchymal Stem Cells:. . .

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Recent Advancements in the Trans-differentiation Capacities of MSCs Over a million people in the world are affected by neurodegenerative disorders. Common examples include Alzheimer’s and Parkinson’s. They are caused by nerve cells in the brain that deteriorate in function and eventually die. They are incurable but the severity of the symptoms is reduced. So, stem cell therapy is being looked into as a promising source for the regeneration of these damaged neurons. Generally, bone marrow MSCs are used for neural regeneration which is accomplished via trans-differentiation. The MSCs obtained from the bone marrow are able to survive within the damaged brain and spinal cord tissue. Later, the MSCs get converted into the precursors of neurons which ameliorate the neurological condition of the person. MSCs have the ability to form different types of neurons such as peptidergic neurons, dopaminergic neurons, and cholinergic neurons. MSCs via chemical induction express markers such as GFAP, NSE, NF- 200, Tau, and NeuN, which are seen on neurons specifically. The MSCs procured from mesoderm are selected as they can be conveniently transdifferentiated into neuronal and glial cells on exposure to several growth inducers. MSCs derived from tooth pulp are being considered as a promising choice for future research on dopaminergic neurons. The menstrual blood–derived MSCs were injected into rats for treatment of incomplete spinal cord injury, and it was observed that their differentiation showed better motor functioning of hind limb [87]. MSCs derived from adipose tissue are differentiated using two differentiation media and the results obtained are elliptical-/spherical-shaped cells. Another differentiation medium was used which included DMEM, FBS, antibiotics, FGF2, EGF, BMP-9, and retinoic acid, which also gave the above results. When these cells further underwent trans-differentiation in appropriate differentiation medium to give Schwann cells, an increased number of cells were observed. Peripheral nerve (PN) injuries occur when the peripheral nerves are damaged due to accidents and in conditions like diabetes or autoimmune diseases. The peripheral nervous system has the ability to regenerate its axons but its unpremeditated response doesn’t give the best outcomes. So the approach toward the treatment of PN injuries comprises autologous nerve grafts but these face a lot of complications. The motor and sensory axons of peripheral nerves are surrounded by myelin sheath which is made up of Schwann cells. These Schwann cells can mend tissues, release neutrophic factors like NGF, BDNF, and GDNF, and restore the neurons and glial support cells for nerve regeneration. It was found that MSCs or ADSCs transdifferentiated into Schwann-like cells when exposed to the nerve lesion environment due to the trophic factors released here. SC-like cells were also obtained by dissociating neurospheres which were formed by

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adipose-derived MSCs. The dissociation of neurospheres is done by culturing them with different factors like bFGF, EGF, and B27 supplement and subsequent mitogen withdrawal. These SC-like cells resulted in myelination and regeneration of damaged nerves. hMSCs were electrically stimulated and differentiated into Schwann cells by passing biphasic electric current for up to four weeks. This showed improvement in the axonal outgrowth and activity of Schwann cells. Similarly, electrical stimuli at spinal cord injury sites accompanied by implantation of MSCs resulted in restricted recovery. The effect of extremely low-frequency electromagnetic fields induces the trans-differentiation of bone-marrowderived MSCs into neural cells. If cholinergic neurons and axons of the basal forebrain in the cortex are affected, it causes progressive deterioration in memory and learning skills which causes Alzheimer’s disease. Human mesenchymal stromal cells (hMSCs) have the ability to differentiate into cholinergic-like neurons (ChLNs). They’re also capable of differentiating into osteoblasts and adipocytes [88]. A newly improved medium called cholinergic-N-Run achieves the trans-differentiation of human Wharton’s jelly-derived mesenchymal stromal cells (WJ–MSCs) into cholinergic-like neurons in just 4 days. MSCs express cholinergic receptors as well as neuronal markers like TUC-4, NF-L, MAP2, and β-tubulin III. MSCs are capable of forming mesoderm and ectoderm germ line and their behavior might be similar to neuronal precursor cells. These findings suggest that neural stem cells and WJ–MSCs may experience the same regulatory mechanisms, and the transdifferentiation of MSCs into ChLNs might be influenced by the medium cholinergic-N-Run. It has been noted that the medium contains bFGF, Hep, SHH, RA, and very low concentration of FBS in DMEM/F12. Hep is found to be capable of pushing the transdifferentiation of MSCs in the direction of ectoderm and neuro epithelium. It is also hypothesized that Hep/bFGF2 can reduce the time taken for the formation of ChLNs. WJ-MSCs were preprogramed to form neural cells, which is proved by the expression of TUC-4, NF-L, β-tubulin III, and MAP2 proteins. RA is a component of the medium and in its presence, WJ–MSCs specifically differentiated into cholinergic-like phenotype and it is also said to accelerate the differentiation process [89]. MSCs derived from human gingiva have several great characteristics such as easy accessibility and high regenerative capacity and has shown positive effects on cutaneous wound healing and skin regeneration. It is hypothesized that HGMSCs can be transdifferentiated into keratinocyte-like cells (KLC) with the help of critical growth factors present in plant extracts. This was applied in developing a wound healing tissue graft which encapsulated the HGMSCs in bioengineered tissue constructs. HGMSCs are capable of differentiating into adipogenic, osteogenic, and chondrogenic tissues. HGMSCs were treated with Acalypha indica and the initial

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Fig. 2 Representation of the multifaceted function of MSC secretome as a therapeutic strategy

spindle-shaped fibroblast was converted to polygonal cobblestoneshaped KLC. This was accompanied by expression of markers, such as cytokeratin 10 (5.4-fold), cytokeratin 14 (6.2-fold), and cytokeratin 5(4.3-fold) [90] as depicted in Fig. 2.

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Conclusion A wide variety of regulatory proteins are expressed by MSC populations, which is a reflection of the complexity of the bone marrow organ system. These proteins contribute to MSCs’ wide therapeutic efficacy but might influence their ability to trans-differentiate. Because MSCs are a sustainable supply for autologous stem cell transplantation, many studies attribute MSCs with therapeutic promise. Due to the scant evidence of trans-differentiation or the ability of transplanted MSCs to replace damaged or lost cells in a functional manner, the beneficial effect of MSC transplantation was previously thought to be supportive rather than substitutive. However, a growing amount of evidence points to the possibility of genetically engineering tissue-specific stem cells to express TFs in order to compel them to prevail the limitations of lineage restriction. In fact, employing this method, MSCs have been made to develop into cells that resemble pancreatic β-cells, dopaminergic neurons, and other particular cell types. These results are encouraging, but more research is needed to see whether these cells can be used as a reliable autologous source to treat conditions like Parkinson’s disease, diabetes mellitus, and cancer.

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Acknowledgments The authors of this paper are thankful to Chettinad Academy of Research and Education (CARE) for providing infrastructure support and to DST (INSPIRE), Government of India, and CARE for the financial support. Author Contributions The study was designed by AB and the literature search and manuscript draft preparation were done by VK, DD, DB, AB. The final version of the manuscript was reviewed and edited by SP and AB.

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INDEX A

F

Amyotrophic lateral sclerosis (ALS).................... 139–150 Astrogenesis................................................................... 102

Fate choice......................................................................... 1 Fibrin .........................................................................85–92 Ficoll-Paque density gradient ................. 79–81, 153, 154 Flow cytometry ........................................ 58, 59, 91, 113, 122, 123, 152, 154, 158, 159, 174, 175, 203

B Bioluminescence.......................................... 178, 183, 188 Blood-derived materials .................................................. 86 Brain tumors stem cells........................................ 177–191 Brown adipose tissue ........................................... 115–124

C Cell encapsulation .....................................................85–92 Cell fate........................................... 1, 5, 31, 39, 165, 210 Cell therapy .....................................................77, 86, 128, 129, 164, 209, 217 Chromosomal analysis ..............................................65–75 Cleavage Under Targets & Release Using Nuclease (CUT&RUN) ................................................. 9–20 Clonal ...................................................... 66, 95–103, 208 Colony-forming unit (CFU) ................ 66, 68, 70–72, 75 Commitment ...................................................1–6, 24, 40, 95, 204, 208, 209, 213 Cryopreservation........................ 123, 195–197, 199–202

D Development .............................1, 2, 5, 9, 10, 23, 24, 40, 54, 63, 67, 101, 106, 107, 127–136, 140, 165, 166, 172, 174, 193, 203, 208, 211–213, 216 Diabetes mellitus (DM) .............................. 128, 129, 219 Disease modeling ...........................................39, 105–113

E Electroporation .................................................... 193–204 Embryoid bodies (EBs) ............................. 41, 44–46, 49, 107, 108, 110–112, 204 Epidermis............................................................ 10, 12, 13 Epigenetics .....................................................9, 10, 40, 66 E14TG2a cell line .....................................................41, 42 Exosomes.............................................................. 163–175 Extracellular vesicles (EVs).................................. 165, 168

G GapmeR.....................................................................23–36 Gene silencing ...........................................................23–36 Genetic modification......................................66, 208, 214 Glioblastoma (GB)............................................... 177, 178

H Hematopoietic lineages ................................... 65–75, 216 Hematopoietic stem cells (HSCs)...........................65–75, 151, 155 Histone modifications................................................. 9–20 HSCs and progenitors ..............................................65–75 Human-induced pluripotent stem cells (hiPSCs) ......................23–36, 107, 141, 146, 194

I Induced pluripotent stem cells (iPSCs) ...........24, 39, 77, 107–113, 116, 144, 193–204 In vitro differentiation ........................... 23, 24, 108–109, 123, 175, 208, 211, 212 In vivo imaging system (IVIS) ........................... 178, 179, 184–188, 190 Islet endocrine cells......................................................... 23 Isolation ..................................... 9, 12–14, 19, 49, 53–63, 68, 69, 77, 80, 81, 119, 120, 124, 151–160, 166, 167, 171, 180, 194–198, 200, 204, 207

K Karyotyping ................................ 66, 68, 71–73, 174, 204

L Leukemia ................................................. 66, 78, 151, 152 Leukemia stem cells (LSCs) ................................ 151–160

Kursad Turksen (ed.), Stem Cells and Lineage Commitment: Methods and Protocols, Methods in Molecular Biology, vol. 2736, https://doi.org/10.1007/978-1-0716-3537-7, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

225

STEM CELLS AND LINEAGE COMMITMENT: METHODS AND PROTOCOLS

226 Index

Lineage .................................................................. 1, 2, 39, 46, 65–75, 78, 86, 95, 123, 178, 208, 210, 212–216, 219 Lineage commitment .....................................1–6, 24, 204 Lineage tracing.............................................................. 1–6

M Magnetic resonance imaging (MRI)...........178, 187–189 Magnetic selection (MS)...................................... 157–160 Mature neurons...................................................... 41, 111 Mesenchymal stem cell-derived exosomes (MSC-EXO) ............................................. 163–175 Mesenchymal stem cells (MSCs)................77–83, 85–92, 115–124, 164–167, 170, 173–175, 207–219 Midbrain dopaminergic neurons (mDMA) ...........................................108–109, 111 Modeling .......................................................39, 105–113, 139–150, 208 Mouse embryonic stem cells (mESCs) ....................39–50 Mouse models ............................................................... 117 Myoepithelial cells.....................................................53–63

N Neural stem cells (NSCs)...................... 98–103, 177, 178 Neurogenesis ................................................................. 211 Neuromuscular junction (NMJ) ......................... 139–150 Neuronal differentiation .................................... 40, 41, 44 Neuronal progenitor cells (NPCs) ..............41, 45–47, 49

Platelet-rich plasma (PRP)........................................85–92 Primary culture..................................57–60, 63, 121, 170

R Regenerative medicine ......................................39, 78, 87, 116, 165, 172, 174, 208 Reporter gene................................................................ 178 Retinoic acid (RA) ............................................27, 29, 30, 39–50, 142, 144, 217, 218

S Salivary glands .................................................... 53, 54, 58 Signaling pathways .......................................151, 207–219 Single-cell transcriptomics ............................................ 1–6 Sphere .......................................57, 58, 62, 181, 183, 190 Spheroid.........................54, 62, 141, 143, 144, 146–149 Standard operating procedure (SOP) .................. 68, 110, 113, 115–124, 170, 174, 175 Stem cell ............................. 1, 10, 23, 40, 53, 66, 77, 86, 95, 107, 116, 128, 140, 152, 164, 177, 193, 208 Stem cell-based models........................................ 105–113 Stem cell culture................................................... 177–191 Surface markers ....................................... 53, 78, 165, 175

T

Optical imaging............................................................. 174 Organ-on-a-chip (OOC) .............................................. 141

Therapies ................................ 77, 78, 86, 106, 116, 128, 129, 152, 163–175, 177, 178, 193, 209, 214–217 Tissue engineering .......................................................... 78 Trans-differentiation ............................................ 207–219 Transfection............................................1, 24, 33, 35, 36, 61, 62, 194–199, 201, 202, 209 Type 2 diabetes mellitus (T2DM) ............................... 128

P

U

Pancreatic endocrine progenitors................................... 24 Parkinson’s disease (PD) ..................................... 105–113 Passaging .............................................181, 196, 199–202 Patient-derived xenografts (PDX)....................... 177–191 Peripheral blood mononuclear cells (PBMCs) .................................. 194–198, 200–202 piggyBac .........................................................58, 193–204 piggyBac transposon vector system................................ 54

Umbilical cord blood..................... 77–83, 116, 160, 207

O

W Wharton’s jelly ........................................ 77–83, 116, 218

Z Zebrafish ............................................................... 127–136