Mouse Genetics: Methods and Protocols (Methods in Molecular Biology, 2224) 1071610074, 9781071610077

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Methods in Molecular Biology 2224

Shree Ram Singh Robert M. Hoffman Amit Singh Editors

Mouse Genetics Methods and Protocols Second Edition

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.

Mouse Genetics Methods and Protocols Second Edition

Edited by

Shree Ram Singh Basic Research Laboratory, National Cancer Institute, Frederick, MD, USA

Robert M. Hoffman AntiCancer, Inc., San Diego, CA, USA; Department of Surgery, University of California, San Diego, CA, USA

Amit Singh Department of Biology, University of Dayton, Dayton, OH, USA

Editors Shree Ram Singh Basic Research Laboratory National Cancer Institute Frederick, MD, USA Amit Singh Department of Biology University of Dayton Dayton, OH, USA

Robert M. Hoffman AntiCancer, Inc. San Diego, CA, USA Department of Surgery University of California San Diego, CA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1007-7 ISBN 978-1-0716-1008-4 (eBook) https://doi.org/10.1007/978-1-0716-1008-4 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Illustration Caption: See Chapter 2, Figure 1 for more details. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Mice and humans share an incredible level of anatomical, physiological, and especially genomic resemblance. Genetically engineered mice are important tools for understanding gene function and modeling various metabolic diseases and cancer. Modeling human disease in mice offers various advantages over other mammalian animal models because mice are small, easy to breed, available in large number of inbred strains, and their genome is fully sequenced. Mouse genetics study provides a basic knowledge of gene function, genetic networks, pathology, preclinical modeling, and development of novel therapeutic targets and precision medicine. Various genetic tools have been developed, such as technologies that efficiently edit the mouse genome in vivo. Gene knockout and knockin or conditional gene modification help in temporal-spatial regulation and cell lineage tracing. Recent discovery of the CRISPR/Cas9 gene editing system revolutionized the genetics field. Over the years, almost all human diseases have been modeled in mice. This second volume of Mouse Genetics: Methods and Protocols provides selected mouse genetic techniques and their application in modeling varieties of human diseases. The chapters are mainly focused on the generation of different transgenic mice to accomplish the manipulation of genes of interest, tracing cell lineages, and modeling human diseases. Composed in the highly successful Methods in Molecular Biology series format, each chapter contains a brief introduction, a list of necessary materials, systematic methods, and a notes section, which shares tips on troubleshooting to avoid known pitfalls. We hope that this second volume of Mouse Genetics: Methods and Protocols will provide fundamental techniques and protocols to geneticists, molecular biologists, cell and developmental biologists, students, and postdoctoral fellows working in the various disciplines of genetics, developmental biology, mouse genetics, and modeling human diseases. We would like to thank Prof. John M. Walker and the staff at Springer for their assistance throughout the preparation of the book for publication. We also would like to express our sincere appreciation and gratitude to the contributors for sharing their precious laboratory expertise with the mouse community. Finally, yet importantly, we would like to thank our family members for their continued support. Frederick, MD, USA San Diego, CA, USA Dayton, OH, USA

Shree Ram Singh Robert M. Hoffman Amit Singh

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

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1 Generation of Mouse Model (KI and CKO) via Easi-CRISPR . . . . . . . . . . . . . . . . Dorjee T. N. Shola, Chingwen Yang, Chiayun Han, Rada Norinsky, and Ruben D. Peraza 2 Analysis of Gene Expression Using lacZ Reporter Mouse Lines . . . . . . . . . . . . . . . Michael Simon Kr€ a mer, Robert Feil, and Hannes Schmidt 3 Linear Density Sucrose Gradients to Study Mitoribosomal Biogenesis in Tissue-Specific Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Benedetta Ruzzenente and Metodi D. Metodiev 4 Mouse Models for Studying Hippocampal Adult Neural Stem Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fatih Semerci, Luke Parkitny, and Mirjana Maletic-Savatic 5 Retina as a Model to Study In Vivo Transmission of α-Synuclein in the A53T Mouse Model of Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . Najiba Mammadova, Thierry Baron, Je´re´my Verche`re, Justin J. Greenlee, and M. Heather West Greenlee 6 Promoting Pro-Endocrine Differentiation and Graft Maturation Following Surgical Resection of the Mouse Pancreas . . . . . . . . . . . . . . . . . . . . . . . . Mugdha V. Joglekar, Rohan R. Patil, Sarang N. Satoor, Wilson K. M. Wong, Mahesh S. Karandikar, and Anandwardhan A. Hardikar 7 Color-Coded Imaging of Cancer and Stromal Cells Interaction in the Pancreatic Cancer Tumor Microenvironment (TME) . . . . . . . . . . . . . . . . . . Atsushi Suetsugu and Robert M. Hoffman 8 Generating Ins2+/ /miR-133aTg Mice to Model miRNA-Driven Cardioprotection of Human Diabetic Heart. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hamid R. Shahshahan, Tyler N. Kambis, Sumit Kar, Santosh K. Yadav, and Paras K. Mishra 9 Generating a Podocyte-Specific Neonatal F Receptor (FcRn) Knockout Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Judith Blaine 10 Mouse Models of Colitis-Associated Colon Cancer . . . . . . . . . . . . . . . . . . . . . . . . . Santhakumar Manicassamy, Puttur D. Prasad, and Daniel Swafford 11 Generation of Colon Cancer Model Based on Colonoscopy Injection . . . . . . . . . Suhwan Chang 12 Generation of Transgenic Fluorescent Reporter Lines for Studying Hematopoietic Development in the Mouse. . . . . . . . . . . . . . . . . . . . . Jeffrey Barminko, Andrei M. Vacaru, and Margaret H. Baron

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Contents

Gene Inactivation in Adult Long-Term Hematopoietic Stem Cells Using Inducible Mouse Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 Kristbjorn Orri Gudmundsson Hematological Humanization of Immune-Deficient Mice . . . . . . . . . . . . . . . . . . . 195 Marina Gergues, Seda Ayer, Sara Morelli, Steven J. Greco, and Pranela Rameshwar Antisense Oligonucleotide Treatment in a Humanized Mouse Model of Duchenne Muscular Dystrophy and Highly Sensitive Detection of Dystrophin Using Western Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . 203 Rika Maruyama and Toshifumi Yokota

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

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Contributors SEDA AYER • Department of Medicine, Rutgers, New Jersey Medical School, Newark, NJ, USA JEFFREY BARMINKO • Department of Medicine, The Tisch Cancer Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA MARGARET H. BARON • Departments of Medicine, Cell, Developmental, and Regenerative Biology, and Oncological Sciences, The Tisch Cancer Institute, The Black Family Stem Cell Institute, and The Graduate School of Biomedical Sciences, Icahn School of Medicine at Mount Sinai, New York, NY, USA THIERRY BARON • Anses, Laboratoire de Lyon, Unite´ Maladies Neurode´ge´ne´ratives, Lyon, France JUDITH BLAINE • Division of Renal Diseases and Hypertension, University of Colorado, Anschutz Medical Campus, Aurora, CO, USA SUHWAN CHANG • Department of Biomedical Sciences, Physiology, College of Medicine, Asan Medical Center, University of Ulsan, Seoul, South Korea ROBERT FEIL • Interfaculty Institute of Biochemistry, Eberhard Karls University Tu¨bingen, Tu¨bingen, Germany MARINA GERGUES • Department of Medicine, Rutgers, New Jersey Medical School, Newark, NJ, USA; Rutgers School of Graduate Studies at New Jersey Medical School, Newark, NJ, USA STEVEN J. GRECO • Department of Medicine, Rutgers, New Jersey Medical School, Newark, NJ, USA JUSTIN J. GREENLEE • Virus and Prion Research Unit, United States Department of Agriculture, National Animal Disease Center, Agricultural Research Service, Ames, IA, USA M. HEATHER WEST GREENLEE • Department of Biomedical Sciences, Iowa State University College of Veterinary Medicine, Ames, IA, USA KRISTBJORN ORRI GUDMUNDSSON • Hematopoiesis and Stem Cell Biology Section, Mouse Cancer Genetics Program, Center for Cancer Research-National Cancer Institute, Frederick, MD, USA; Basic Research Program, Frederick National Laboratory for Cancer Research, Leidos Biomedical Research Inc., Frederick, MD, USA CHIAYUN HAN • CRISPR and Genome Editing Resource Center, The Rockefeller University, New York, NY, USA ANANDWARDHAN A. HARDIKAR • Diabetes and Islet Biology Group, School of Medicine, Western Sydney University, Campbelltown, NSW, Australia ROBERT M. HOFFMAN • AntiCancer, Inc., San Diego, CA, USA; Department of Surgery, University of California, San Diego, CA, USA MUGDHA V. JOGLEKAR • Diabetes and Islet Biology Group, School of Medicine, Western Sydney University, Campbelltown, NSW, Australia TYLER N. KAMBIS • Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA MAHESH S. KARANDIKAR • Department of Physiology, DY Patil Medical College, DY Patil University, Pune, India

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SUMIT KAR • Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA € MICHAEL SIMON KRAMER • Interfaculty Institute of Biochemistry, Eberhard Karls University Tu¨bingen, Tu¨bingen, Germany MIRJANA MALETIC-SAVATIC • Department of Pediatrics, Baylor College of Medicine, Houston, TX, USA; Jan and Dan Duncan Neurological Research Institute at Texas Children’s Hospital, Houston, TX, USA; Department of Neuroscience, Baylor College of Medicine, Houston, TX, USA NAJIBA MAMMADOVA • Virus and Prion Research Unit, United States Department of Agriculture, National Animal Disease Center, Agricultural Research Service, Ames, IA, USA SANTHAKUMAR MANICASSAMY • Georgia Cancer Center, Augusta University, Augusta, GA, USA; Department of Biochemistry and Molecular Biology, Medical College of Georgia, Augusta University, Augusta, GA, USA; Department of Medicine, Medical College of Georgia, Augusta University, Augusta, GA, USA RIKA MARUYAMA • Faculty of Medicine and Dentistry, Department of Medical Genetics, University of Alberta, Edmonton, AB, Canada METODI D. METODIEV • Laboratory for Genetics of Mitochondrial Disorders, INSERM U1163, Universite´ Paris Descartes-Sorbonne Paris Cite´, Institut Imagine, Paris, France PARAS K. MISHRA • Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA SARA MORELLI • Department of Obstetrics, Gynecology and Women’s Health, Rutgers, New Jersey Medical School, Newark, NJ, USA RADA NORINSKY • Transgenic and Reproductive Technology Center, The Rockefeller University, New York, NY, USA LUKE PARKITNY • Department of Pediatrics, Baylor College of Medicine, Houston, TX, USA; Jan and Dan Duncan Neurological Research Institute at Texas Children’s Hospital, Houston, TX, USA ROHAN R. PATIL • DY Patil Medical College, DY Patil University, Pune, India RUBEN D. PERAZA • Laboratory of Molecular Genetics and Immunology, The Rockefeller University, New York, NY, USA PUTTUR D. PRASAD • Department of Biochemistry and Molecular Biology, Medical College of Georgia, Augusta University, Augusta, GA, USA PRANELA RAMESHWAR • Department of Medicine, Rutgers, New Jersey Medical School, Newark, NJ, USA; Rutgers School of Graduate Studies at New Jersey Medical School, Newark, NJ, USA BENEDETTA RUZZENENTE • Laboratory for Genetics of Mitochondrial Disorders, INSERM U1163, Universite´ Paris Descartes-Sorbonne Paris Cite´, Institut Imagine, Paris, France SARANG N. SATOOR • DNA Sequencing Facility, National Centre for Cell Science, Pune, India HANNES SCHMIDT • Interfaculty Institute of Biochemistry, Eberhard Karls University Tu¨bingen, Tu¨bingen, Germany FATIH SEMERCI • Department of Pediatrics, Baylor College of Medicine, Houston, TX, USA; Jan and Dan Duncan Neurological Research Institute at Texas Children’s Hospital, Houston, TX, USA HAMID R. SHAHSHAHAN • Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA

Contributors

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DORJEE T. N. SHOLA • CRISPR and Genome Editing Resource Center, The Rockefeller University, New York, NY, USA ATSUSHI SUETSUGU • Gifu University Graduate School of Medicine, Gifu, Japan; AntiCancer, Inc., San Diego, CA, USA; Department of Surgery, University of California, San Diego, CA, USA DANIEL SWAFFORD • Georgia Cancer Center, Augusta University, Augusta, GA, USA ANDREI M. VACARU • Department of Medicine, The Tisch Cancer Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Institute for Cellular Biology and Pathology, Bucharest, Romania JE´RE´MY VERCHE`RE • Anses, Laboratoire de Lyon, Unite´ Maladies Neurode´ge´ne´ratives, Lyon, France WILSON K. M. WONG • Diabetes and Islet Biology Group, School of Medicine, Western Sydney University, Campbelltown, NSW, Australia SANTOSH K. YADAV • Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA CHINGWEN YANG • CRISPR and Genome Editing Resource Center, The Rockefeller University, New York, NY, USA TOSHIFUMI YOKOTA • Faculty of Medicine and Dentistry, Department of Medical Genetics, University of Alberta, Edmonton, AB, Canada; The Friends of Garrett Cumming Research & Muscular Dystrophy Canada HM Toupin Neurological Science Research Chair, Edmonton, AB, Canada

Chapter 1 Generation of Mouse Model (KI and CKO) via Easi-CRISPR Dorjee T. N. Shola, Chingwen Yang, Chiayun Han, Rada Norinsky, and Ruben D. Peraza Abstract Recent development of Easi-CRISPR (Efficient additions with ssDNA inserts-CRISPR) that utilizes long single-stranded DNA (lssDNA) of 0.2–2 kbases in length as donor templates to insert large segments of novel DNA sequences or to replace endogenous genes at precise locations in the genome has enabled CRISPR-assisted genome editing to make strides toward a more simple and rapid workflow. By leveraging the notion that short single-stranded DNA oligo (10 PCR (50 μL volume) to get enough lssDNA (see Note 6). The final lssDNA yield would depend on multiple factors (e.g., obtaining single-band starting PCR product with robust amount is essential); the acquired lssDNA can be validated through size separation on agarose gel as described in Subheading 3.4. 3.3.2 Design and Generation of LssDNA for Exon-Floxed CKO Model (GENE-Y)

Based on the CRISPR guides selected (guide-C and guide-D), the donor template for creating the CKO allele for GENE-Y consists of a exon 2 floxed cassette, flanked with 50 HA (longer) and 30 HA (shorter) that contain upstream genomic sequence of guide-C and downstream genomic sequence of guide-D, respectively (see Fig. 2b). Due to the presence of various segments of repetitive sequences, including different types of SINEs (short interspersed nuclear elements) and LTRs (long terminal repeats) in the intron 1 and intron 2 regions, we extend the length of 30 HA to 400 bp and 50 HA to 500 bp to span those unusual repeats with enough good sequences. Furthermore, since LoxP sequence lacks credible RE sites, we additionally introduce two unique RE cut sites (XbaI and EcoRV) adjacent to LoxPs into the donor for genotyping purpose. The total length of the donor DNA is 2.2 kb. To ensure the final assembled donor not to carry any undesired splicing elements, GENESCAN software can be used for verification. We obtained the dsDNA template (anchored in pUC57 backbone) through gene synthesis via vendor (Genewiz). The optimal lssDNA donor for efficient HDR, based on the CRISPR gRNA designed, is the top (sense) strand of the dsDNA vector (see Figs. 2b and 5a) (reasoned in Subheading 3.3). We initially attempted to obtain the lssDNA template by using Takara’s kit, but kept encountering multiple nonspecific DNA fragments being amplified, even with numerous adjustments to primer pairs and PCR conditions. As intron 1 and intron 2 regions of the GENE-Y are patched with multiple segments of unusual repeats, which tends to cause PCR amplification issues as evidenced, we therefore adopted a PCR-free route to directly retrieve lssDNA from the plasmid vector through CRIPSR-CLIP method [10]. Specifically, Cpf1 and Cas9n (D10A mutant nickase version of Cas9) with the corresponding gRNAs are used to create a dsDNA cleavage and a nick, respectively, on the donor-carrying plasmid at two junction sites flanking the lssDNA cassette, where Cas9n is particularly exerted on the strand of interest (see Fig. 5b). Upon the incisions, the plasmid results in three stand-alone ssDNA units,

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Spacer-NGG (SpCas9 PAM) TTTN (Cpf1 PAM)-Spacer

Fig. 5 Design and generation of lssDNA donor using CRISPR-CLIP (GENE-Y). (a) Donor DNA anchored in the default plasmid. The sense ssDNA (top stand) is the donor template of choice (lssDNA). (b) CRISPR-CLIP uses Cpf1 to create a dsDNA incision on one end of the lssDNA cassette while Cas9n(D10A) creates an ssDNA incision on the other end; Cas9n is applied on the strand to be procured. (c) Multiple reverse primers (red bent arrow) can be used to sequence the lssDNA (sense DNA) acquired (~50 bases at the 30 end cannot be read due to Sanger sequencing limitation). (d) Upon incisions by Cpf1 and Cas9n, followed by DGLB treatment, the entire plasmid resolved (in 0.9% agarose gel electrophoresis) into three stand-alone ssDNA units with distinct sizes 4.9 kbases, 2.7 kbases, and 2.2 kbases (lane 5B)

Generation of Mouse Model via Easi-CRISPR

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which are of distinct sizes hence are able to be separated on agarose gel electrophoresis upon denaturing gel-loading buffer (DGLB) treatment (see Fig. 5d). The target strand of our interest (2.2kbase lssDNA donor) can thus be identified and clipped out through the gel extraction procedure. 1. The assembled donor DNA for exon 2 floxing contains 50 HA (500 bp), XbaI+LoxP+exon 2+LoxP+EcoRV (1300 bp), and 30 HA (400 bp). The beginning of 50 HA and toward the end of 30 HA contain Cas9 PAM and Cpf1 PAM, respectively. 2. To digest the donor DNA plasmid, prepare both ctRNPCas9n (crRNA + tracrRNA + Cas9n Protein) and cRNPCpf1 (crRNA + Cpf1 Protein) complex first: the ratio of protein:crRNA:trRNA is 1:2:2 for ctRNPCas9n, whereas protein:crRNA is at a 1:2 ratio for cRNPCpf1. 3. Accordingly, to form ctRNPCas9n complex, mix 6.1 μL of 100 μM crRNA and 6.1 μL of 100 μM tracrRNA in an  RNase-free tube. Heat at ~100 C for 2 min, and cool at RT for 10 min. Add 5 μL of 10 μg/μL of Cas9n protein, incubate  at 37 C for 10 min. 4. For cRNPCpf1 complex, add 6.25 μL of 100 μM crRNA in an  RNase-free tube, heat at ~100 C for 2 min, cool at RT for 10 min, and add 5 μL of 10 μg/μL of Cpf1 protein, incubate at  37 C for 10 min. 5. Take 100 μg of donor plasmid DNA, add 20 μL of NEBuffer 3.1, and bring the volume to 171.55 μL with DEPC-treated H2O. Add the above 17.2 μL of ctRNP Cas9n complex as well as 11.25 μL of cRNPCpf1 complex, and gently mix. Incubate at  37 C for overnight. 6. Take 20 μL of the digestion mixture, column purify using Purelink Quick Combo kit, elute in 10 μg of lssDNA). About 1 μg of lssDNA will be needed for injection and another 1 μg for QC (described in Subheading 3.4). 3.4 Validation of LssDNA Donor

The integrity (length intactness, single-stranded feature) of the lssDNA generated can be verified through size comparison (by resolving on gel electrophoresis) as well as restriction enzyme diagnostics via dsDNA-specific or ssDNA-specific (e.g., exonuclease I) digestion. To validate the sequence fidelity, complementary primers can be used for lssDNA sequencing read (see Note 8).

3.4.1 Check Length Intactness

Take ~200 ng of the acquired lssDNA for size comparison with its counterpart of source dsDNA by resolving on 0.9% agarose gel at constant 100 volts for desired time, and stain with EtBr for at least 30 min to enhance lssDNA visualization (see Note 9). The lssDNA procured through the Takara kit is compared with ~200 ng PCR product of its dsDNA source (see Fig. 4c). For validating the lssDNA procured through CRISPR-CLIP , prepare its dsDNA counterpart as a control reference by using WT Cas9 and Cpf1 to digest the lssDNA-carrying plasmid (see Subheading 3.4.2); the latter comprises 2.2 kb dsDNA template, plus 2.7 kb pUC57 backbone. Take ~200 ng lssDNA to resolve on gel in parallel to the digested control plasmid (which is 4.9 kb, about twice in length of the lssDNA, thus take ~400 ng accordingly): the plasmid digested by WT Cas9 and Cpf1 would resolve at 2.2 kb and 2.7 kb, while the 2.2-kbase lssDNA would resolve around 1.1 kb (see Fig. 6a, lane “a” vs. “c”).

3.4.2 Check Single-stranded Feature

We use dsDNA-specific RE (BamHI) to digest the lssDNA donor that carries a BamHI cut site (CRISPR-CLIP case demonstrated here). To validate BamHI digestion, additional control reference on dsDNA needs to be included for the diagnostic; the dsDNA control is generated by using WT Cas9 and Cpf1 (+/ BamHI) to digest the 4.9 kb lssDNA-carrying plasmid as described below. Make sure the RE chosen does not cut the backbone. 1. To digest the plasmid, first prepare 5 μM cRNACpf1 by adding 1 μL of 100 μM crRNACpf1 in 19 μL of Duplex buffer into an  RNase-free tube, heat at 100 C for 2 min, and cool at RT for 10 min. Take 2.6 μL out and mix with 1 μg of Cpf1 protein in  another tube, incubate at 37 C for 10 min to form cRNPCpf1 complex. 2. Take 2.6 μL of 10 μM ctRNA (from Subheading 3.2.2, step 1),  add 1 μg of WT Cas9, incubate at 37 C for 10 min to form ctRNP complex.

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Fig. 6 Validation of lssDNA procured via CRISPR-CLIP. (A) Check length intactness: Cpf1 and WT Cas9 digest entire plasmid (4.9 kb) into 2.7 kb and 2.2 kb DNA fragments (lane a) vs. acquired lssDNA of 2.2 kbases resolved around ~1.1 kb in size (lane c; red arrow). (B) Check single-stranded feature using dsDNA-specific RE (BamHI) digest: (lane a) entire plasmid digested by Cpf1 and WT Cas9; (lane b) same as in lane a, with extra BamHI which digests the 2.2 kb dsDNA template into ~1 kb and ~1.2 kb; (lane c) lssDNA without BamHI digestion; (lane d) same as in lane c, with BamHI digestion but does not cleave lssDNA despite bearing BamHI cut site (red arrow). Resolved in 0.9% agarose gel electrophoresis

3. Gently mix cRNPCpf1 and ctRNP complexes from the above, add ~400 ng of plasmid, 3 μL of NEB3.1, +/ BamHI (10–20 U), and bring the total volume to 30 μL with DEPC treated H2O, incubate at 37 C for 2 h. 4. Also take ~200 ng of lssDNA and digest +/ BamHI (10–20 U), followed by resolving all four samples on 0.9% agarose gel at constant 100 volts for desired time: the 2.2 kb lssDNA source dsDNA is fragmented into 1 kb and 1.2 kb by BamHI (see Fig. 6b, lane “a” vs. “b”), indicating BamHI efficacy, while lssDNA remains a single fragment despite carrying BamHI cut site, confirming its single-stranded feature (see Fig. 6b, lane “c” vs. “d”).

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3.4.3 Check Sequence Fidelity

1. The sense lssDNA (top strand) can be sequenced with reverse primers (see Fig. 5c), whereas antisense lssDNA (bottom strand) can be sequenced with forward primers (see Fig. 4b). 2. As a negative control, include some forward primers to read top strand or include reverse primers to read bottom strand; both should not pick up correct reading, reflecting the lssDNA purity (i.e., not mixed with dsDNA). Obtaining lssDNA with better purity will result in higher editing efficiency.

3.5 Delivery of CRISPR Reagents and LssDNA into Mouse Zygote 3.5.1 Preparation of CRISPR Reagents and LssDNA Mixture

The reaction to form ctRNP complex (crRNA+tracrRNA+Cas9 protein) needs to take place first, followed by mixing with lssDNA before delivering into mouse zygotes. The concentration for Cas9 protein is 20 ng/μL (0.13 μM); for both crRNA and tracrRNA, we routinely use two concentrations 0.1 μM and 0.05 μM; the lssDNA is 10 ng/μL per kbase in length. It is noteworthy that higher crRNA/tracrRNA (0.1 μM) tends to result in lower or null number of live pups (in C57BL/6), which may be caused by probable off-target effects, yet for some loci, the positive founder mice were produced exclusively from this group. The protocol below is to prepare ctRNP complex with 0.1 μM crRNA/tracrRNA (i.e., 0.2 μM ctRNA) concentration. 1. For each ctRNP complex, take 1.2 μL of 10 μM ctRNA mixture from Subheading 3.2.2, step 1, add 1.2 μg of Cas9 protein in a  1.5 ml tube, gently mix with a pipettor, incubate at 37 C for 10 min to form the ctRNP complex. 2. Add lssDNA by adjusting its concentration to 20 ng/μL (~2 kbase length) and bring the total volume to 60 μL with IDTE. 3. Filter to remove particles that can clog injection needle; transfer the ctRNP+lssDNA cocktail to Ultrafree-MC filter unit and centrifuge at 12,000
g for 3 min. Store the cocktail on ice before the injection.

3.5.2 Pronuclear Injection

The Easi-CRISPR reagents thus prepared are delivered into mouse zygote through pronuclear injection following standard injection protocol [20]. 1. Female mice (C57BL/6 J) of 3- to 4-week-old are housed in an animal facility for a week to acclimatize to a 14 h light cycle, then hormone primed with 5 IU of PMSG (IP injection) and 48 h later, with 5 IU of hCG. 2. Following hCG injection, each female is mated with individually caged stud male. The next morning, euthanize those females displaying copulation plug and retrieve 0.5-day zygotes from the oviduct after dissecting the swollen ampulla in FHM medium.

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3. Oocytes are isolated from the cumulus-oocyte complex via treatment with hyaluronidase solution (300 μg/mL in FHM). Treated oocytes are washed with FHM, transferred to a dish with KSOM medium, and may be temporarily kept in an  incubator (37 C, 5% CO2) to await further manipulation. 4. For pronuclear injection, our Transgenic group uses Narishige manipulation system and four-chamber Lab-Tek slides with FemtoJet microinjector. Injection needles are pulled fresh for every injection session on horizontal pipette puller. For each injection session, 30–40 zygotes are placed in FHM droplets covered liquid paraffin. Injected zygotes are transferred to a  dish with KSOM and placed in a 37 C incubator with 5% CO2. 5. One to two hours after the injection session, embryos are transferred to pseudo-pregnant female CD-1 mice prepared by mating with vasectomized males. A total of 20–25 zygotes are distributed evenly and transferred to both horns of oviduct of each surrogate. After surgery, surrogate mice are placed on the warming pad and monitored until recovery from anesthesia. 3.6 Screening and Genotyping of Founder (F0) Mice 3.6.1 PCR Screening Primers and Genotyping: Novel DNA Insertion (GENE-X)

The insertion of large exogeneous cassette, such as 2A-GFP-2AGOI, is typically validated with several pairs of screening primers, where one primer (external primer) is designed to anneal to the genomic sequence that is external to the donor cassette, while the other primer (internal primer) is to anneal to the exogenous DNA sequence inserted (see Note 10); the entire KI cassette and partial genomic sequence in context should be covered by screening primers (see Fig. 7a). The primer pairs are recommended to be tested before the actual mouse genotyping. We routinely create gDNA template in mESCs for primer testing (described in the following steps 1–3) because large numbers (~0.5
106) of mESCs can be easily and economically pooled together where a subset carrying the correct insert is more likely to be found than in finite samples of zygotes. 1. Clone the CRISPR guide into px330 or any DNA vector with strong promoter (CAG/CBh) in mESCs. 2. Transfect sgRNA-px330 (200–500 ng) into 2
105 mESCs along with 500 ng of DNA donor plasmid (source of lssDNA) using lipofectamine-2000 in suspension condition at ~10 min RT incubation or any other transfection system that delivers >50% efficiency. Culture on feeder coated 12-well plate at  37 C in a humid 5% CO2 incubator. Use mESCs derived from the same mouse strain as the zygote injection is performed.

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Fig. 7 Founder mice genotyping. (a) KI of GFP-GOI at GENE-X: PCR primer pairs annealing to genomic sequence external to the donor cassette (external primer) and to the donor cassette (internal primer). (b) Floxing exon of GENE-Y: A pair of external primers for PCR amplification of the region spanning 2.2 kb insert and genomic sequence in context. (c) (top) PCR products of six pups resolved on the gel; (bottom) the same PCR products digested with EcoRV; partial PCR product resolving into two smaller fragments (a and b) indicates downstream LoxP inserted into intron 2 in one copy of the chromosome (lane 4)

3. After 48–72 h, harvest the cells and extract gDNA using High Pure PCR Template kit or any other available gDNA extraction system of your choice. 4. Test all screening primer pairs with different PCR systems using Touch-Down PCR conditions listed in Table 1. It is crucial to include one external primer from each pair to exclude random integration of the donor template in the mice that are being genotyped.

Generation of Mouse Model via Easi-CRISPR

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5. Pick one primer pair that gives robust amplification and do the actual genotyping on the mice gDNA (QIAmp Fast DNA kit for gDNA extraction); positive controls from step 3 need to be included. 6. Use the remaining primer pairs that cover the entire region for PCR screening on those positive candidate mice from step 5, followed by Sanger sequencing. 7. The other chromosome (without the 2A-GFP-2A-GOI insertion) should be checked for indels or homozygous targeting. This information will be helpful for breeding scheme. 3.6.2 PCR Screening Primers and Genotyping: Exon-Floxed CKO (GENE-Y)

For screening founder mice (F0) that carry the floxed cassette for creating GENE-Y CKO model, it is crucial to use external screening primer pairs to amplify the entire region that contains the 2.2 kb replacement donor insert and neighboring sequences in a single PCR amplification (see Fig. 7b), instead of screening for each individual LoxP. The former screening strategy is more informative and able to identify whether two LoxP sites reside on the same chromosome. Furthermore, to facilitate a simplified and efficient screening process, it is highly recommended to include RE sites adjacent to LoxP sequences in the donor template. Specifically, upon PCR amplification using an external primer pair, the PCR fragment can be digested with XbaI and EcoRV in two different reactions, which can be easily resolved on agarose gel to verify the copy number of the LoxP insert. The digested PCR products are subject to gel extraction, followed by Sanger sequencing to verify the sequence integrity of the entire floxed allele. Once the mouse line is established (beyond F1 generation), alternative screening system(s) that is less laborious, such as qPCR-based TaqMan assay, can be devised to expedite the genotyping process. 1. Extract gDNA from mice tail using the method of your choice (we routinely use QIAmp Fast DNA kit). 2. Use the primer pairs and PCR system that are empirically tested to give the most robust outcome. 3. Resolve the PCR product on 0.9% agarose gel electrophoresis to validate the amplified DNA fragment. Take 5–8 μL of the PCR product, add RE (XbaI for upstream LoxP; EcoRV for downstream LoxP), 3 μL of NEBuffer, and bring the volume to  30 μL with dH2O. Incubate at 37 C for 1 h. 4. Resolve the digested PCR product on 0.9% agarose gel. Appearance of smaller DNA fragments of predicted size on the gel indicates the insertion of the LoxP site (see Fig. 7c). The two smaller fragments can be gel purified and sequenced to determine the sequence integrity of each LoxP insert as well as the exon flanked.

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5. Alternative to step 4, if RE sites were not included in the donor, the PCR product can be TOPO cloned (in the case of heterozygous/hemizygous targeted), followed by Sanger sequencing to validate the integrity of the LoxP sequence in context. However, it may not be an efficient workflow especially when a large number of mice needs to be genotyped. 3.6.3 Results and Breeding Scheme

1. Across all the mouse models we have generated via EasiCRISPR so far, we observed that the insertion efficiency (at target site) of lssDNA donor tends to be higher for novel genes KI models than for CKO models although more data are warranted. Upon genotyping result analysis, certain mutations (e.g., nucleotide change and indels) were found in the repaired sequence as reported [21] (see Note 11) and thus those mice were excluded, which concludes editing efficiencies ranging from 8% to 23%. Given that the sequence fidelity of the lssDNA templates have been comprehensively validated before delivering into mouse zygotes, we presume these mutations might have taken place during the recombination process. 2. We have used internal primers to perform PCR-screening on those negative F0 mice and observed random (off-target) integration of exogenous DNA in all Easi-CRISPR projects. Of note, the use of primer pairs aimed for shorter segments amplication would increase the likelihood of randon insertion being detected. Nonetheless, random integration is not an exclusive issue for Easi-CRISPR, and such issue can be offset by several cycles of back-crossing or simply including wild-type mice from the F1 generation in the control group for experimental design setup, which could also be adopted to address potential CRISPR off-target effects.

4

Notes 1. Guide-B alone was used in our first round of zygote pronuclear injection but only indels were created without donor insertion; therefore, guide-A was additionally included along with guideB in the second round of injection and the targeting rate increased to 10%. We have observed in two other KI projects that the efficiency of lssDNA donor insertion can be significantly enhanced (; vs. 15%; ; vs. 23%) when using two guides simultaneously for a single target site. On the other hand, for creating gene replacement (e.g., humanization, floxed allele) model where DNA break is required to be induced at two target sites flanking the genomic sequence being replaced, we observed that using single guide for each target site is sufficient for lssDNA donor integration (with 10%–20% efficiency).

Generation of Mouse Model via Easi-CRISPR

25

Based on these data, we suspect that using two guides simultaneously to cut on the genome plausibly facilitate dislocation of a genomic segment, and such scenario may be crucial for the integration of donor lssDNA to take place. 2. Excision of exon 2 leads to removal of 2 nucleotides (nts) each from its 50 and 30 ends (see Fig. 2c), allowing 1 nt from exon 1 to disrupt the triplet nature of amino acid codon on exon 3 onward hence creating FS mutation; in addition, a de novo stop codon thus created in exon 4 will truncate the translation. 3. Exceeding Cas9 protein over 1 μg per agarose well (~1.5 mm thick, ~7 mm width) could sometime result in DNA hang-up in the well; excessive protein could obstruct the agarose pores thus immobilizing the DNA, which can be prevented by column purifying the DNA. 4. Storing the morula/blastocyst QE lysed crude gDNA at  20 C overnight results in more robust PCR amplification. 5. Herculase II Fusion system was adopted here, and a single band was generated. As long as the PCR-generated DNA band of interest constitutes the major population, other faint nonspecific bands are still tolerable. Gel purifying the dsDNA band of interest prior to Strandase reaction could be an option but would result in low yield of final product (lssDNA). 6. Zygote injection requires very little lssDNA amount (10 ng/μ L per kbase in length). The starting input of 10 robust PCR amplifications should yield >6 μg in the first eluate and >2 μg in the second eluate, which are sufficient for injection and QC. 7. Since ssDNA stains (EtBr) poorly compared to dsDNA, the DNA agarose gel is recommended to be stained with EtBr (>0.5 μg/mL) for at least 30 min after the electrophoresis (even pre-stained gel). The three ssDNA units will separate into three major bands; the 2.2 kbases of interest should run around 1.1 kb of dsDNA ladder (see Fig. 5d). 8. Although Sanger sequencing are typically performed on plasmid or PCR amplified DNA, during the sequencing reaction the dsDNA are denatured, subjecting only one strand to be read by its complementary primer. 9. The lssDNA may not resolve at exact predicted size (1.1 kb in this case), depending on certain factors, such as buffer condition and potential secondary structure formation due to sequence composition, in which case additional DGLB treatment on the acquired lssDNA is needed for size separation with more precise outcome. The integrity of the procured lssDNA is considered of good quality as long as the majority of the lssDNA resolved close to the predicted size.

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10. Since PCR preferentially amplifies the shorter DNA fragment (WT without insert), using a pair of only external primers tend not to robustly amplify the entire 1.5 kb insert sequence (2A-GFP-2A-GOI). 11. The lssDNA used in Codner GF et al. [21] was generated via ivTRT method, which lacks a proof reading feature hence could introduce mutations in the lssDNA donor. The mutations found in repaired sequence could also be attributable to the lssDNA supplied, if the sequence fidelity was not verified (e.g., by using complementary primer for Sanger sequencing as described in Subheading 3.4) before zygote injection.

Acknowledgments We thank our lab (CGERC) members Vhy-Shelta Kewaldar, Pradip Kar, and Jing Gao for their help and assistance with the experiments; Transgenic (TRTC) members Jahnney Torres, Roxana Cubias, William Ramirez, and Eunyoung Kim for zygote microinjection and mouse husbandry; Patrick Smith and David Knorr (LMGI) for collaborative efforts in trying out the Easi-CRISPR system while we were at early stage establishing relevant methods and protocols. We gratefully acknowledge the support and help of Ravi Tolwani, Associate Vice President, Comparative Bioscience Center of the Rockefeller University, for providing resources to make this manuscript possible. We also give our thanks to Betty Shih, Enlightagen Consulting for the editorial contribution and helpful discussions on the manuscript. References 1. Cong L, Ran FA, Cox D et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339:819–823 2. Mali P, Yang L, Esvelt KM et al (2013) RNA-guided human genome engineering via Cas9. Science 339:823–826 3. Suzuki K, Tsunekawa Y, Hernandez-Benitez R et al (2016) In vivo genome editing via CRISPR/Cas9 mediated homologyindependent targeted integration. Nature 540:144–149 4. Yang H, Wang H, Shivalila CS et al (2013) One-step generation of mice carrying reporter and conditional alleles by CRISPR/Casmediated genome engineering. Cell 154:1370–1379 5. Gu B, Posfai E, Rossant J (2018) Efficient generation of targeted large insertions by

microinjection into two-cell-stage mouse embryos. Nat Biotechnol 36:632–637 6. Quadros RM, Miura H, Harms DW et al (2017) Easi-CRISPR: a robust method for one-step generation of mice carrying conditional and insertion alleles using long ssDNA donors and CRISPR ribonucleoproteins. Genome Biol 18:92 7. Roth TL, Puig-Saus C, Yu R et al (2018) Reprogramming human T cell function and specificity with non-viral genome targeting. Nature 559:405–409 8. Yoshimi K, Kunihiro Y, Kaneko T et al (2016) ssODN-mediated knock-in with CRISPR-Cas for large genomic regions in zygotes. Nat Commun 7:10431 9. Miura H, Quadros RM, Gurumurthy CB et al (2018) Easi-CRISPR for creating knock-in and

Generation of Mouse Model via Easi-CRISPR conditional knockout mouse models using long ssDNA donors. Nat Protoc 13:195–215 10. Shola DTN, Yang C, Kewaldar V-S et al (2020) New additions to the CRISPR toolbox: CRISPR-CLONInG and CRISPR-CLIP for donor construction in genome editing. The CRISPR J 3(2):109–122 11. Gurumurthy CB, O’brien AR, Quadros RM et al (2019) Reproducibility of CRISPR-Cas9 methods for generation of conditional mouse alleles: a multi-center evaluation. Genome Biol 20:171 12. Terada R, Johzuka-Hisatomi Y, Saitoh M et al (2007) Gene targeting by homologous recombination as a biotechnological tool for rice functional genomics. Plant Physiol 144:846–856 13. Aida T, Chiyo K, Usami T et al (2015) Cloning-free CRISPR/Cas system facilitates functional cassette knock-in in mice. Genome Biol 16:87 14. Mashiko D, Young SA, Muto M et al (2014) Feasibility for a large scale mouse mutagenesis by injecting CRISPR/Cas plasmid into zygotes. Develop Growth Differ 56:122–129 15. Chen S, Lee B, Lee AY et al (2016) Highly efficient mouse genome editing by CRISPR

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ribonucleoprotein electroporation of zygotes. J Biol Chem 291:14457–14467 16. Bell CC, Magor GW, Gillinder KR et al (2014) A high-throughput screening strategy for detecting CRISPR-Cas9 induced mutations using next-generation sequencing. BMC Genomics 15:1002 17. Sentmanat MF, Peters ST, Florian CP et al (2018) A survey of validation strategies for CRISPR-Cas9 editing. Sci Rep 8:888 18. Li H, Beckman KA, Pessino V et al (2017) Design and specificity of long ssDNA donors for CRISPR-based knock-in. bioRxiv:178905 19. Richardson CD, Ray GJ, Dewitt MA et al (2016) Enhancing homology-directed genome editing by catalytically active and inactive CRISPR-Cas9 using asymmetric donor DNA. Nat Biotechnol 34:339–344 20. Behringer RGM, Nagy KV, Nagy A (2014) Manipulating the mouse embryo: a laboratory manual, 4th edn. Cold Spring Harbor Laboratory Press, New York 21. Codner GF, Mianne J, Caulder A et al (2018) Application of long single-stranded DNA donors in genome editing: generation and validation of mouse mutants. BMC Biol 16:70

Chapter 2 Analysis of Gene Expression Using lacZ Reporter Mouse Lines Michael Simon Kr€amer, Robert Feil, and Hannes Schmidt Abstract Reporter mice transgenically expressing the bacterial (E. coli) lacZ gene encoding β-galactosidase (β-gal, EC 3.2.1.23) are a versatile and extensively used tool to study gene expression and cell lineage patterns. Enzymatic activity of the β-gal reporter can be effectively visualized at cellular resolution either histochemically using 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-gal) or by immunofluorescent detection using a β-gal-specific antibody. Here, we summarize protocols for the localization of β-gal expressing cells in whole embryos or organs as well as in histological tissue sections of lacZ reporter mice and discuss their limitations and common pitfalls. Key words Gene expression, Reporter mice, lacZ, β-galactosidase, X-gal, Immunofluorescence staining

1

Introduction

1.1 The lacZ Reporter System

A key aspect to elucidate the function of a given protein is the determination of its temporal and spatial expression pattern. Mouse lines engineered to express a reporter transgene under control of a given gene promoter provide an invaluable resource to identify and mark cellular subsets defined by expression of a gene of interest, based on the detection of the reporter protein. The bacterial β-galactosidase (β-gal, EC 3.2.1.23) that is encoded by the lacZ gene of Escherichia coli (E. coli) represents one of the most popular reporters owing to its use as a sensitive histochemical marker that does not interfere with cellular function [1–3]. The β-gal protein, comprising 1024 amino acids [4, 5], is arranged into a 464-kDa homotetrameric enzyme [6, 7] that catalyzes the hydrolysis of β-galactosides into monosaccharides. This enzymatic activity has been utilized for histochemical detection of β-gal through the conversion of the soluble colorless glycoside 5-bromo-4chloro-3-indolyl-β-D-galactopyranoside (X-gal, also termed BCIG) [8] into an insoluble blue product. This assay involves two

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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consecutive steps: First, X-gal is hydrolyzed into galactose and the colorless 5-bromo-4-chloro-3-hydroxyindole. In the second step, this becomes oxidized and spontaneously dimerizes into the blue indigo precipitate 5,50 -dibromo-4,40 -dichloro-indigo [9, 10] (see Fig. 1a). The latter reaction is aided by ferric and ferrous ions that serve as electron acceptors [11]. Meanwhile, several other chromogenic substrates of β-gal are available that produce differently colored reaction products including 6-chloro-3-indolyl-β-Dgalactopyranoside (Salmon-gal), 5-bromo-6-chloro-3indolyl-β-D-galactopyranoside (Magenta-gal), and 5-bromo-3indolyl-β-D-galactopyranoside (Bluo-gal) [10, 12–15]. To substitute for potassium ferri- and ferrocyanide, tetrazolium salts like nitroblue tetrazolium (NBT), tetranitroblue tetrazolium (TNBT), and iodonitrotetrazolium (INT) have been used together with β-Dgalactopyranosides to detect β-gal activity through the generation of colored formazan precipitates [16]. Several reports indicated that the combination of X-gal with TNBT, X-gal with NBT, Salmon-gal with TNBT, or Salmon-gal with ferric and ferrous ions could increase the sensitivity and speed of β-gal detection in comparison with conventional X-Gal/FeCN staining [15, 17– 19]. However, this might also pose a higher risk for background staining due to endogeneous β-gal activity. In addition to the use of chromogenic substrates, the localization of β-gal in fixed tissues can also be analyzed by immunofluorescence staining using commercially available antibodies against β-gal. Furthermore, fluorogenic β-gal substrates have been developed that enable the sensitive identification of β-gal expressing living cells [20–22] using fluorescence microscopy or fluorescence-activated cell sorting. 1.2 lacZ Reporter Mice

The high sensitivity and relative ease of β-gal detection made lacZ reporter mice a favorite tool for the dissection of cis-acting gene regulatory elements, the analysis of gene expression patterns, and the investigation of cellular lineages. For these purposes, transgenic lacZ reporter mouse lines have been generated either by (1) random genomic integration of a transgene containing a lacZ expression cassette under control of the transcriptional regulatory elements of interest [23–25], or by (2) targeted insertion of lacZ into the gene of interest (knock-in). This gene-specific knock-in of a lacZ cassette can also be used to generate a null allele (knockout) of the respective gene. A further variant of lacZ reporter mice is obtained via (3) the genomic insertion of a conditional lacZ construct that only becomes expressed after Cre-mediated removal of a preceding floxed STOPcassette. A frequently used example for the latter is the Rosa26R line in which the conditional lacZ reporter is inserted into the broadly expressed Rosa locus [26]. Combination of the Rosa26R line with Cre driver lines with either cell type-specific or tamoxifen-inducible expression of Cre recombinase [27] confers cell type-specificity and/or temporal control of lacZ reporter expression which can be applied for genetic inducible fate mapping [28–31].

β-galactosidase Detection

31

Fig. 1 Detection of β-gal activity in lacZ reporter mice via X-gal or immunofluorescence staining. (a) The two-step mechanism of β-gal-mediated conversion of X-gal into the insoluble blue 5,50 -dibromo-

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Since the introduction of the first transgenic lacZ reporter mouse in 1987 [23], there has been a constant rise in reports on lacZ mice and many are now readily available from public mouse repositories, e.g., from the Knockout Mouse Project (KOMP) which aims to produce a lacZ-tagged null mutation for every protein-coding gene in the mouse [32] (see Note 1). If the generation of a new lacZ reporter mouse model is required, the appropriate type of lacZ expression cassette should be adapted to the desired applications. The use of a lacZ construct that lacks a subcellular localization sequence (see Table 1) results in a cytosolic distribution of the β-gal reporter [23, 33]. Thus, detection of β-gal by histochemical or antibody staining will mark the whole cellular extent of positive cells. However, for the unambiguous identification of individual β-gal expressing cells, it might be helpful to use a lacZ expression cassette that is targeted to a specific subcellular compartment. A tau-lacZ cassette which encodes the fusion of the microtubule-associated protein tau with β-gal [34] (see Table 1) has been instrumental in the visualization of axonal projections [35]. Widely used are also lacZ constructs with a nuclear localization sequence (NLS; see Table 1) that confine the localization of β-gal to the nuclei of positive cells (see Fig. 1b–g; see Table 1) and thereby allow the easy identification of individual positive cells [36]. In addition, the R26NZG line enables the Cre-dependent expression of nuclear targeted β-gal under control of the Rosa26 locus [37] (see Table 1). In this chapter, we provide protocols that describe in detail the assays for histochemical or immunological detection of β-gal expression in whole-mount embryos, organs, and tissue sections of lacZ reporter mice. Furthermore, we discuss the limitations and common pitfalls of these techniques. ä Fig. 1 (continued) 4,40 -dichloro-indigo precipitate. (b–g) Exemplary results are shown for lacZ reporter mouse lines for C-type natriuretic peptide (CNP) and natriuretic peptide receptor 2 (Npr2)–a ligand/receptor-pair that controls bifurcation of afferent axons in dorsal root ganglion (DRG) neurons during development [43]. (b, e) X-gal staining of whole-mount embryos at E11.5 reveals expression of CNP in the dorsal spinal cord and in the hindbrain, whereas Npr2 expression is detected in DRGs, cranial sensory ganglia, the hind brain, and the mesencephalon. (c, f) X-gal staining of transversal tissue sections of the embryonic spinal cord reveals a complementary localization of CNP and Npr2 in the dorsal spinal cord and DRGs, respectively. (d, g) The latter observation is confirmed by immunofluorescence staining of cryosections using an antibody specific for β-gal (see Subheading 2.3.1, item 5). Note that due to the presence of a nuclear localization sequence in the lacZ expression cassette, β-gal activity is confined to the nuclei of CNP or Npr2 expressing cells. For better orientation, the margins of the spinal cord and the DRGs are indicated by broken lines in c, d, f, and g. While X-gal staining of whole-mount tissue provides a first impression of the overall expression pattern of the gene of interest, the staining of sections enables a more detailed observation of its cellular expression profile, which might be further characterized by double immunostaining using a β-gal-specific antibody in combination with an antibody for a cell type-specific marker (modified from [44]). Scale bars, 1 mm in b and e, 100 μm in c, d, f, and g

β-galactosidase Detection

2

33

Materials

2.1 Dissection and Fixation of Specimen

1. Dissecting instruments (scissors, large forceps, Vannas-style spring scissors, Dumont No. 5 dissecting forceps, and a spoon for tissue transfer). 2. Dissecting stereomicroscope. 3. Peristaltic pump for perfusion fixation. 4. Dissection pad and sticky tape. 5. 12-well plates and 10-cm petri dishes. 6. 4 mL glass vials. 7. Horizontal shaker, roller shaker. 8. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 9.58 mM Na2HPO4, 1.47 mM KH2PO4, pH 7.4. 9. Zamboni’s fixative solution for subsequent X-gal staining (modified from [38]): 2% (w/v) paraformaldehyde (PFA), 0.15% (v/v) picric acid in PBS, pH 7.4. 10. Methanol. 11. Dent’s fixative: 80% methanol, 20% dimethyl sulfoxide. 12. Needles (27-gauge).

2.2 Histochemical Detection of β-Galactosidase Activity 2.2.1 X-Gal Staining of Whole Embryos or Isolated Organs

1. X-gal stock solution: 20 mg/mL X-gal (5-bromo-4-chloro-3indolyl-β-D-galactoside) in dimethylformamide (see Note 2), protect from light and store at 20  C. 2. X-gal equilibration solution: 2 mM MgCl2, 0.01% (w/v) sodium deoxycholate, 0.02% (v/v) nonidet-P40 substitute in PBS. 3. X-gal staining solution: 2 mM MgCl2, 5 mM K3[Fe(CN)6], 5 mM K4[Fe(CN)6], 0.01% (w/v) sodium deoxycholate, and 0.02% (v/v) nonidet-P40 substitute in PBS, store at 4  C. Before use, freshly add X-gal to the pre-warmed staining solution to a final concentration of 0.5 mg/mL. Protect from light. 4. X-gal wash buffer: 2 mM MgCl2 in PBS. 5. PFA fixative: 4% PFA in PBS, pH 7.4.

2.2.2 Clearing of X-Gal Stained Embryos/Organs

1. ScaleA2 clearing solution [39]: 4 M urea, 10% glycerol, and 0.1% Triton X-100.

2.2.3 X-Gal Staining of Frozen Sections

1. PBS (see Subheading 2.1, item 8). 2. Cryopreservation solution: 30% (w/v) sucrose in PBS, store at 4  C. 3. Tissue-Tek (O.C.T. Compound from Sakura Finetek). 4. Silicone embedding molds.

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Table 1 lacZ reporter gene constructs targeted to different subcellular compartments

Construct

Targeted compartment Advantage

Selected references

lacZ

Cytosol

Visualization of cellular morphology

Differentiation of neighboring individual cells may be complicated

[23, 33]

Tau-lacZ

Axon

Visualization of axonal projection patterns

Differentiation of neighboring individual cells may be complicated

[34, 35]

NLS-lacZ

Nucleus

Easy differentiation of individual cells

No information about cellular morphology

[36]

R26-LacZ (R26R)

Cytosol

Enables tracing of cell lineages

Differentiation of neighboring individual cells may be complicated

[26]

R26-NLS- Nucleus LacZEGFP (R26 NZG)

Enables tracing of cell lineages

No information about cellular morphology

[37]

Disadvantage

5. Cryostat. 6. Superfrost Plus slides. 7. X-gal equilibration solution (see Subheading 2.2.1, item 2). 8. X-gal staining solution (see Subheading 2.2.1, item 3). 9. Parafilm. 10. X-gal wash buffer: (see Subheading 2.2.1, item 4). 11. Nuclear Fast Red. 12. Aqueous mounting medium, e.g., ImmuMount (Thermo Scientific). 13. Transparent nail polish. 14. Standard light microscope attached to a color camera. 2.3 Immunofluorescent Detection of β-Galactosidase 2.3.1 Immunostaining of Whole Embryos or Isolated Tissues

1. PBS (see Subheading 2.1, item 8). 2. 4 mL glass vials and 50 mL Falcon tubes. 3. Blocking buffer: 1% Normal Donkey Serum and 0.1% Triton X-100 in PBS. 4. Washing buffer: 0.1% Triton X-100 in PBS. 5. Chicken-anti-β-galactosidase ab9361, 1 mg/mL).

primary

antibody

(Abcam,

6. Donkey-anti-chicken-AlexaFluor488 secondary antibody (Jackson ImmunoResearch Laboratories, Inc., 703-545-155, 1.5 mg/mL).

β-galactosidase Detection

35

7. DAPI (4,6-Diamidine-2-phenylindole): stock solution of 0.5 mg/mL DAPI in autoclaved distilled water, store aliquots at 20  C. 8. Aqueous mounting medium, e.g., ImmuMount (Thermo Scientific). 9. Storage solution: 80% glycerol in PBS. 2.3.2 Clearing of Immunofluorescently Labeled Whole Embryos/ Organs

1. 4% PFA in PBS. 2. PBS (see Subheading 2.1, item 8). 3. Methanol. 4. Clearing medium: 1 in 2 mixture of benzyl alcohol and benzyl benzoate (BABB).

2.3.3 Immunostaining of Frozen Sections

1. Cryopreservation solution: 30% (w/v) sucrose in 1 PBS. 2. Silicon embedding molds. 3. Tissue-Tek (O.C.T. SA62550).

Compound

from

Sakura

Finetek,

4. Cryostat. 5. PAP-Pen (Science Services GmbH, Germany, N71310). 6. PBS (see Subheading 2.1, item 8). 7. Superfrost Plus slides. 8. Humid staining chamber with horizontal slide holders. 9. Blocking buffer (see Subheading 2.3.1, item 3). 10. Washing buffer (see Subheading 2.3.1, item 4). 11. Chicken-anti-β-galactosidase ab9361, 1 mg/mL).

primary

antibody

(Abcam,

12. Donkey-anti-chicken-AlexaFluor488 secondary antibody (Jackson ImmunoResearch Laboratories, Inc., 703-545-155, 1.5 mg/mL). 13. DAPI (4,6-Diamidine-2-phenylindole): stock solution of 0.5 mg/mL DAPI in autoclaved distilled water, store aliquots at 20  C. 14. Aqueous mounting medium, e.g., ImmuMount (Thermo Scientific). 2.4 Identification of β-Galactosidase Expressing Live Cells Using a Fluorogenic Substrate

1. PBS (see Subheading 2.1, item 8). 2. Dimethyl sulfoxide (DMSO). 3. Fluorescein di-β-D-galactopyranoside (FDG; Thermo Fisher, F1179): stock solution of 10 mM FDG in DMSO, store at 20  C in the dark.

36

3 3.1

€mer et al. Michael Simon Kra

Methods Breeding of Mice

1. Set up breeding to generate experimental mice with either heterozygous and/or homozygous expression of the lacZ reporter allele (see Notes 1 and 3). If lacZ reporter expression resulting from Cre-mediated recombination should be analyzed, devise a mating strategy combining parental animals from a conditional lacZ reporter strain and an appropriate Cre-driver line. From the resulting litters identify animals with the appropriate genotype by PCR analysis of genomic DNA acquired from ear biopsies. 2. For analysis of gene expression at embryonic stages, set up timed matings and check for vaginal plugs every 12 h. The time point of plug detection is determined as embryonic day (E) 0.5. In order to minimize the numbers of experimental animals, plugged mice should be weighed after plug detection to enable the determination of pregnancy at later stages through monitoring of weight gain (see Note 4).

3.2 Dissection and Fixation of Specimen 3.2.1 Immersion-Fixation

1. Sacrifice the pregnant dam or experimental mouse at the desired age using an approved method of euthanasia. 2. Collect embryos by cesarean section from the uterine horns of pregnant dams or dissect other tissues of interest. Transfer the embryos or tissues for trimming to a 10 cm petri dish with ice-cold PBS. 3. Under microscopic control, free the embryos from their embryonic sheaths and decapitate them, trim other tissues, and subsequently transfer them to individual wells of a 12-well plate each filled with 2.5 mL ice-cold PBS or to a 4 mL glass vial containing 3 mL ice-cold PBS (see Note 5). Keep the yolk sac or a part of the head of each embryo for genotyping. 4. Replace the PBS in the wells of the 12-well plate or in the 4 mL glass vial with 2.5 or 3 mL ice-cold Zamboni’s fixative solution, respectively, and agitate for 10–120 min at 4  C on a horizontal or a roller shaker (see Notes 6 and 7).

3.2.2 Fixation of Whole Animals by Perfusion

To secure a rapid and uniform fixation especially of larger specimen, whole lacZ reporter mice might be perfused transcardially with fixative solution instead of immersion fixation of tissues. 1. In a fume hood, set up a peristaltic pump system for perfusion and connect the outlet port of the tubing with a 27-gauge needle. Flush the tubing with perfusion buffer until the tubing is free of any bubbles.

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2. Sacrifice the experimental mouse using an approved method of euthanasia and mount the animal postmortem with sticky tape to a dissection pad. Place the pad into a tray to enable the collection of used fixative solution for proper disposal. 3. Wet the animal’s fur with 70% ethanol and open the abdominal wall beneath the rib cage. Cut through the diaphragm and lift the chest plate after two lateral incisions through the ribs to the collarbone. Take care not to damage either liver or lungs. 4. After a small incision into the right atrium of the heart to create an outlet, puncture the left ventricle to insert the perfusion needle towards the ascending aorta and perfuse the animal for 3–4 min with PBS (see Note 8). 5. Subsequently, switch perfusion to ice-cold Zamboni’s fixative solution for further 5 min (see Note 9). 6. Isolate the organs of interest from the perfused mouse and briefly trim the tissue in ice-cold PBS. 7. Post-fix tissues in a well of a 12-well plate filled with 2.5 mL of Zamboni’s fixative solution for 10–120 min on ice with gentle shaking (see Notes 6 and 7). 3.2.3 Fixation for Immunostaining of Whole-Mount Embryos or Isolated Tissue

1. Following immersion-fixation (see Subheading 3.2.1) or fixation by perfusion (see Subheading 3.2.2), wash the embryo or tissue twice for 30 min in ice-cold PBS. 2. Transfer the embryo or tissue into a 4 mL glass vial and dehydrate the tissue in an ascending methanol series (25%, 50%, and 75% methanol) at 4  C with 20 min incubation per step. 3. Incubate the specimen overnight at 4  C in Dent’s fixative on a roller shaker. After this step, wash once with 80% methanol/ PBS. At this point, the tissue can be stored for longer times in 100% methanol at 20  C.

3.3 Histochemical Detection of β-Galactosidase Activity

1. Following fixation (see Subheadings 3.2.1 or 3.2.2), collect the fixation solution for proper disposal and equilibrate the samples in X-gal equilibration solution three times for 30 min at ambient temperature with gentle agitation.

3.3.1 X-Gal Staining of Whole-Mount Mouse Embryos or Isolated Organs

2. Incubate whole embryos or organs in X-gal staining solution for up to 24 h at 37  C under constant agitation in the dark (see Notes 10–13). 3. After development of the blue X-gal precipitate, stop the reaction by washing three times with ice-cold PBS containing 2 mM MgCl2 for 10 min each. 4. Post-fix tissue in 4% paraformaldehyde solution at 4 overnight.



C

5. Wash tissue three times for 30 min in PBS at room temperature.

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3.3.2 Clearing of X-Gal Stained Whole-Mount Embryos/Organs for Documentation

1. Discard the PBS and incubate the samples in ScaleA2 clearing solution for 1 week at room temperature under light agitation to increase transparency of the tissue.

3.3.3 X-Gal Staining of Frozen Sections

1. Following fixation of the specimen (see Subheadings 3.2.1 or 3.2.2), collect the fixative solution for proper disposal and wash the tissue three times for 30 min each in ice-cold PBS to remove any traces of the fixative.

2. After sufficient clearing, analyze X-gal stained whole-mount tissues in a petri dish completely covered with ScaleA2 using a dissecting microscope attached to a color camera. For best results, the specimen should be imaged before a white background using indirect illumination.

2. Cryo-protect the tissue by overnight incubation in 30% sucrose in PBS at 4  C. 3. Remove excess liquid and place individual samples into appropriately sized embedding molds filled with OCT compound. Eliminate any air bubbles using a micropipette and place the molds onto dry ice to freeze the specimen. After freezing, store the appropriately labeled specimen in air-tight containers at 80  C until sectioning. 4. Prepare 25 μm cryosections and mount them onto Superfrost plus slides. Air-dry the slides with the sections for 30 min at room temperature. The slides can then be stored in air-tight slide boxes at 80  C before further use. 5. Equilibrate the slides at room temperature and incubate the sections three times for 10 min with approx. 300 μL X-gal equilibration solution at room temperature (see Note 14). 6. Aspirate X-gal equilibration solution and pipette approx. 200 μL X-gal staining solution onto the slide. Cover the liquid with a cut-to-fit strip of parafilm. Turn the slide upside down and place it onto horizontal slide holders in a humid chamber. Make sure that no liquid bridge forms between the parafilmcovered area and the lateral slide holders to avoid leaking out of the X-gal staining solution. Incubate for 2–24 h at 37  C in the dark until a blue precipitate becomes visible (see Notes 10, 11, 13, and 15). 7. Invert the slide and remove the parafilm cover slip using forceps. Aspirate remaining X-gal staining solution and stop the reaction by 3 washes with cold PBS with 2 mM MgCl2 for 10 min each. 8. Wash the slide(s) once with tap water. 9. Counterstain the sections in Nuclear Fast Red solution for 3–10 min at room temperature (see Note 16). 10. Aspirate excess Nuclear Fast Red solution and dip the slide twice into a cuvette filled with PBS to wash.

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11. Mount the sections using an aqueous mounting medium, e.g., ImmuMount, and seal the coverslips with nail polish the following day. 12. For photo documentation, image the X-gal stained sections using a standard light microscope attached to a color camera (see Note 17). 3.4 Immunofluorescent Detection of β-Galactosidase 3.4.1 Immunostaining of Whole-Mount Mouse Embryos or Isolated Tissue

Confocal microscopy of immunostained whole-mount embryos or tissues provides a good overview of the localization of β-gal within the specimen. Best results are obtained with embryos up to E12.5. Larger embryos or organs should be dissected into smaller pieces for staining. 1. Rehydrate the fixed embryo or tissue (see Subheading 3.2.3) in graded methanol (80%, 50%, and 25% methanol, PBS) at 4  C with 20 min incubation per step. 2. Incubate the specimen in 2 mL of blocking buffer overnight at 4  C under constant agitation to block nonspecific antigens. 3. Remove the blocking buffer and incubate the tissue with the chicken-anti-β-galactosidase primary antibody (diluted at 1 in 2000 in blocking buffer) for 2 days at 4  C. 4. Transfer the samples to a 50 mL Falcon tube filled with 45 mL washing buffer and incubate for 1 h under gentle agitation. Repeat this step 6 times. 5. Transfer the samples to a 4 mL glass vial and incubate tissue in the donkey-anti-chicken-AlexaFluor488 secondary antibody (diluted at 1 in 1000 in blocking buffer) for 2 days under gentle agitation. For nuclear counter staining, include DAPI diluted at 1 in 1000 in blocking buffer. 6. Transfer the samples to a 50 mL Falcon tube filled with 45 mL washing buffer and incubate for 1 h under gentle agitation. Repeat this step 6 times. 7. Add few drops of ImmuMount to a slide and mount the sample with a coverslip. Prior to mounting, remove excess liquid from the sample. Seal the coverslip the following day with nail polish for long-term storage. 8. Alternatively, larger tissues can be stored in 80% glycerol in PBS for partial clearing or processes for clearing with BABB (see Subheading 3.4.2).

3.4.2 Clearing of Immunofluorescent Stained Whole Embryos/ Tissues for Documentation

1. Transfer the samples to a 4 mL glass vial and post-fix for 4 h at 4  C with 3 mL 4% PFA in PBS. 2. Wash three times for 30 min each with 3 mL PBS.

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3. Dehydrate the tissue in an ascending methanol series (25%, 50%, 80%, and 100% methanol) with 20 min incubation per step. 4. For rapid optical clearing, transfer the tissue into a glass vial with BABB. Incubate the tissue for 3–48 h, change the solution 1–2 times until the samples become transparent (see Note 18). 5. For photo documentation, place the specimen into an appropriately sized dish and record Z-stacks using a confocal microscope (see Note 19). 3.4.3 Immunostaining of Cryosections

1. Following fixation of the specimen (see Subheadings 3.2.1 or 3.2.2), collect the fixative for proper disposal and wash the tissue three times for 30 min in ice-cold PBS. 2. Incubate tissue in cryopreservation solution at 4  C overnight. 3. Embed tissue into appropriately sized silicone molds filled with Tissue-Tek. 4. Prepare frozen sections of a suitable thickness and mount them onto Superfrost Plus slides. Air-dry the slides for 30 min at room temperature. The slides can then be stored in air-tight slide boxes at 80  C before further processing. 5. Equilibrate the slide(s) at room temperature and encircle the sections with hydrophobic barrier using a PAP-Pen. 6. Carefully pipette 300 μL PBS onto the slide and incubate for 5 min to remove Tissue-Tek. 7. Aspirate the PBS and incubate the sections in blocking buffer for 1 h at room temperature to block nonspecific antigens. 8. Incubate sections in the primary chicken-anti-β-galactosidase antibody (diluted at 1 in 2000 in blocking buffer) overnight at 4  C. 9. Wash the slide(s) three times for 10 min at room temperature in a staining cuvette with washing buffer. 10. Incubate sections in the secondary donkey-anti-chicken-AlexaFluor488 antibody (diluted at 1 in 1000 in blocking buffer) for 2 h at room temperature in the dark. 11. Thereafter, wash the slide(s) under slight agitation in a glass cuvette filled with washing buffer for 10 min at room temperature. Repeat twice. 12. Wash sections once for 10 min in PBS. 13. Remove excess liquid, add few drops of aqueous mounting medium and mount the sections with a coverslip. 14. On the following day, seal the coverslips with nail polish and store the slides at 4  C in the dark. 15. Image the samples using fluorescence microscopy.

β-galactosidase Detection

3.5 Identification of β-Galactosidase Expression in Living Cells

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Several fluorogenic substrates are available for detection of β-gal activity in vivo and in vitro, e.g., fluorescein di-β-D-galactoside (FDG) – a very sensitive fluorogenic substrate that upon hydrolysis by β-gal releases strongly fluorescent fluorescein. 1. Dilute the 10 mM stock solution of FDG in the respective cell culture medium at 1 in 1000 to produce a 10 μM working solution (final DMSO concentration: 0,1%). 2. Incubate cells with FDG working solution for 10 min at 37  C in the incubator. 3. Wash cells with pre-warmed culture medium for 1 min. Repeat twice. 4. Incubate cells in culture medium for further 20 min at 37  C in the incubator. 5. Subsequently, lacZ-positive cells can be detected by fluorescence microscopy or flow cytometry using an excitation/emission setting of 490 nm and 514 nm, respectively.

4

Notes 1. LacZ reporter mouse lines are available for many genes from repositories like the European Mouse Mutant Archive (EMMA, https://www.infrafrontier.eu/search), the International Mouse Strain Resource (IMSR, http://www.findmice. org/), the Jackson Laboratory (https://www.jax.org/jaxmice-and-services), the Knockout Mouse Project (KOMP, https://www.komp.org/), or the Mutant Mouse Resource & Research Centers (MMRRC, https://www.mmrrc.org/). 2. Special care should be taken when handling dimethylformamide since it is a potent liver toxin which is readily absorbed through the skin. The X-gal stock solution in dimethylformamide should be prepared in either glass or polypropylene vials. Aliquots should be kept at 20  C in the dark. 3. Apart from studying the expression of a certain gene of interest, the knock-in of a lacZ cassette into a gene locus can also be used to generate a knockout of this specific gene. Cross-breeding of heterozygous mice will generate litters with wild-type, heterozygous, and homozygous (knockout) offspring. The phenotypical comparison of littermates with different genotypes, e.g., the analysis of the localization of β-gal-positive cells, might provide a clue about the function of the protein encoded by the gene of interest. If homozygous lacZ reporter mice are viable and fertile, they can be mated with wild-type animals to generate litters in which all individuals will be heterozygous for the lacZ reporter. Furthermore, it should be noted that due to

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the presence of two lacZ alleles in homozygous animals the intensity of X-gal staining might be somewhat stronger than that observed in β-gal-positive cells of heterozygous animals. 4. Weight gains of 2 g or more between plug detection at E0.5 and E8.5 indicate a developing pregnancy. 5. If separation of individual samples with different genotypes is not an issue, all tissues might be collected into a single 15 mL Falcon tube for further processing. 6. The time for optimal fixation depends on tissue size, the level of β-gal activity, and further use (β-gal detection in whole-mount tissues or sections) and should be individually determined. The fixation for subsequent preparation of cryostat sections requires longer times than for the detection of β-gal activity in whole embryos or organs. However, prolonged fixation might impair the enzymatic activity of β-gal leading to falsenegative results. 7. Apart from Zamboni’s fixative, other fixative solutions based on paraformaldehyde or glutaraldehyde might be used for subsequent detection of β-gal. However, fixation using glutaraldehyde causes an increase in background fluorescence of the tissue that impedes immunofluorescence analysis. 8. Gradual clearing of the liver from remaining blood gives a good indication for the quality of the perfusion. 9. The onset of fixation is accompanied by little tremors observed especially at whisker hairs and the tail. 10. The time for the development of X-gal staining varies depending on the strength of the individual gene promotor activity and must be empirically optimized for each lacZ reporter line. Therefore, the first X-gal staining of a tissue from a previously not yet analyzed lacZ reporter mouse should be closely monitored at regular intervals. 11. The interpretation of the results of X-gal staining, especially with longer staining times, can be confounded by the presence of endogenous β-gal activity. While some background staining might be the consequence of the presence of lacZ-containing bacteria, e.g., in the lumen of the gastrointestinal tract, other background staining might result from endogenous non-bacterial β-galactosidases in a number of murine tissues, e.g., in neonatal bone, mandibular and lacrimal glands, small intestine, kidney, ovary, testis, and choroid plexus. To minimize background activity, X-gal staining should be performed at a pH of 7.4 which is the optimal pH for bacterial β-gal, whereas the endogenous mammalian β-galactosidases have their optimum at a pH between 3.5 and 5.5 [10, 11]. For unequivocal identification of genuine lacZ reporter activity,

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the use of a lacZ cassette with a nuclear localization sequence might be helpful since the resulting X-gal staining will be confined to the nucleus. Thus, it can easily be discriminated from a cytosolic background staining. Additionally, the specificity of the observed X-gal staining should always be confirmed by a parallel control staining of corresponding tissue from a wild-type littermate. 12. Due to incomplete penetration of the staining solution, the success of X-gal staining in bigger objects might be limited to peripheral layers. In order to examine β-gal activity in embryos older than E12.5 or in larger organs, it is thus advisable to further dissect the specimen or to use tissue sections. 13. For interpretation of the results of X-gal staining in tissues from lacZ reporter mice, it should be considered that, despite the high sensitivity of the detection system, positive X-gal staining requires a certain number of gene transcripts and β-gal proteins (threshold). Therefore, if β-gal proteins are expressed at very low copy numbers, X-gal staining might be negative. Alternatively, it might be useful to switch to immunofluorescent detection of β-gal using an appropriate antibody (see Subheading 3.4) as several studies reported a more sensitive detection of β-gal by immunostaining in comparison with X-gal staining [40–42]. Furthermore, the lack of X-gal-positive cells does not necessarily imply that the protein encoded by the gene of interest (in which the lacZ cassette had been inserted) is also not expressed. For instance, a “false-negative” X-gal staining might also be caused by a shorter half-life of the β-gal protein as compared to the investigated protein. Vice versa, a longer half-life of β-gal relative to the protein of interest might result in a “false-positive” X-gal staining. 14. Do not use a PAP-Pen to encircle the sections with a hydrophobic boundary as this reacts with the X-gal staining solution and produces an unwanted precipitate. 15. To prevent drying-out of the sections, it might be necessary to freshly add X-gal staining solution from time to time and to seal the humid chamber with parafilm. 16. After longer storage, precipitates might form in the Nuclear Fast Red solution which could compromise the staining result. Therefore, the Nuclear Fast Red solution should be centrifuged for 2 min at maximum speed in a benchtop centrifuge to sediment any particles. Then, carefully remove the supernatant without disturbing the pellet. Re-centrifuge the supernatant once again before application to the sections. 17. Since the pink Nuclear Fast Red staining will gradually fade in the aqueous mounting medium, images should be acquired soon after mounting.

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18. While incubation in BABB enables rapid clearing of tissues, it will cause the samples to shrink. Tissues that have been left in BABB for too long will eventually become brittle and finally break apart. BABB should only be applied in glass vials or resistant plastic ware, e.g., Lumox dishes. 19. Since BABB can damage microscope lenses, care should be taken to avoid any leakage. Alternatively, special objectives with a resistant front lens are available for the use with BABB immersion medium (e.g., Leica HCX APO L20/0.95 IMM).

Acknowledgments We thank Dr. Maria TK Zaldivia for critically reading the manuscript. This work was supported by the DFG (FOR 2060 project SCHM 2371/1) to HS. References 1. Cui C, Wani MA, Wight D et al (1994) Reporter genes in transgenic mice. Transgenic Res 3:182–194 2. Takahashi E, Miyamoto N, Kajiwara N et al (2000) Expression analysis of Escherichia coli lacZ reporter gene in transgenic mice. Brain Res Brain Res Protoc 5:159–166 3. Burn SF (2012) Detection of betagalactosidase activity: X-gal staining. Methods Mol Biol 886:241–250 4. Fowler AV, Zabin I (1978) Amino acid sequence of beta-galactosidase. XI Peptide ordering procedures and the complete sequence. J Biol Chem 253:5521–5525 5. Kalnins A, Otto K, Ruther U et al (1983) Sequence of the lacZ gene of Escherichia coli. EMBO J 2:593–597 6. Jacobson RH, Zhang XJ, DuBose RF et al (1994) Three-dimensional structure of betagalactosidase from E. coli. Nature 369:761–766 7. Juers DH, Matthews BW, Huber RE (2012) LacZ beta-galactosidase: structure and function of an enzyme of historical and molecular biological importance. Protein Sci 21:1792–1807 8. Horwitz JP, Chua J, Curby RJ et al (1964) Substrates for cytochemical demonstration of enzyme activity. I. Some substituted 3-indolylbeta-D-glycopyranosides. J Med Chem 7:574–575 9. Cotson S, Holt SJ (1958) Studies in enzyme cytochemistry. IV. Kinetics of aerial oxidation of indoxyl and some of its halogen derivatives. Proc R Soc Lond B Biol Sci 148:506–519

10. Pearson B, Wolf PL, Vazquez J (1963) A comparative study of a series of new indolyl compounds to localize beta-galactosidase in tissues. Lab Investig 12:1249–1259 11. Lojda Z (1970) Indigogenic methods for glycosidases. II An improved method for beta-Dgalactosidase and its application to localization studies of the enzymes in the intestine and in other tissues. Histochemie 23:266–288 12. Aguzzi A, Theuring F (1994) Improved in situ beta-galactosidase staining for histological analysis of transgenic mice. Histochemistry 102:477–481 13. Brunet LJ, McMahon JA, McMahon AP et al (1998) Noggin, cartilage morphogenesis, and joint formation in the mammalian skeleton. Science 280:1455–1457 14. Schmidt A, Tief K, Foletti A et al (1998) lacZ transgenic mice to monitor gene expression in embryo and adult. Brain Res Brain Res Protoc 3:54–60 15. Kishigami S, Komatsu Y, Takeda H et al (2006) Optimized beta-galactosidase staining method for simultaneous detection of endogenous gene expression in early mouse embryos. Genesis 44:57–65 16. Altman FP (1976) Tetrazolium salts and formazans. Prog Histochem Cytochem 9:1–56 17. Gugliotta P, Pacchioni D, Bussolati G (1992) Staining reaction for beta-galactosidase in immunocytochemistry and in situ hybridization. Eur J Histochem 36:143–148 18. Sundararajan S, Wakamiya M, Behringer RR et al (2012) A fast and sensitive alternative for

β-galactosidase Detection beta-galactosidase detection in mouse embryos. Development 139:4484–4490 19. Trifonov S, Yamashita Y, Kase M et al (2016) Overview and assessment of the histochemical methods and reagents for the detection of beta-galactosidase activity in transgenic animals. Anat Sci Int 91:56–67 20. Nolan GP, Fiering S, Nicolas JF et al (1988) Fluorescence-activated cell analysis and sorting of viable mammalian cells based on beta-Dgalactosidase activity after transduction of Escherichia coli lacZ. Proc Natl Acad Sci U S A 85:2603–2607 21. Macgregor GR, Nolan GP, Fiering S et al (1991) Use of Escherichia coli (E. coli) lacZ (beta-galactosidase) as a reporter gene. Methods Mol Biol 7:217–235 22. Zhang YZ, Naleway JJ, Larison KD et al (1991) Detecting lacZ gene expression in living cells with new lipophilic, fluorogenic betagalactosidase substrates. FASEB J 5:3108–3113 23. Goring DR, Rossant J, Clapoff S et al (1987) In situ detection of beta-galactosidase in lenses of transgenic mice with a gamma-crystallin/ lacZ gene. Science 235:456–458 24. Bonnerot C, Nicolas JF (1993) Application of LacZ gene fusions to postimplantation development. Methods Enzymol 225:451–469 25. Sekerkova G, Katarova Z, Joo F et al (1997) Visualization of beta-galactosidase by enzyme and immunohistochemistry in the olfactory bulb of transgenic mice carrying the LacZ transgene. J Histochem Cytochem 45:1147–1155 26. Soriano P (1999) Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat Genet 21:70–71 27. Feil R, Wagner J, Metzger D et al (1997) Regulation of Cre recombinase activity by mutated estrogen receptor ligand-binding domains. Biochem Biophys Res Commun 237:752–757 28. Petit AC, Legue E, Nicolas JF (2005) Methods in clonal analysis and applications. Reprod Nutr Dev 45:321–339 29. Joyner AL, Zervas M (2006) Genetic inducible fate mapping in mouse: establishing genetic lineages and defining genetic neuroanatomy in the nervous system. Dev Dyn 235:2376–2385 30. Watson CM, Trainor PA, Radziewic T et al (2008) Application of lacZ transgenic mice to cell lineage studies. Methods Mol Biol 461:149–164 31. Feil S, Krauss J, Thunemann M et al (2014) Genetic inducible fate mapping in adult mice

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using tamoxifen-dependent Cre recombinases. Methods Mol Biol 1194:113–139 32. Skarnes WC, Rosen B, West AP et al (2011) A conditional knockout resource for the genome-wide study of mouse gene function. Nature 474:337–342 33. Hall CV, Jacob PE, Ringold GM et al (1983) Expression and regulation of Escherichia coli lacZ gene fusions in mammalian cells. J Mol Appl Genet 2:101–109 34. Callahan CA, Thomas JB (1994) Tau-betagalactosidase, an axon-targeted fusion protein. Proc Natl Acad Sci U S A 91:5972–5976 35. Mombaerts P, Wang F, Dulac C et al (1996) Visualizing an olfactory sensory map. Cell 87:675–686 36. Bonnerot C, Rocancourt D, Briand P et al (1987) A beta-galactosidase hybrid protein targeted to nuclei as a marker for developmental studies. Proc Natl Acad Sci U S A 84:6795–6799 37. Yamamoto M, Shook NA, Kanisicak O et al (2009) A multifunctional reporter mouse line for Cre- and FLP-dependent lineage analysis. Genesis 47:107–114 38. Stefanini M, De Martino C, Zamboni L (1967) Fixation of ejaculated spermatozoa for electron microscopy. Nature 216:173–174 39. Hama H, Kurokawa H, Kawano H et al (2011) Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat Neurosci 14:1481–1488 40. Cheng G, Thompson RP, Gourdie RG (1999) Improved detection reliability of betagalactosidase in histological preparations. BioTechniques 27:438–440 41. Mahony D, Karunaratne S, Rothnagel JA (2002) Improved detection of lacZ reporter gene expression in transgenic epithelia by immunofluorescence microscopy. Exp Dermatol 11:153–158 42. Couffinhal T, Kearney M, Sullivan A et al (1997) Histochemical staining following LacZ gene transfer underestimates transfection efficiency. Hum Gene Ther 8:929–934 43. Dumoulin A, Ter-Avetisyan G, Schmidt H et al (2018) Molecular analysis of sensory axon branching unraveled a cGMP-dependent signaling cascade. Int J Mol Sci 19:1266 44. Ter-Avetisyan G, Rathjen FG, Schmidt H (2014) Bifurcation of axons from cranial sensory neurons is disabled in the absence of Npr2-induced cGMP signaling. J Neurosci 34:737–747

Chapter 3 Linear Density Sucrose Gradients to Study Mitoribosomal Biogenesis in Tissue-Specific Knockout Mice Benedetta Ruzzenente and Metodi D. Metodiev Abstract Like bacterial and cytoplasmic ribosomes, mitoribosomes are large ribonucleoprotein complexes with molecular weights in the range of several million Daltons. Traditionally, studying the assembly of such high molecular weight complexes is done using ultracentrifugation through linear density gradients, which remains the method of choice due to its versatility and superior resolving power in the high molecular weight range. Here, we present a protocol for the analysis of mitoribosomal assembly in heart mitochondrial extracts using linear density sucrose gradients that we have previously employed to characterize the essential role of different mitochondrial proteins in mitoribosomal biogenesis. This protocol details in a stepwise manner a typical mitoribosomal assembly analysis starting with isolation of mitochondria, preparation and ultracentrifugation of the gradients, fractionation and ending with SDS-PAGE, and immunoblotting of the gradient fractions. Even though we provide an example with heart mitochondria, this protocol can be directly applied to virtually all mouse tissues, as well as cultured cells, with little to no modifications. Key words Mitoribosome, Sedimentation analysis, Sucrose gradient, mt-SSU, mt-LSU, 55S, Heartspecific knockout mice

1

Introduction Mitochondria contain a proprietary translation system that is responsible for the intramitochondrial synthesis of mtDNAencoded polypeptides, which in turn are essential for the biogenesis and function of the ATP-producing oxidative phosphorylation system in the inner mitochondrial membrane. The mitochondrial translation system includes a unique, mitochondria-specific ribosome (mitoribosome), mtDNA-encoded tRNAs, as well as nuclearencoded translation factors and mitochondrial tRNA synthetases most of which differ from those in the cytosol. Coordinated expression of the nuclear and mitochondrial genomes ensures proper mitochondrial translation and continuous supply of energy needed to sustain the functions of most eukaryotic cells.

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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The mitoribosome has been the subject of extensive scrutiny for over 50 years, which has recently peaked with the publication of high-resolution structural models for the two mitoribosomal particles and the assembled monosome [1–5]. Like its bacterial or cytosolic counterparts, the mitoribosome is composed of two multisubunit particles termed small and large or mt-SSU and mt-LSU, respectively. The two particles are ribonucleoprotein complexes composed of mtDNA-encoded 12S and 16S rRNA in mt-SSU and mt-LSU, respectively, and ~30 mt-SSU and ~50 mt-LSU proteins that are encoded by nuclear genes. In contrast to bacterial and cytosolic ribosomes, mt-LSU also contains either tRNAPhe or tRNAVal, which may fulfill a role similar to 5S rRNA [6, 7]. Mitoribosomes also feature significantly shorter rRNAs and a larger number of proteins some of which have no bacterial orthologs while others have acquired additional C- or N-terminal extensions. As a result, the two mitoribosomal particles and the monosome have different sedimentation properties when compared to their bacterial and cytosolic counterparts. Despite our significantly increased understanding of mitoribosomal structure, many aspects of its biogenesis and function remain unknown. We and our colleagues have previously studied the function of several mitochondrial proteins with important roles in mitoribosomal biogenesis and/or function. These include the mitochondrial methyltransferases TFB1M [8] and NSUN4 [9] that are essential for 12S rRNA maturation and mt-SSU biogenesis; the mt-LSU maturation factor and NSUN4 interaction partner MTERF4 [10] as well as MTERF3 whose exact mechanistic role in mitoribosomal biogenesis remains unclear [11]. These proteins are essential for mitochondrial function and their whole-body knockout results in embryonic lethality. Therefore, to study the roles of these proteins, we used a conditional Cre-loxP recombination system wherein the expression of the Cre-recombinase and consequent recombination-mediated gene inactivation are restricted to specific tissues. In our work, we used mice that express the Cre-recombinase under the control of a muscle creatine kinase promoter (ckmm) whose expression is restricted to skeletal muscle and cardiomyocytes [12]. Generation of tissue-specific knockout mice follows a standardized protocol that was described in the previous edition of this book [13] whereas we have previously published breeding schemes to obtain heart/muscle-specific knockout mice—for an example see [8]. We present here the protocol that we have developed to study mitoribosomal biogenesis— specifically, the analysis of mitoribosomal assembly using linear density sucrose gradients. This protocol is based on seminal works conducted in the 70s through 90s of the twentieth century and especially on work by Tom O’Brien, Carol C. Cunningham and their coworkers [14–16].

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Traditionally, the monosome and its particles are also described using their specific sedimentation coefficients or S values (S, Svedberg unit) which is a measure of the rate at which a molecule, macromolecular complex or cellular organelle travels under the influence of a centrifugation force depending on its size, shape, and density. The human and mouse mitoribosome, mt-LSU, and mt-SSU are thus referred to as 55S, 39S, and 28S, respectively, but these values vary between different species: mitoribosomes from Xenopus are ~60S while those from yeast and Neurospora are relatively larger at ~73–80S; bacterial and cytosolic ribosomes have sedimentation coefficients of ~70–80S and dissociate into 40S and 60S particles. In practice, this means that when a mitochondrial lysate is layered on top of a density gradient and subjected to specific g force for a given time, the mitoribosome and its two particles will sediment into different zones of the gradient which enables their specific identification, isolation, and analysis. Density gradient ultracentrifugation remains the most widely used method for analysis of mitoribosomal assembly due to its superb resolving power, near-native conditions, and versatility in regard to buffer composition, amount of starting material, and centrifugation conditions.

2

Materials

2.1 Specialized Equipment

1. A 15 mL glass/Teflon Potter-Elvehjem homogenizer attached to a motor-driven mechanical stirrer/drill—for example, Heidolph RZR2021—is used for tissue and cell disruption in the procedure “Isolation of Mitochondria.” 2. Linear density sucrose gradients are prepared using the gradient former “Gradient Master” from BioComp Instruments (Fredericton, New Brunswick, Canada). Other methods for gradient preparation exist and are described elsewhere (see Note 1). 3. Polyallomer tubes 14  89 mm from Beckman—soak in 0.1% diethyl pyrocarbonate (DEPC) in Milli-Q water for at least 12 h, drain in a bottle, and autoclave both the tubes and the DEPC solution, for 20 min at 180  C to inactivate the DEPC. The autoclaved 0.1% DEPC solution can be disposed of or used to wash the silicon tube caps supplied with the Gradient Master (see below). 4. Beckman Coulter Optima L-90 K refrigerated, floor model ultracentrifuge, and a rotor SW41Ti.

2.2 Isolation of Mitochondria (See Note 2)

1. 1 M sucrose, used for preparation of mitochondrial isolation buffer. Prepare by dissolving 342.3 g of sucrose in 500–600 mL of Milli-Q H2O, mix until all of the sucrose is

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fully dissolved and adjust the volume to 1 L with Milli-Q H2O. Because sucrose breaks down to glucose and fructose at high temperature, it should not be autoclaved. Instead, filter sterilize the stock solution using a 0.2 μm filtration unit and store at 4  C. 2. Mitochondrial Isolation Buffer (MIB): 320 mM sucrose, 10 mM Tris–HCl (pH 7.5), 1 Complete EDTA-free protease inhibitor. MIB is prepared by mixing 320 mL of 1 M sucrose (see above) with 10 mL 1 M Tris–HCl (pH 7.5) and adjusting the volume to 1 L with MiliQ H2O. Sterilize by filtration using a 0.2 μm filtration unit and store at 4  C. Prior to use, transfer the required volume of MIB to Falcon tube(s) and to each 50 mL of MIB add 1 pill of Roche’s Complete EDTA-free protease inhibitor (see Note 3). 2.3 Sucrose Gradients

Maintain RNAse-free environment during sample handling and while preparing the different buffers. Excessive RNAse activity in the samples or gradients can reduce the yield of assembled mitoribosomal particles, monosomes, and polysomes and will prevent downstream analysis of mitoribosome-mRNA association if such is intended. Most powders and solutions, as well as distilled H2O, can be purchased free of RNAse/DNase contaminations. Although Milli-Q water is essentially free of contaminating nucleases, we recommend that it is treated with 0.1% DEPC for at least 12 h while stirring followed by autoclaving for 20 min at 180  C to inactivate the DEPC. Similarly, glassware, plasticware, and the metal cannulas used for gradient preparation should be soaked in 0.1% DEPC for overnight followed by autoclavation. On the day of the experiment, clean the short (rate-zonal) plastic caps supplied with the gradient former, forceps and scissors with RNAse-ZAP, and rinse them with copious amounts of DEPC-treated water; use RNAse-free filter tips. 1. Prepare 50% w/v (~1.46 M) sucrose stock solution by dissolving 250 g sucrose in RNase-free H2O for a final volume of 500 mL. Sterilize by filtration using an RNAse-free filtration unit. This solution is used in the formulation of the lysis buffer and the light and heavy gradient solutions and can be stored either at room temperature or at 4  C. In the latter case, it will need to be equilibrated to room temperature before use (see Note 4). 2. Prepare 50 concentrated stock solution of Roche’s Complete EDTA-free protease inhibitor by dissolving 1 tablet in 1 mL of RNAse-free water. Keep on ice until needed. Remaining volumes can be stored at –20  C . 3. Prepare fresh 1.43 M dilution of β-mercaptoethanol used for the preparation of the lysis buffer by diluting 10 μL of commercially available 14.3 M stock of β-mercaptoethanol in 90 μL

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of RNAse-free water. Because β-mercaptoethanol is volatile and a health hazard, handle in a well-ventilated hood wearing gloves, lab coat, and protective eyewear. 4. Mitochondrial lysis buffer (MLB): 260 mM sucrose, 100 mM KCl, 20 mM MgCl2, 10 mM Tris–HCl (pH 7.5), 1% TritonX100, 5 mM β-mercaptoethanol, and 1 Complete EDTAfree protease inhibitor. Prepared just before use. For 1 mL MLB (sufficient for lysing 10 mg of mitochondria), mix 178 μL of 50% sucrose (prepared above) with 50 μL of 2 M KCl, 20 μL of 1 M MgCl2, 10 μL 1 M Tris–HCl (pH 7.5), 100 μL of 10% Triton X-100, 3.5 μL 1.43 M β-mercaptoethanol, 20 μL of Complete EDTA-free protease inhibitor, and adjust the volume to 1 mL with 618.5 μL of RNAse-free water. If downstream analysis aims to detect mitochondrial RNAs, fortify against RNAse activity by adding RNAsin Plus RNAse inhibitor to 0.08 U/mL—as we have done in [17]—prior to adjusting the volume to 1 mL. Keep on ice until needed. 5. Light and heavy sucrose gradient buffers contain 10% or 30% sucrose, 100 mM KCl, 20 mM MgCl2, 10 mM Tris–HCl (pH 7.5), 5 mM β-mercaptoethanol, and 1 Complete EDTA-free protease inhibitor. To prepare 15 mL of each, which are sufficient for two gradients, mix 3 mL or 9 mL of 50% sucrose with 750 μL of 2 M KCl, 300 μL of 1 M MgCl2, 150 μL of 1 M Tris–HCl (pH 7.5), 300 μL of 50 Complete EDTA-free protease inhibitor (see above), 5.2 μL of 14.3 M β-mercaptoethanol, and adjust volume to 15 mL with RNAsefree H2O. These buffers have to be equilibrated to room temperature before preparation of the gradient. 6. Using the cleaned scissors (especially important if downstream RNA analysis is intended), cut the tips off of enough 1 mL pipette tips—30 tips are enough for fractionation of two gradients (see Note 5). Make sure that the tips are cut approximately at the same position to maintain reproducible fraction volumes. 2.4 TCA Precipitation, SDS-PAGE, and Immunoblot Analysis of the Gradient Fractions

One way to monitor the sedimentation of the mitoribosome and the two particles is by determining the migration of individual proteins from each particle within the gradient (see Note 6). Therefore, after gradient fractionation, the proteins in the fractions will be precipitated by the addition of trichloroacetic acid (TCA) and sodium deoxycholate. 1. Prepare 100% TCA by dissolving 500 g TCA in 227 mL of Milli-Q H2O. This will be a 5 Stock solution and store at 4  C.

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2. Prepare a 2% w/v stock solution of sodium deoxycholate by dissolving 1 g in 50 mL distilled H2O. Store at room temperature. Also, prepare a 1:10 dilution of the stock in distilled H2O (0.2%) for precipitation of the input sample. Due to its insolubility in the presence of low TCA concentrations, sodium deoxycholate will be used as a co-precipitant. 3. Ice-cold acetone will be used to wash away co-precipitated TCA and deoxycholate. Store at –20  C until needed. 4. SDS-PAGE and blotting equipment, as well as running and blotting buffers are supplied by different vendors. We use precast polyacrylamide gels, buffers, and equipment from BioRad. 5. Immunoblot washing buffer is composed of Tris-buffered saline (TBS) supplemented with 0.05% v/v Tween-20. 6. Immunoblot blocking solution: 5% milk in washing buffer. To prepare 50 mL, weight 2.5 g of skimmed milk powder and dissolve in 50 mL of washing buffer. 7. Different antisera are available for many of the ~80 mitoribosomal proteins. Whereas some of them can detect both human and mouse MRPS/L proteins, others are species-specific. Moreover, finding antisera that reliably recognize their target proteins may be challenging. When analyzing gradient fractions of mouse mitochondrial extracts, we have had success with the following MRPS and MRPL antisera: anti-MRPS15 (Proteintech, 17006-1-AP), anti-MRPS35 (Proteintech, 16457-1-AP), anti-MRPS25 (Proteintech, 15277-1-AP), anti-MRPS34 (Sigma, HPA042112), anti-MRPS24 (Proteintech, 16213-1-AP), anti-MRPL37 (Sigma, HPA025826), anti-MRPL12 (Proteintech, 14795-1-AP), and anti-MRPL44 (Proteintech, 16394-1-AP).

3

Methods

3.1 Isolation of Mitochondria from Mouse Heart

It is imperative that all buffers and the homogenizer are kept on ice; all steps should be carried out on ice and as quickly as possible. 1. Sacrifice the mouse by cervical dislocation and quickly place the explanted heart in a petri dish or other vessel containing 5–10 mL of ice-cold PBS. Remove as much blood as possible by squeezing it out using clean forceps. Destroying the heart structure in this process makes it easier to remove most of the blood. 2. Place the heart tissue in a new 10 cm petri dish and cut it in small pieces using a scalpel.

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3. Transfer the pieces to a 15 mL tube (Falcon or round-bottom) containing 14 mL of ice-cold MIB and keep on ice until all hearts have been processed in the above manner. 4. Transfer the heart tissue and MIB to a pre-chilled 15 mL glass homogenizer and disrupt the tissue with 20 strokes at 1000 rpm. 5. Return the homogenate to the 15 mL tube. 6. Rinse the homogenizer with Milli-Q water and repeat the homogenization of the next heart. Keep on ice and repeat steps 4 and 5 for all hearts. 7. Centrifuge the homogenates for 10 min at 1000  g at 4  C (see Note 7). 8. Recover the supernatant containing mitochondria to a new 15 mL tube. 9. Centrifuge for 10 min at 12,000  g at 4  C to recover crude mitochondria in the pellet fraction. 10. Discard the supernatant and resuspend the pellet in 10 mL of ice-cold MIB. 11. Repeat centrifugation step 8 and discard the supernatant. 12. Resuspend the pellet in 300–500 μL of MIB and determine the concentration of mitochondrial protein using your favorite assay—we use Bradford reagent and follow the manufacturer’s recommendations. Always determine the concentration of the controls and test samples at the same time. After this step, it is possible to snap-freeze the mitochondrial suspensions in liquid nitrogen for long-term storage at –80  C. This is useful if the mitochondria will be utilized in several different experiments. If sucrose gradients are the only downstream application for the isolated mitochondria, proceed to steps 13 and 14. 13. Aliquot 0.6–1.1 mg of heart mitochondrial suspensions in 1.5 mL RNAse-free tubes and centrifuge for 10 min at 12,000  g at 4  C. 14. Discard the supernatant and either proceed with mitochondrial lysis for sucrose gradient analysis or snap-freeze the pellets in liquid nitrogen and store at -80  C until needed. 3.2 Ultracentrifugation through a Linear Density Sucrose Gradient

1. Resuspend the mitochondrial pellets prepared above in freshly made MLB at a protein concentration of 10 mg/mL—for example, lyse 1.1 mg of mitochondria in 110 μL of MLB. Mix well by pipetting. 2. Incubate on ice for 20 min. Occasionally, flick the tubes to mix the lysate. 3. During the above incubation step, prepare the light and heavy gradient solutions.

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4. Clear the mitochondrial lysates by centrifugation for 45 min at 9200  g at 4  C. This step aims to remove detergent-insoluble material that can disrupt the gradient and can interfere with the sedimentation of the mitoribosomal particles and the monosome. While the lysates are clearing in step 4, prepare the sucrose gradients. As mentioned in the beginning, we use the BioComp’s Gradient Master equipment as other preparation methods are more laborious, time-consuming, and less reproducible. Gradient forming is carried out at room temperature. 5. Take the needed number of DEPC-treated polyallomer tubes (max 6 per run) using clean forceps and place them in the supplied two-step marker block; mark the tubes’ half-full point for short 4 mm plastic caps. 6. Place the tubes in the provided MagnaBase tube holder and fill each with the light gradient solution (~7 mL per tube) until it reaches ~2–3 mm above the half-full mark. Pay attention to avoid introducing bubbles that can attach to the tube walls as they will prevent proper gradient formation. 7. Attach the supplied DEPC-treated cannula to a 10 mL sterile syringe and withdraw ~8 mL of heavy sucrose gradient solution. Ascertain that no bubbles are trapped in the cannula or at the tip of the syringe by ejecting ~1 mL of the gradient buffer back into the tube. 8. Make sure that there are no drops forming at the end of the cannula and quickly insert it all the way down to the bottom of the centrifuge tube that contains the light sucrose solution. Slowly eject the heavy sucrose buffer while monitoring the upward migration of the visible interface between the two buffers until it reaches the half-full mark on the tube. During this step, also move the tip of the cannula up so that it is always ~5 mm or less under the buffer interface. This will ensure the least possible disturbance of the interface and mixing of the two gradient solutions during ejection of the heavy buffer and removal of the cannula. When done, stop pressing the plunger and withdraw the cannula in a single smooth motion. Some of the light buffer may overflow when the heavy solution reaches the half-full mark. This is normal and ensures that there will be no air left between the tube cap and the top of the light solution after inserting the cap. 9. Carefully close the tubes with the cleaned short plastic tube caps and remove all liquid trapped in the top reservoir of the caps. 10. Place the MagnaBase tube holder with the tubes on the magnetic plate of the Gradient Master and select the program for 10–30% sucrose gradient, with short cap and SW41Ti rotor.

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The magnetic plate will tilt to a preset angle of 81.5 and will mix the gradient solutions for 1.54 min. Then, the plate will return to its starting position indicating that the gradient is ready. 11. Transfer the MagnaBase tube holder with the tubes in a container with ice slurry to cool down the gradient for at least 30 min. The level of the slurry should be just under the edge of the tubes without reaching the plastic cups. 12. While the gradient is cooling, the preclearing of the lysates (started in step 4) will finish. Transfer the supernatants to new RNAse-free Eppendorf tubes and wait for the gradients to cool down. 13. After cooling the gradients, remove the silicone caps from the tubes and withdraw 400 μL from the top of the gradient. This is meant to avoid spilling of the gradient during the centrifugation even after adding the volume of the sample. Carefully load 100 μL of the lysate (equivalent to 1 mg of mitochondrial protein) on top of each gradient. Optional: to the remaining 10 μL (equivalent to 100 μg mitochondrial protein or one-tenth of the lysate loaded on the gradient), add 2 μL of the 100% TCA and 1 μL of 0.2% sodium deoxycholate, mix well, and incubate on ice for at least 20 min before placing at 20  C for storage (see Note 8). 14. Carefully wipe the sides of the tubes containing the gradient, load them into the buckets of a swing-out SW41Ti rotor, attach the buckets to the rotor, and centrifuge for 15 h at 71,000  g at 4  C. 3.3 Gradient Fractionation and TCA Precipitation (See Note 9)

While the centrifugation is ongoing, label the needed number of tubes. 1. At the end of the ultracentrifugation, carefully detach the buckets, extract the tubes, and place them in the MagnaBase tube holder. Using a cut pipette tip, collect 750 μL fractions from the top of the gradient. The volume of the first fraction should be increased to accommodate the sample volume—in this example, 750 + 100 μL sample volume. 2. To each fraction, add 100% TCA to final concentration of ~20% and sodium deoxycholate to a final concentration of 0.02%, mix by vortexing, and incubate for at least 20 min on ice before proceeding to the next step. At this stage, the samples can also be stored at –20  C. 3. To pellet the protein precipitates, centrifuge the samples including the input sample for 30 min at 12,000  g at 4  C. 4. Discard the supernatant in the appropriate waste container, add 1 mL of ice-cold acetone and mix by vortexing.

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5. Repeat steps 3 and 4 one more time. 6. Air-dry the pellets for no more than 30–60 min (see Note 10) and resuspend in 25 μL 1 SDS-PAGE loading buffer. If needed, rise the pH of the samples by adding 1 μL of 1 M Tris (see Note 11). 7. Mix vigorously until the pellets are dissolved. Incubation for 5 min at 65  C will improve pellet solubilization. Samples can be stored at –20  C until needed or you can proceed to SDSPAGE and immunoblotting. 3.4 Analysis of the Fractions by SDS-PAGE and Immunoblotting

For detection of mt-SSU and mt-LSU, we recommend choosing MRPS/MRPL proteins that have different electrophoretic mobility in SDS-PAGE such that the blotting membranes can be cut into several pieces according to the molecular weights of the target proteins. Thus, immunodetections can be carried out simultaneously for several MRPS/MRPL proteins in two or more gradients. This reduces the required volume of primary antisera preparations and the processing time of the immunoblots. 1. Fractionate the samples through a 12% criterion TGX gel (see Note 12) and transfer onto a PVDF or nitrocellulose membrane using your favorite SDS-PAGE/blotting system. Keep in mind that if using a PVDF membrane, it needs to be activated by incubation in absolute ethanol (see Note 13). 2. After the transfer, rinse the membrane in distilled H2O and visualize the proteins on the membrane by incubating for 5 min in commercially available Ponceau S solution. Most of the proteins will be concentrated in the first few gradient fractions. At this point, the membranes can be dried and stored protected from dust for 1–2 weeks. Alternatively, proceed with immunodetection. 3. Immunodetection is carried out using a standard protocol. First, reactivate the PVDF membranes (see above), rinse with distilled H2O, and pre-equilibrate in washing buffer for 10 min at room temperature. 4. Incubate the membranes in blocking solution for 1 h at room temperature. During this incubation, the proteins in the milk will bind to the protein-free regions of the PVDF membrane preventing nonspecific binding of the primary and secondary antisera to these regions. 5. Discard the blocking solution and incubate the membranes with the primary antisera diluted in blocking solution. The optimal length of the hybridization and the dilution of the antisera need to be determined empirically for each antibody. In our hands, the antisera suggested in Subheading 2.3.6 can

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be diluted at 1:1000 or 1:2000 in blocking buffer and hybridization can be carried out for at least 1 h. In most cases, the antibody dilutions can be stored at –20  C and reused several times. 6. Perform three 10 min washes with washing buffer. Make sure that the membranes are well covered with the buffer. 7. Incubate the membranes with a 1:5000 or 1:10,000 dilution of horse radish peroxidase (HRP)- or fluorophore-conjugated secondary antisera in blocking buffer depending on the downstream detection system—HRP-conjugated antisera are used for chemiluminescent detection whereas fluorophoreconjugated secondary antisera are used in multiplex fluorescence detection systems. 8. Repeat step 6 and proceed to develop the membranes. Figure 1 shows a typical sedimentation profile for the mt-SSU and mt-LSU and their association into the monosome. When adhering to the suggested protocol, mt-SSU proteins form a peak in fraction 6 whereas mt-LSU proteins form a peak in fraction 8 and trail into the monosomal region—peak fraction 10.

Fig. 1 Typical sedimentation profile of mt-SSU and mt-LSU proteins in a 10–30% linear density sucrose gradient. 100 μL of mitochondrial lysate (1 mg wild-type mitochondria lysed in buffer containing 1% Triton X-100) were layered on top of a 10–30% linear density sucrose gradient and centrifuged at 71,000  g for 15 h. Fractions were collected from the top of the gradient, TCA precipitated and fractionated through a 12% criterion TGX gel. Immunoblotting was carried out on PVDF membranes using antisera against MRPS35 and MRPS15 proteins from the small mitoribosomal subunit (mt-SSU) or MRPL37 and MRPL12 proteins from the large mitoribosomal subunit (mt-LSU). The top (fraction 1) and bottom (fraction 15) of the gradient are indicated as are the typical fractions in which mt-SSU, mt-LSU, and the monosome can be detected using our protocol. Input sample is not shown

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Notes 1. Several other methods for gradient preparation exist. For example, the freeze-thaw method involves overlaying three or more sucrose cushions with decreasing sucrose concentration followed by freezing and thawing at 4  C. We suggest the publications by Luthe [18] and Cooper et al. [19] as a starting points if preparing gradients by this method is more convenient to the reader. Alternatively, gradients can be prepared by overlaying equal volumes of heavy and light sucrose solutions in the centrifugation tubes, sealing the tubes with parafilm and slowly tipping them to one side. Let the gradients slowly diffuse for overnight at 4  C and gently return them to vertical position. The optimal diffusion time can be determined empirically by adding a colorant to the heavy solution and building a spectrophotometric profile of the dye distribution along the gradient. 2. The method for mitochondrial isolation and ribosomal analysis presented here can be directly applied to virtually all other mouse tissues with minimal adaptation. This method was also used to analyze mitoribosomal assembly in cultured patient fibroblasts with minimal modifications of the buffer compositions—for example see [20]. 3. Adding the protease inhibitor helps decrease protein degradation in the sample whereas we have decided to omit EDTA to avoid chelation of Mg2+ ions, which are important for monosome stability [15]. However, it has to be kept in mind that exclusion of EDTA can lead to contamination with cytosolic ribosomes [21]. This will not pose a problem for downstream analyses by immunoblotting with specific antisera or qRT-PCR/Northern blot but may be interfere if, for example, RNA absorption spectra are being recorded during fractionation. 4. Gradient forming conditions were empirically determined for ambient temperature and can vary as solutions’ viscosity may change with changing the temperature. Therefore, it is recommended that gradients are formed at room temperature and then cooled to 4  C. 5. Widening the pipette tip entrance will help ensure that the fraction is withdrawn at the level of the tip entrance into the gradient rather than from the underlying gradient zones. 6. Alternatively, sedimentation of the mitoribosome and mt-SSU and mt-LSU can be monitored spectrophotometrically by determining the absorption of rRNA at 260 nm either in the individual fractions or prior to fractionation using capillary spectrophotometric unit.

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7. Using a swing-out rotor though not essential is recommended as it will make it easier to transfer as much of the supernatant as possible in the next step. One can also dissolve the resulting pellet into additional volume of MIB and repeat the low speed centrifugation to further increase the mitochondrial yield. We find that a single centrifugation step combined with maximal, but careful, supernatant recovery yields more than enough mitochondria for gradient analysis. 8. The input sample can be used to calculate a protein or RNA—if RNA is extracted using Trizol LS—distribution profiles along the gradient relative to the starting material. 9. The gradient fractions can be split in order to carry out both, SDS-PAGE/immunoblotting and TaqMan qRT-PCR analysis. We developed this protocol while characterizing the role of the mRNA stability factor LRPPRC [17]. Briefly, one-third of each fraction is transferred to a new RNAse-free Eppendorf tube and the RNA in the fraction is extracted using Trizol LS. TaqManbased qRT-PCR is then used to build a profile of the relative abundance of the target mRNAs or rRNAs along the gradient. 10. Avoid overdrying the pellets as this may reduce their solubility in the SDS-PAGE loading buffer. 11. TCA remaining after the washes can decrease the pH of the samples. This will be visible by the yellow color of the acid indicator bromophenol blue in the sample buffer. Acidity of the sample may result in unspecific protein breakdown, decreased solubility, and aggregation or aberrant migration in SDS-PAGE. Adding 0.5–1 μL of 1 M Tris without pH adjustment to each sample is usually enough to rise the pH. 12. For the combination of antisera shown in Fig. 1, we used a 12% polyacrylamide precast Criterion TGX gel. These gels provide sufficient separation of the target MRPS/MRPL proteins and also have 18 wells that can accommodate up to 30 μL of sample making them suitable for loading all gradient fractions including the input sample. However, the choice of polyacrylamide concentration of the gel should be based on the molecular weight of target proteins. 13. Both PVDF and nitrocellulose work well in our hands. However, PVDF membranes allow to repeatedly remove (strip) the primary and secondary antisera without significant loss of detection sensitivity. References 1. Brown A, Amunts A, Bai XC, Sugimoto Y et al (2014) Structure of the large ribosomal subunit from human mitochondria. Science 346 (6210):718–722. https://doi.org/10.1126/ science.1258026

2. Greber BJ, Boehringer D, Leibundgut M, Bieri P et al (2014) The complete structure of the large subunit of the mammalian mitochondrial ribosome. Nature 515(7526):283–286. https://doi.org/10.1038/nature13895

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3. Kaushal PS, Sharma MR, Booth TM, Haque EM et al (2014) Cryo-EM structure of the small subunit of the mammalian mitochondrial ribosome. Proc Natl Acad Sci U S A 111 (20):7284–7289. https://doi.org/10.1073/ pnas.1401657111 4. Amunts A, Brown A, Toots J, Scheres SH et al (2015) Ribosome. The structure of the human mitochondrial ribosome. Science 348 (6230):95–98. https://doi.org/10.1126/sci ence.aaa1193 5. Greber BJ, Bieri P, Leibundgut M, Leitner A et al (2015) Ribosome. The complete structure of the 55S mammalian mitochondrial ribosome. Science 348(6232):303–308. https:// doi.org/10.1126/science.aaa3872 6. Rorbach J, Gao F, Powell CA, D’Souza A et al (2016) Human mitochondrial ribosomes can switch their structural RNA composition. Proc Natl Acad Sci U S A 113(43):12198–12201. https://doi.org/10.1073/pnas.1609338113 7. Chrzanowska-Lightowlers Z, Rorbach J, Minczuk M (2017) Human mitochondrial ribosomes can switch structural tRNAs - but when and why? RNA Biol 14(12):1668–1671. https://doi.org/10.1080/15476286.2017. 1356551 8. Metodiev MD, Lesko N, Park CB, Camara Y et al (2009) Methylation of 12S rRNA is necessary for in vivo stability of the small subunit of the mammalian mitochondrial ribosome. Cell Metab 9(4):386–397. https://doi.org/ 10.1016/j.cmet.2009.03.001 9. Metodiev MD, Spahr H, Loguercio Polosa P, Meharg C et al (2014) NSUN4 is a dual function mitochondrial protein required for both methylation of 12S rRNA and coordination of mitoribosomal assembly. PLoS Genet 10(2): e1004110. https://doi.org/10.1371/journal. pgen.1004110 10. Camara Y, Asin-Cayuela J, Park CB, Metodiev MB et al (2011) MTERF4 regulates translation by targeting the methyltransferase NSUN4 to the mammalian mitochondrial ribosome. Cell Metab 13(5):527–539. https://doi.org/10. 1016/j.cmet.2011.04.002 11. Wredenberg A, Lagouge M, Bratic A, Metodiev MD et al (2013) MTERF3 regulates mitochondrial ribosome biogenesis in invertebrates and mammals. PLoS Genet 9(1):e1003178.

https://doi.org/10.1371/journal.pgen. 1003178 12. Bruning JC, Michael MD, Winnay JN, Hayashi T et al (1998) A muscle-specific insulin receptor knockout exhibits features of the metabolic syndrome of NIDDM without altering glucose tolerance. Mol Cell 2(5):559–569 13. Sakamoto K, Gurumurthy CB, Wagner KU (2014) Generation of conditional knockout mice. Methods Mol Biol 1194:21–35. https://doi.org/10.1007/978-1-4939-12155_2 14. Cahill A, Baio DL, Cunningham CC (1995) Isolation and characterization of rat liver mitochondrial ribosomes. Anal Biochem 232 (1):47–55. https://doi.org/10.1006/abio. 1995.9962 15. O’Brien TW (1971) The general occurrence of 55 S ribosomes in mammalian liver mitochondria. J Biol Chem 246(10):3409–3417 16. Hamilton MG, O’Brien TW (1974) Ultracentrifugal characterization of the mitochondrial ribosome and subribosomal particles of bovine liver: molecular size and composition. Biochemistry 13(26):5400–5403. https://doi. org/10.1021/bi00723a024 17. Ruzzenente B, Metodiev MD, Wredenberg A, Bratic A et al (2012) LRPPRC is necessary for polyadenylation and coordination of translation of mitochondrial mRNAs. EMBO J 31 (2):443–456. https://doi.org/10.1038/ emboj.2011.392 18. Luthe DS (1983) A simple technique for the preparation and storage of sucrose gradients. Anal Biochem 135(1):230–232 19. Cooper AJ, Smallwood JA, Morgan RA (1984) The preparation of freeze-thaw density gradients with homogeneous solute concentrations. J Immunol Methods 71(2):259–264 20. Lake NJ, Webb BD, Stroud DA, Richman TR et al (2017) Biallelic mutations in MRPS34 lead to instability of the small mitoribosomal subunit and leigh syndrome. Am J Hum Genet 101(2):239–254. https://doi.org/10.1016/j. ajhg.2017.07.005 21. Lambowitz AM (1979) Preparation and analysis of mitochondrial ribosomes. Methods Enzymol 59:421–433

Chapter 4 Mouse Models for Studying Hippocampal Adult Neural Stem Cell Biology Fatih Semerci, Luke Parkitny, and Mirjana Maletic-Savatic Abstract The mammalian hippocampus shows a remarkable capacity for continued neurogenesis throughout life. Newborn neurons, generated by the radial neural stem cells (NSCs), are important for learning and memory as well as mood control. During aging, the number and responses of NSCs to neurogenic stimuli diminish, leading to decreased neurogenesis and age-associated cognitive decline and psychiatric disorders. Thus, adult hippocampal neurogenesis has been the subject of intense investigation, generating both excitement and controversy. Identifying the core molecular machinery responsible for NSC preservation is of fundamental importance if we are to use neurogenesis to halt or reverse hippocampal age-related pathology. Here, we briefly overview the most frequently used mouse models to study hippocampal neurogenesis and then focus on a unique mouse model that allows NSC-specific studies based on their unique expression of lunatic fringe (Lfng). The Lfng-eGFP and Lfng(BAC)-CreERT2;RCL-tdT transgenic mice provide us with an excellent tool to resolve long-standing questions regarding the properties of NSCs, such as their specific molecular composition, potency, and plasticity, in isolation from any other cell in the hippocampal neurogenic niche. Key words Adult hippocampal neurogenesis, Mouse models, Lunatic fringe, Neurogenic cascade

1

Introduction The adult human hippocampus is a site of life-long neurogenesis [1–10], which is estimated to replace about 30% of the dentate gurus over the life span [8]. Thus, it has attracted great attention as a possible target for regenerative therapies and novel therapeutic approaches for a range of neurological and neuropsychiatric diseases associated with impaired neurogenesis [11–19]. Based on rodent studies, generation of new neurons from primary neural stem cells (NSCs, radial NSC (rNSC), Type 1) located in the subgranular zone (SGZ) of the dentate gyrus progresses along a relatively well-defined path from NSC to mature granule cell. Although marked heterogeneity exists within each cell population [20–23], distinct stages can be defined by cell

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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morphology and specific markers [24]. Knowledge of these markers has resulted in the development of multiple in vivo rodent models to study neurogenesis, some of which are listed below. These mouse models have been used to gain greater insights into population dynamics, life cycle, migration, and interactions of different cell types in the neurogenic niche. 1.1 The Most Common Mouse Models Used to Study Hippocampal Neurogenesis

1. Nestin-based mouse models. Nestin is an intermediate filament protein that was initially identified in stem cells of the embryonic neural tube [25] but since has been shown to be expressed in regenerative neural and non-neural tissue [26–31], including in cancer stem cells [32]. Nestin-GFP reporter mice label GFP in the cytoplasm of cells in the dentate gyrus, subventricular zone, and the rostral migratory stream [33, 34]. Due to the abundance of cells expressing Nestin-GFP in the adult neurogenic niche, a nuclear localized CFP-expressing version, Nestin::CFPnuc, has been developed to increase the accuracy of quantification [35]. Doxycycline- and tamoxifen-inducible nestin strains have also been developed to permit temporal manipulations of NSCs and their progeny [36, 37]. 2. GFAP (glial fibrillary acid protein)- and GLAST (glial high affinity glutamate transporter)-based mouse models. GFAP and GLAST have been used to label adult NSCs of the dentate gyrus, as NSCs share these classical astrocytic markers [38]. Glast-CreERT2-based lineage tracing analysis revealed that the contribution of GLAST-expressing NSCs to the granule neuron population seemed to continuously increase over time rather than to plateau over time, as observed in NestinCreERT2 mice [39], highlighting that although both models label cells with similar morphologies, these are stem cells with different potentials. 3. ASCL1 (Aschaete-scute homolog 1)-based mouse model. ASCL1 is a transcription factor that is critical for the neuronal differentiation [40]. The ASCL1 reporter model Ascl1GFPnuc predominantly labels ANPs and only some NSCs [41, 42]. Indeed, expression of ASCL1 in adult NSCs is restricted to only activated NSCs that give rise only to the neuronal lineage [43]. 4. Gli1-based mouse model. Gli1 is a transcription factor and a sensitive readout of sonic hedgehog activity. It has been shown to be important for NSC proliferation [44]. The Glil1-CreERT2 line targets NSCs that give rise to both neurons and astrocytes [45]. 5. Hes5 (hairy/enhancer of split)-based mouse model. Hes5 gene is downstream target of Notch signaling, which is critical to determining differentiation, proliferation, and cell survival [46–48]. Hes5::GFP+ labels only a subset of NSCs and ANPs,

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reflecting the heterogeneity of NSCs and ANPs based on their Notch activity [49, 50]. Hes5::CreERT2 line has been used to delineate the proliferation dynamics of different ANP subtypes [51]. 6. Spot14: Thrsp (thyroid hormone-responsive protein; SPOT14)based mouse model. Spot14 has been shown to be important for NSC proliferation. SPOT-14::GFP mouse line labels mostly quiescent NSCs and ANPs, and Spot14::CreERT2 mice have been used to demonstrate that Spot14+ cells respond to neurogenic stimuli and give rise to neurons [52, 53]. In the remainder of this chapter, we will focus on a murine model that utilizes Lunatic fringe (Lfng), a regulator of important pathway for NSC biology, namely Notch signaling [54], that is exclusively expressed in the hippocampal NSCs [55]. It was proposed that the Lfng-mediated Notch signaling provides an avenue for communication between the NSC progeny and the mother NSCs to regulate preservation of the NSC pool and neurogenesis [55]. Thus, Lfng transgenic mouse models such as Lfng-eGFP and Lfng-CreERT2 provide a novel tool for studies of NSC biology.

2

Materials

2.1 Transgenic Reporter Mice for Selective Labeling of Neural Stem Cells 2.1.1 Lfng-eGFP Mouse 2.1.2 Lfng-CreERT2 Mouse

Lfng-eGFP (RRID:MMRRC_015881-UCD) mice were obtained from GENSAT [56]. RP23-270N2 BAC plasmids with eGFP inserted upstream of the Lfng locus (see Fig. 1) were injected into the pro-nuclei of RVB/N-fertilized oocytes. Hemizygous progeny were then bred with C57BL/6J mice (The Jackson Laboratory) to maintain the line in this background. To generate the Lfng-CreERT2 transgenic mouse model for lineage tracing, the parent plasmid containing the Lfng locus was modified by recombineering to insert a CreERT2 sequence in frame at the transcriptional start site (see Fig. 1) [55]. Lfng-CreERT2 mice were then generated by pronuclear injection of the modified BAC construct into fertilized FVB/N embryos.

Fig. 1 Map of bacterial artificial chromosome (BAC) construct that has been used to generate Lfng-CreERT2;RCL-tdT and Lfng-eGFP mice

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Hemizygous progeny were crossed to C57BL/6J mice. To allow tracing of the progeny of Lfng-expressing cells, LfngCreERT2 mice were subsequently crossed with Ai9 (RCL-tdT) reporter mice [57].

3

Methods

3.1 Brain Perfusion and Post-Fixation

1. Deeply anesthetize the animal according to applicable institutional guidelines and approved protocols. 2. Perfuse transcardially with 30 mL of ice-cold phosphate-buffered saline (PBS), pH 7.4 (see Notes 1 and 2), followed by 30 mL of 4% (w/v) ice-cold paraformaldehyde (PFA) in PBS (see Note 3). 3. Remove the brain and immerse in 15–20 mL 4% PFA either overnight at 4  C or 4 h at room temperature. Following this, remove the brain from PFA, rinse in PBS, then store in PBS at 4  C up to 6 months (see Notes 4 and 5).

3.2

Sectioning

1. Prepare 24-well plates (preferably black to limit fluorophore exposure to light) by filling each well with 1.5 mL of PBS with 0.1% sodium azide solution to prevent contamination (see Note 6). Five wells are required per brain hemisphere. 2. If only one hemisphere is to be used for further processing and analysis, then it needs to be selected randomly to account for possible variance between the right and left hippocampi. 3. Six hemispheres can be sectioned at a time by carefully organizing them on a vibratome cutting platform. To assist with the placement of hemispheres, the margins within which the razor travels can be labeled on the bottom of the glass plate so that they can be seen by the operator. A thin layer of super glue (ethyl 2-cyanoacrylate) is then spread across the glass surface and the thoroughly dried hemispheres (medial surface) are attached in a 2  3 pattern with gentle downward pressure to ensure proper attachment (see Note 7). 4. Trim and discard the most lateral 300μm of the hemispheres by gradual cutting, then start collecting 50μm-thick sections into the five consecutive wells: slice 1 to well 1, slice 2 to well 2, slice 3 to well 3, slice 4 to well 4, slice 5 to well 5, slice 6 to well 1, and so on. 5. Adjust the speed and the vibration intensity (amplitude) of the vibratome as required to attain a consistent cut throughout the sectioning process. Avoid changing the settings when the razor is placed over/within the tissue to prevent tissue damage. 6. At completion, expect each well to contain 12–14 slices that include the dentate gyrus.

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7. These consecutive slices are separated by approximately 250μm. Slices from any well may then be used as a representative sample to quantify the entire brain in an unbiased manner using stereology (see Subheading 3.4.2) [58]. 8. Cover the sections with Parafilm to prevent evaporation of the storage solution and store sections at +4  C. 3.3 Immunofluorescence

1. Antigen retrieval. For immunostaining of nuclear antigens such as BrdU, CldU, IdU, Ki67, and Olig2, sections should be treated with 2N HCl for 30 min at 37  C for antigen retrieval, then rinsed with PBS, incubated in 0.1M sodium tetraborate (pH 8.5) for 10 min at room temperature, then rinsed in PBS prior to blocking and permeabilizing (see Notes 8–10). 2. Initial permeabilization and blocking. For antigens that do not require antigen retrieval step, incubate the brain sections for 1 h at room temperature with PBS containing 0.1 to 0.5% Triton X-100 (PBST), 3% Bovine Serum Albumin (BSA), and 5% Normal Donkey Serum (NDS) (see Notes 11–13). 3. Incubation with primary antibodies. Incubate the slices with primary antibodies diluted in the permeabilization and blocking solution overnight at 4  C or 2 h at room temperature (see Note 14). Then, wash these sections three times in PBST for 5 min (see Note 15). 4. Incubation with secondary antibodies. Incubate the sections with secondary antibodies (targeted against the primary antibody species) diluted in the permeabilization and blocking solution at room temperature for 1.5– 2 h in the dark (see Note 16). Wash sections three times in PBST for 5 min. Counterstain with DAPI (5μg/mL) at 1:1000, to visualize nuclei (see Note 17). 5. Mounting the slices. Mount the slices on coated glass slides and remove excess liquid by gently shaking and drying. Add appropriate mounting media (such as 80μl of DakoCytomation Fluorescent Mounting Medium per slide), then carefully cover the slide with a cover glass avoiding air bubbles (see Note 18).

3.4 Confocal Microscopy and Stereology 3.4.1 Identification of a Cell Type

1. Neural Stem Cells (NSCs). As reported in several mouse models used to study hippocampal neurogenesis [24, 50, 59], NSCs in the SGZ have a characteristic triangular cell body from which a radial process that spans throughout the granule cell layer originates, ending with fine arborizations within the molecular layer of the dentate gyrus. Morphology of the radial processes (single long process or multiple short branches) and the range of their extension into the granule cell layer and/or the

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Fig. 2 Confocal photomicrograph of the dentate gyrus of an Lfng-eGFP mouse in low (a) and high magnification (b) showing the characteristic morphology of NSCs with arborizations ending in the molecular layer.

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molecular layer vary due to the heterogeneity of different populations of NSCs and the age of the animal [20, 22, 23]. These processes are positive for markers such as GFAP, Nestin, Vimentin, BLBP, while NSC nuclei are positive for Sox2 and for Hes5, Ascl1, Sox1, Sox21 in subset of NSCs [24, 60–62]. In Lfng-eGFP transgenic mice, the NSCs can be visualized easily due to expression of the GFP reporter protein (see Fig. 2a, b). In the Lfng-CreERT2;RCL-tdT lineage tracing mice, the NSCs can be visualized following Cre induction with tamoxifen, as they express tdTomato (see Fig. 2c, d). In the absence of a reporter, such as tdTomato-negative clones in the Lfng-CreERT2;RCL-tdT mice or simply in wild-type mice, radial GFAP-positive processes originating from Sox2-positive nuclei (see Fig. 2e) located in the SGZ are a reliable indicator of NSCs and can be used to quantify this population. 2. Amplifying Neuroprogenitors (ANPs). Since Lfng expression is exclusive to NSCs in the dentate gyrus, only very immediate ANP progeny show remnants of Lfng-eGFP expression in SGZ due to sequestration of the fluorescent protein during the formation of the progeny. Thus, in these mice, ANPs require independent staining to visualize. Type2a (early ANPs) and Type2b (late ANPs) can be identified as GFAP-negative, DCX-negative, Sox2-positive (early ANPs), or Tbr2(Eomes)positive (late ANPs) round cells in the SGZ without a radial process (see Fig. 3a, b). In Lfng-CreERT2; RCL-tdT transgenic mice, in which all progeny of the tamoxifen-induced NSC is expressing tdTomato, the ANPs are round tdTomato-positive, GFAP-negative, DCX-negative, Sox2-positive (early ANPs), or Tbr2(Eomes)-positive (late ANPs) cells located next to NSCs. 3. Neuroblasts (NB) and Immature Neurons (IN). Neuroblasts and immature neurons are DCX-positive cells with single or multiple process(es). In Lfng-CreERT2; RCL-tdT transgenic mice, the NBs and INs will also be labeled with tdTomato (see Fig. 3c). Their localization is not strictly restricted to the SGZ as they can be found in the lower layers of the molecular layer or the upper hilus near the SGZ. These cells are also positive for other markers, such as PSA-NCAM, Prox1, and NeuroD. 4. Granule Cells (GC). Granule cells are mature neurons in the dentate gyrus granule cell layer that have axons spanning through the hilus to CA3 region and dendrites branching in the molecular layer. They are NeuN and Calbindin-positive and ä Fig. 2 (continued) Following tamoxifen induction, Lfng-CreERT2;RCL-tdT mice labels NSCs (c and d). NSCs can be identified by their GFAP radial process (arrows) originating from Sox2+ nuclei (arrow heads) located in the SGZ. Scale bars 50μm for a and c; 20μm for b and d

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Fig. 3 (a) tdTomato+ NSC(arrow) with GFAP+ radial process and Sox2+ nuclei is next to its newborn Sox2+ ANP localized to SGZ (arrow head) (b) Lfng-eGFP-Tbr2+ late ANPs localized to SGZ. (c) 1 week after induction with tamoxifen Lfng-CreERT2;RCL-tdT mice can give rise to DCX+ immature neurons. (d) 1 month after tamoxifen injection, granule neurons with prominent dendritic arborizations are localized to the granule cell layer. (e) 1 month following tamoxifen induction S100β+ astrocytes could be observed in granular cell layer (arrow head) and molecular layer (arrow). (f) Iba1+ microglia (arrow heads) localized to the granule cell layer of dentate gyrus. Scale bars 20μm for a, b, c, and f; and 50μm for d and e

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in Lfng-CreERT2; RCL-tdT transgenic mice, they will also be labeled with tdTomato (see Fig. 3d). 5. Astrocytes. Astrocytes are GFAP-positive, S100β-positive glial cells localized in nearly all parts of the dentate gyrus with multiple processes of similar lengths (see Fig. 3e). 6. Microglia. Microglia are resident immune cells of the central nervous system and are an intrinsic part of the hippocampal SGZ neurogenic niche. They express Cd11b and Iba1 (Aif1), which can be used to visualize their ramified morphology (see Fig. 3f). Microglia can be found in all parts of the dentate gyrus, but they are mostly localized to the hilus and some parts of the SGZ due to their role in active phagocytosis of apoptotic ANPs and neuroblasts [63–65]. 3.4.2 Cell Counting

1. Estimating the total number of different cell types or cells that are positive for different antigens: thymidine analogs, proliferation markers, such as Ki67, PCNA, MCM2, and PH3, without falling into spatial bias requires the use of an optical dissector method [58]. 2. After completion of the immunofluorescence staining for one set of sections (see Subheading 3.2, step 4), an observer blind to the experimental groups should acquire 20μm thick optical sections (Z-stacks) with a confocal microscope. Cells located in the lowermost and the uppermost focal plane should be excluded from quantification. 3. After cells are counted in 13–14 sections, the total number of events is multiplied by five to obtain a total number across the dentate gyrus.

4

Notes 1. Ensure that peristaltic pump tubing is thoroughly flushed with 40 mL of dH2O after each cycle of perfusion to remove any remnant PFA. Failure to do so may cause premature fixation and result in incomplete removal of blood from the circulation. 2. Clearance of blood from the liver is a proxy indicator of good PBS perfusion. 3. Fasciculations in the hind limb and tail muscles are an indication of a good PFA perfusion. 4. If the tissue is stored for long time, a small amount of PFA (0.01%) may be added to the PBS to prevent the growth of bacteria and fungi.

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5. Long-term storage may be accomplished by storage at 20  C in a cryopreservation solution (FD Section Storage Solution Cat#PC101). 6. Avoid using sodium azide solution for storage if the sections are to be used in azide-dependent kits, such as EdU staining, to prevent high background signal. 7. Excess glue should be removed from the hemisphere/surface contact locations with a Kimwipe to prevent an obstructive barrier from forming around the tissue. 8. Although the HCl method is required for retrieval of the aforementioned nuclear antigens, other antigens might require a modified version, such as the use of a citrate buffer. 9. In slices from transgenic reporter animals that carry eGFP and/or tdTomato reporter proteins, harsh antigen retrieval processing may decrease or entirely deplete the fluorescent signal and antigenicity of the fluorescent proteins. Thus, staining against the fluorescent proteins with the following antibodies might be required to prevent loss of the signal: chicken anti-GFP (Aves Labs Cat# GFP-1020 RRID:AB_10000240) at 1:1000, rabbit anti-RFP (Rockland Cat# 600–401-379 RRID:AB_2209751) at 1:500, goat anti- RFP (SICGEN, Cantanhede, PORTUGAL) at 1:200. 10. The antigen retrieval process may also be completed after staining for antigens that do not require the retrieval step. 11. Choosing the right detergent and concentration is critical to achieving an effective stain. Thus, Triton X-100 may be replaced with Saponin or the concentration of Triton X-100 may need to be changed for optimal staining. Higher concentrations of Triton X-100 (0.3–0.5%) may be required to stain nuclear and cytoplasmic antigens in relatively thick (50μm) sections, and Saponin might be a better choice for sensitive antigens located in the cellular membrane. 12. The serum should be selected according to the host of the secondary antibodies to help minimize background signal. 13. BSA and NDS should be excluded from the permeabilization and blocking solution for staining against some antigens, such as PSA-NCAM, as BSA may block the antigenic regions of the polysialylated version of neural cell adhesion molecules. 14. Sections obtained from transgenic mice carrying fluorescent reporter proteins need to be protected from light during incubation with primary antibodies. 15. Diluted primary antibody solutions may be reused for subsequent staining; this needs to be determined based on tissue conditions and antibody performance in a case-by-case manner.

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16. If several primary antibodies are used for staining and are developed in closely related species, such as mouse and rat, pre-adsorbed secondary antibodies should be used to prevent cross-reactivity between different antibodies and thus nonspecific binding. 17. If any secondary antibody is conjugated to a fluorophore with an emission close to the emission of the nuclear counterstaining dye (DAPI or Hoechst), such as AlexaFluor® 405, it is important to avoid overlapping emission spectra. Thus, if sections are treated with DAPI or Hoechst, different brushes need to be used to mount and wash the slides. 18. Ensure that the cover glass thickness (~170μm thickness; 1.5 or 1.5H) is appropriately matched to the confocal microscope setup for optimal image acquisition.

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promotes hippocampal adult neurogenesis via the transcriptional repression of the Hes5 gene. J Neurosci 32(36):12543–12557 62. Venere M, Han YG, Bell R, Song JS, AlvarezBuylla A, Blelloch R (2012) Sox1 marks an activated neural stem/progenitor cell in the hippocampus. Development 139 (21):3938–3949 63. Sierra A, Encinas JM, Deudero JJ, Chancey JH, Enikolopov G, Overstreet-Wadiche LS et al (2010) Microglia shape adult hippocampal neurogenesis through apoptosis-coupled phagocytosis. Cell Stem Cell 7(4):483–495

64. Diaz-Aparicio I, Paris I, Sierra-Torre V, PlazaZabala A, Rodrı´guez-Iglesias N, Ma´rquezRopero M et al (2019) Microglia actively remodels adult hippocampal neurogenesis through the phagocytosis secretome. bioRxiv:583849 65. Lucassen PJ, van Dam A-M, Kandel P, Bielefeld P, Korosi A, Fitzsimons CP et al (2019) The orphan nuclear receptor TLX: an emerging master regulator of crosstalk between microglia and neural precursor cells. Neuronal Signal 3(2):NS20180208

Chapter 5 Retina as a Model to Study In Vivo Transmission of α-Synuclein in the A53T Mouse Model of Parkinson’s Disease Najiba Mammadova, Thierry Baron, Je´re´my Verche`re, Justin J. Greenlee, and M. Heather West Greenlee Abstract Parkinson’s disease is a neurodegenerative disorder characterized by accumulation of misfolded α-synuclein within the central nervous system (CNS). Retinal manifestations have been widely described as a prodromal symptom; however, we have a limited understanding of the retinal pathology associated with Parkinson’s disease. The strong similarities between the retina and the brain and the accessibility of the retina has potentiated studies to investigate retinal pathology in an effort to identify biomarkers for early detection, as well as for monitoring the progression of disease and efficacy of therapies as they become available. Here, we discuss a study conducted using a transgenic mouse model of Parkinson’s disease (TgM83, expressing human α-synuclein containing the familial PD-associated A53T mutation) to demonstrate the effect of the A53T α-synuclein mutation on the retina. Additionally, we show that “seeding” with brain homogenates from clinically ill TgM83 mice accelerates the accumulation of retinal α-synuclein. The work described in this chapter provides insight into retinal changes associated with Parkinson’s disease and identifies retinal indicators of Parkinson’s disease pathogenesis that could serve as potential biomarkers for early detection. Key words Parkinson’s disease, Retina, Mouse model, A53T, α-synuclein

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Introduction Parkinson’s disease (PD) is a degenerative disorder of the nervous system, characterized by two prominent disease processes: progressive degeneration of dopaminergic neurons of the substantia nigra pars compacta (SNpc) in the midbrain resulting in motor deficits (e.g., tremors, bradykinesia, and rigidity), and the accumulation of intraneuronal Lewy bodies, that contain misfolded α-synuclein [1– 3]. Epidemiological data over the past several decades has reported a dramatic increase in the occurrence of Parkinson’s disease, concluding that the worldwide burden of PD has more than doubled from 2.5 million patients in 1990 to 6.1 million patients in 2016 [4]. This has motivated researchers to identify sensitive and reliable

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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methods to screen individuals at risk prior to permanent damage. Although classically considered a movement disorder, emerging evidence reports a broad spectrum of non-motor manifestations associated with PD (e.g., cognitive dysfunction and hallucinations, sleep and mood disorders, gastrointestinal dysfunction, hyposmia, visual impairments, etc.) that occur decades before the onset of the hallmark motor symptoms [5–8]. Among these, hyposmia and visual dysfunction (impaired visual acuity, contrast sensitivity, and deficits in color vision have been most widely reported in PD [9– 12], with ~96% of PD patients reporting a reduced ability to smell or detect odors, and ~ 80% of PD patients reporting some extent of visual deficit [10, 13–15]. In the present study, we used a transgenic mouse model (TgM83) to investigate the underlying retinal changes that may contribute to the visual manifestations experienced by PD patients. Moreover, we aimed to identify retinal indicators of PD pathogenesis that could serve as potential biomarkers for early detection as well as for monitoring the effect of therapeutic interventions as they become available. Visual manifestations have been widely reported as a preclinical phase in several neurodegenerative processes including Parkinson’s disease and other synucleinopathies (dementia with Lewy bodies (DLB) and multiple system atrophy (MSA)), described to occur at least 10 years before the cardinal motor symptoms [9–19]. Additionally, several reports describe the accumulation of misfolded α-synuclein aggregates in the retinas of postmortem PD patients [20–24]. However, there is a scarcity of information on the distinguishing events that cause retinal change. The strong similarities between the retina and the brain has led to an increase of studies investigating retinal pathology resulting from PD, particularly using existing mouse models, both toxin-induced and genetic, that can recapitulate varying aspects of the human disease including the progressive accumulation of α-synuclein throughout the central nervous system (reviewed in [25]). Specifically, several studies have described α-synuclein phosphorylated at serine-129 (pSer129) [23, 24] in the retinas of PD patients. Phosphorylation of α-synuclein at serine 129 is one of the several post-translational modifications to α-synuclein known to occur in PD. Studies attribute this post-translational modification to the increased formation and self-propagation of α-synuclein aggregates, and Lewy body formation [26–30]. While the self-propagating ability of pathological a-synuclein (pSer129) has been widely postulated [1, 26, 31– 47], more conclusive evidence for the presence and spread of α-synuclein in the retina and its correlation to the cerebral a-synuclein burden is lacking. In a recent report, we described retinal pathology in a transgenic mouse model (TgM83) expressing the human Ala53Thr α-synuclein mutation. Specifically, we demonstrated that the accumulation of α-synuclein (pSer129) in retinas of

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5-month-old transgenic mice is accelerated upon “seeding” or intracerebral inoculation with brain homogenate from clinically ill transgenic mice [48]. This study is based on in vivo experiments using the TgM84 mouse model that demonstrated that “seeding” with an inoculum derived from the brain of clinically ill mice results in acceleration of α-synuclein-associated disease and shortening of survival time [36]. Our studies use the TgM83 mouse model to demonstrate the effect of α-synuclein propagation in the central nervous system, on the progression of retinal changes associated with the A53T mutation, specifically the accumulation of misfolded α-synuclein. This in vivo model provides insight into the effect of Parkinson’s disease on the retina and warrants further exploration into the potential use of retinal α-synuclein as a biomarker for early detection.

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Methods

2.1 In Vivo Animal Experiments

1. For these studies, we used TgM83 transgenic mice, that express the A53T-mutated human α-synuclein protein (B6; C3H-Tg[SNCA]83Vle/J, RRID:MGI:3603036, The Jackson Laboratory, Bar Harbor, ME, USA), and spontaneously develop motor deficits between 8 and 16 months of age.

2.2

1. Animals include: B6C3H mice (genetic background of the TgM83 mouse line) at 5 and 8 months of age; TgM83 mice inoculated with brain homogenate obtained from a 2-monthold healthy TgM83 mouse; C57Bl/6S mice, presenting a deletion of the α-synuclein locus [49] inoculated with brain homogenate obtained from a clinically ill TgM83 mouse to gauge for unspecific toxicity of the inoculum; homozygous TgM83 mice at 5 months of age (corresponding to a preclinical stage), 8 months of age, and clinically ill animals (~12–18 months of age); homozygous TgM83 mice inoculated at 2 months of age with brain homogenate obtained from two clinically ill (12 and 18 months) TgM83 mice, and euthanized 3 months after inoculation (5 months of age). Eyes are only collected from the following animals: B6C3H mice at 5 and 8 months of age; TgM83 mice at 5 months of age, 8 months of age, and clinically ill animals; and TgM83 mice inoculated at 2 months of age with brain homogenate obtained from two clinically ill (12 and 18 months) TgM83 mice, and euthanized 3 months after inoculation (5 months of age).

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Methods

3.1 Preparation and Inoculation of Brain Homogenates

1. Mice are euthanized by an intraperitoneal injection of lethal dose of sodium pentobarbital. All homogenates are prepared from half sagittally sectioned brains. Each half is weighed and placed in a ribolysis tube containing grinding balls. High Salt (HS) buffer [50 mM Tris–HCl, pH 7.5, 750 mM NaCl, 5 mM EDTA, 1 mM DTT, 1% phosphatase and protease inhibitor cocktails] is added to the brain halves to obtain 20% (wt/vol) homogenates. Brain extract inoculation samples are prepared using a mechanical homogenizer at 6.0 m/s for 23 sec twice. After the first 23 sec homogenization, the tubes contained the homogenates are placed on ice for 2 min before the second 23 s homogenization cycle. 2. Before brain extract inoculation, mice are anesthetized with a xylazine (10 mg/kg) and ketamine (100 mg/kg) mixture. Mice are subjected to intracerebral inoculation (IC) with 20 μL of 1% (wt/vol in glucose 5%) brain homogenates obtained from half of the brain of a clinically ill TgM83 mouse. Mouse brains are stored at 80  C for Western blot (WB) analyses, or fixed in buffered 10% formalin for immunohistochemical (IHC) studies [50, 51].

3.2 Clinical Monitoring of Mice

1. Care and housing of mice, as well as ethical approval is described as per Mougenot et al., 2011 [36]. Mice are monitored daily and clinically examined individually three times a week to detect any symptoms of TgM83 disease. Indications of clinical motor disease include: reduced mobility and/or persistent immobility, partial paralysis of the hind leg (e.g., freezing of a hind limb during spontaneous walking that lasts a few seconds), and balance impairment. Balance impairment can be examined by observing the spontaneous mobility of the mice (e.g., falling after rearing up, gently pushing the mouse on its side to see if the mouse can recover). These clinical signs progress to feeding difficulty and subsequent weight loss, prostration and/or hunched back, and general paralysis rapidly requiring sacrifice of the mouse for ethical reasons. A mouse is thus considered “clinically ill” as soon as the hind limb paralysis is detected by two independent observers [50].

3.3 In Vitro Animal Experiments

1. After mice are euthanized, eyes are enucleated before the brain has been removed. The eyelids are pulled apart to improve access to the eyeball surface. Curved forceps are placed under the globe in the orbit and is used to gently push the eye out of its socket and grip the globe from underneath without squeezing the globe. The globe is gently moved from left to right and upward until released from the socket, with the optic nerve still

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attached (see Notes 1 and 2). The eye globe is transferred into a Petri dish containing phosphate-buffered saline, after which the extraocular muscles are gently removed, and a single puncture wound is made into the cornea allowing fixative to penetrate the eye tissue. The entire eye is immediately placed into and fully immersed in 10% formalin for 24 h, then 70% ethanol. Eye globes are embedded in paraffin and sectioned sagittally at 4-μm onto Superfrost plus glass slides. 2. In the brain and the retina, phosphorylated α-synuclein (pSer129) was detected by WB and IHC analyses as previously described [48, 52]. For WB and IHC analyses, pSer129 was detected with anti-rabbit pSer129 α-synuclein monoclonal antibody (mAB) (WB 1:1000, IHC 1:300) (see Notes 3–5). 3.4 Representative Results

1. Survival curves were obtained by Kaplan–Meier method [52]. There was no statistical significance between survival times of uninoculated TgM83 mice (435 days old), and TgM83 mice inoculated with a brain homogenate from a healthy 2-month-old TgM83 mouse (359 days old) [36]. There was a significant difference between survival times of the aforementioned mice and TgM83 mice inoculated with brain homogenate obtained from two clinically ill (12 and 18 months) TgM83 mice (193 and 182 days old, respectively) (see Fig. 1a) [36]. 2. Western blot analysis of brains from TgM83 mice inoculated with brain homogenate obtained from two clinically ill (12 and 18 months) TgM83 mice, revealed pSer129 protein expression as early as 97 dpi (161 days old), while Western blot analysis of brains from age-matched uninoculated TgM83 mice revealed no pSer129 (see Fig. 1c) [36]. IHC analysis revealed accumulation of pSer129 in the dorsal raphe nucleus and lateral vestibular nucleus of 1) M83 mouse (339 days old) inoculated with brain homogenate obtained from a healthy mouse (2 months of age), 2) M83 mouse (198 days old) inoculated with brain homogenate obtained from a clinically ill mouse (12 months of age) and 3) M83 mouse (328 days old) inoculated with brain homogenate obtained from a clinically ill mouse (18 months of age) (see Fig. 1d) [36]. 3. Immunohistochemistry and Western blot analysis were used to detect α-synuclein (pSer129) and total α-synuclein in retinas of transgenic and control mice (see Fig. 2) [48]. Immunohistochemical analysis of retinas from B6C3H mice at 5 and 8 months of age revealed no pSer129 or total α-synuclein immunolabeling. In retinas of 5-month-old transgenic mice, pSer129 immunolabeling was evident only in the outer nuclear layer. However, pSer129 immunoreactivity was minimal, with two to three patches of labeling per retinal section. Compared

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Survival time (days) Uninoculated TgM83 (n=20) TgM83 inoculated with brain of a healthy (2m) mouse (n=11) TgM83 inoculated with brain of a sick (12m) mouse (n=9)

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TgM83 inoculated with brain of a sick (18m) mouse (n=12) C57BL/6S Δ α-syn inoculated with brain of a sick (18m) mouse (n=10)

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Fig. 1 (a) Survival times of TgM83 mice inoculated with brain homogenates from sick, 12-month-old (red dotted line) or 18-month-old (red line) TgM83 mice, compared with uninoculated TgM83 (black line) or TgM83 inoculated with a brain homogenate from a 2-month-old healthy TgM83 mouse (blue line). (b) A 161-day-old TgM83 mouse inoculated with a brain homogenate from a sick TgM83 mouse shows (97 days post-inoculation [d.p.i.]) paralysis of the hind limbs and impaired rotation to upright posture. (c) Western blot detection of insoluble pSer129 α-synuclein (α-syn) in brains of uninoculated sick (S) TgM83 mice and in brains of TgM83 mice inoculated with brain homogenates from sick TgM83 mice (1–4). pSer129 α-syn was not detected in brains of uninoculated healthy (H) (2- or 6-month-old) TgM83 mice. Below each lane, immunodetection of glyceraldehyde 3-phosphate dehydrogenase (GAPDH) using a mouse monoclonal antibody (mAb) [1:10,000] (Millipore, Molsheim, France) is shown, as well as a figure indicating the age at death of the corresponding mouse. (d) Immunohistochemical detection of pSer129 α-syn. The right upper photomicrograph attests the specificity of the labeling. The α-syn pathology is indicated by dystrophic neuritis and labeling of the neuronal perikarya (upper lane in the raphe nucleus of TgM83 mice which died when 339 or 198 days old). Bars: 16 μm. Lower panel: diffuse perikaryal inclusions as well as spheroid-like inclusions (arrows) in the gray matter of the lateral vestibular nucleus were also detected in a TgM83 mouse that died at 328 days old after inoculation with a brain homogenate from a sick TgM83 mouse or in a TgM83 mouse which died when 339 days old after inoculation with a brain homogenate from a healthy 2-month-old TgM83 mouse. Bars: 64 μm

to 5-month-old non-inoculated transgenic mice, retinas of inoculated 5-month-old transgenic mice revealed marked pSer129 immunoreactivity detected throughout the inner and outer retina. Retinas of 8-month-old and clinically ill transgenic mice had pSer129 immunolabeling similar to that of

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Fig. 2 Accumulation of α-synuclein. (a, b) Retinas of B6C3H mice had no phospho-α-synuclein (p129S) immunoreactivity. (c–f) Retinas of transgenic mice showed a similar trend, with sparse perinuclear and

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inoculated 5-month-old mice (see Fig. 2g–l). Western blot analysis of pSer129 and total α-synuclein protein revealed a similar trend. Retinas of inoculated 5-month-old mice, 8-month-old, and clinically ill mice had significantly more pSer129 (~two fold compared to control and 5-month-old transgenic mice; Fig. 2m, n). This result suggests accelerated accumulation of the pathogenic form of α-synuclein, pSer129, in the seeded model.

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Notes 1. In order for the retina to remain intact, mouse eyes must be enucleated before the brain has been removed. Care must be taken to ensure that the optic nerve is not damaged during enucleation. The use of curved dressing forceps makes this easier. When the curved forceps are placed under the globe in the orbit to push the eye out of its socket, it is best to grasp the extraocular muscles/connective tissue behind the globe to avoid squeezing the globe and damaging the retina [53]. A 30-gauge needle can be used to make a single puncture wound 1–2 mm into the eyeball [53]. Curved Westcott dissection scissors can be used to remove extraocular muscles and other connective tissues from the eye globe prior to fixation. 2. For optimum results, eye globes should be fixed in 10% formalin or Bouin’s fixative for 12–24 h. 3. Western blot analysis of pSer129 protein expression in retinas of transgenic mice was carried out using paraffin-embedded tissues as per Mammadova et al. 2019 [48]. However, similar results are anticipated with frozen tissues. 4. For protein extraction from paraffin-embedded retinas for Western blot analysis, retinal tissues are collected into 1.5 mL microcentrifuge tubes, and then deparaffinized and rehydrated

ä Fig. 2 (continued) extracellular phospho α-synuclein (p129S) immunoreactivity at 5 months of age, evident in the inner retina at the stage of clinical disease. (e) Retinas of inoculated mice showed increased α-synuclein (p129S) immunoreactivity, localized to ONL as well as the inner retina. (g, h) Retinas of B6C3H mice had no α-synuclein immunoreactivity. (i) Perinuclear and extracellular α-synuclein accumulation first evident at 5 months of age, localized to ONL. (j–l) As disease progressed, α-synuclein accumulation increased in intensity, distributed throughout the inner retina. (k) Retinas of inoculated TgM83 mice are comparable to retinas of clinically ill TgM83 mice, with α-synuclein localized to ONL and inner retina. Abbreviations: GCL ganglion cell layer, INL inner nuclear layer, IPL inner plexiform layer, OPL outer plexiform layer, ONL outer nuclear layer. Insets: High magnification images of α-synuclein immunoreactivity. Scale bars: 40 μm; insets 15 μm. (m) α-synuclein (pS129) (15 kDA), α-synuclein (15 kDA), immunoreactive bands. (n) Representative bar graph showing quantitative densitometric analysis of α-synuclein (pS129)/α-synuclein SD **P < 0.01 vs. B6C3H and TgM83 (5 m); ***P < 0.001 vs. B6C3H and TgM83 (5 m)

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using xylene, followed by a decreasing ethanol concentration gradient (100%, 95%, 70%) by incubation at room temperature in each solution for 10 min. After each incubation, tissue is pelleted at 14000  g for 3 min, and incubation/centrifugation is repeated two times. For optimum results, 100 microns of retinal tissue and at least 500 μL of solution (e.g., xylene or various concentrations of ethanol) must be used each time to ensure proper deparaffinization and rehydration. Further detail has been described by Guo et al. 2012 [54]. Alternatively, the Qproteome FFPE Tissue kit (Cat No./ID: 37623 Qiagen, Germantown, Maryland, USA) can potentially be used. 5. Besides immunohistochemistry and Western blot analysis, detection of disease-associated α-synuclein (pSer129) has been previously described in the TgM83 mouse model using an enhanced ELISA technique as per Be´temps et al., 2015 [51]. References 1. Recasens A, Dehay B (2014) Alpha-synuclein spreading in Parkinson’s disease. Front Neuroanat 8:159 2. Parkinson J (2002) An essay on the shaking palsy. J Neuropsychiatry Clin Neurosci 14 (2):223–236 3. Kempster PA, Hurwitz B, Lees AJ (2007) A new look at James Parkinson’s Essay on the Shaking Palsy. Neurology 69(5):482–485 4. Rocca WA (2018) The burden of Parkinson’s disease: a worldwide perspective. Lancet Neurol 17(11):928–929 5. Braak H et al (2004) Stages in the development of Parkinson’s disease-related pathology. Cell Tissue Res 318(1):121–134 6. Guo L et al (2018) Oculo-visual abnormalities in Parkinson’s disease: possible value as biomarkers. Mov Disord 33(9):1390–1406 7. Mahlknecht P, Seppi K, Poewe W (2015) The concept of prodromal Parkinson’s disease. J Parkinsons Dis 5(4):681–697 8. Poewe W (2008) Non-motor symptoms in Parkinson’s disease. Eur J Neurol 15:14–20 9. Armstrong RA (2011) Visual symptoms in Parkinson’s disease. Parkinson’s Dis 2011:908306 10. Archibald NK et al (2009) The retina in Parkinson’s disease. Brain 132(5):1128–1145 11. Bodis-Wollner I (2013) Foveal vision is impaired in Parkinson’s disease. Parkinsonism Relat Disord 19(1):1–14 12. Ridder A et al (2017) Impaired contrast sensitivity is associated with more severe cognitive impairment in Parkinson disease. Parkinsonism Relat Disord 34:15–19

13. Jones RD, Donaldson IM, Timmings PL (1992) Impairment of high-contrast visual acuity in Parkinson’s disease. Mov Disord 7 (3):232–238 14. Matsui H et al (2006) Impaired visual acuity as a risk factor for visual hallucinations in Parkinson’s disease. J Geriatr Psychiatry Neurol 19 (1):36–40 15. Sartucci F, Porciatti V (2006) Visual-evoked potentials to onset of chromatic red-green and blue-yellow gratings in Parkinson’s disease never treated with L-dopa. J Clin Neurophysiol 23(5):431 16. De Groef L, Cordeiro MF (2018) Is the eye an extension of the brain in central nervous system disease? J Ocul Pharmacol Ther 34 (1-2):129–133 17. Normando EM et al (2016) The retina as an early biomarker of neurodegeneration in a rotenone-induced model of Parkinson’s disease: evidence for a neuroprotective effect of rosiglitazone in the eye and brain. Acta Neuropathol Commun 4(1):86 18. Schneider JS, Ault ME, Anderson DW (2014) Retinal Pathology detected by optical coherence tomography in an animal model of Parkinson’s disease. Mov Disord 29 (12):1547–1551 19. Turcano P et al (2019) Early ophthalmologic features of Parkinson’s disease: a review of preceding clinical and diagnostic markers. J Neurol 266(9):2103–2111 20. Bodis-Wollner I et al (2014) α-synuclein in the inner retina in parkinson disease. Ann Neurol 75(6):964–966

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21. Bodis-Wollner I, Miri S, Glazman S (2014) Venturing into the no-man’s land of the retina in Parkinson’s disease. Mov Disord 29 (1):15–22 22. Ho CY et al (2014) Beta-amyloid, phospho-tau and alpha-synuclein deposits similar to those in the brain are not identified in the eyes of Alzheimer’s and Parkinson’s disease patients. Brain Pathol 24(1):25–32 23. Beach TG et al (2014) Phosphorylated α-synuclein-immunoreactive retinal neuronal elements in Parkinson’s disease subjects. Neurosci Lett 571:34–38 ˜ o-Lizara´n I et al (2018) Phosphorylated 24. Ortun α-synuclein in the retina is a biomarker of Parkinson’s disease pathology severity. Mov Disord 33(8):1315–1324 25. Veys L et al (2018) Retinal α-synuclein deposits in Parkinson’s disease patients and animal models. Acta Neuropathol:1–17 26. Angot E et al (2010) Are synucleinopathies prion-like disorders? Lancet Neurol 9 (11):1128–1138 27. Oueslati A (2016) Implication of alphasynuclein phosphorylation at S129 in synucleinopathies: What have we learned in the last decade? J Parkinsons Dis 6(1):39–51 28. Chen L et al (2009) Tyrosine and serine phosphorylation of α-synuclein have opposing effects on neurotoxicity and soluble oligomer formation. J Clin Invest 119(11):3257 29. Fujiwara H et al (2002) α-Synuclein is phosphorylated in synucleinopathy lesions. Nat Cell Biol 4(2):160–164 30. Smith WW et al (2005) α-Synuclein phosphorylation enhances eosinophilic cytoplasmic inclusion formation in SH-SY5Y cells. J Neurosci 25(23):5544–5552 31. Desplats P et al (2009) Inclusion formation and neuronal cell death through neuron-toneuron transmission of α-synuclein. Proc Natl Acad Sci 106(31):13010–13015 32. Lee S-J et al (2010) Cell-to-cell transmission of non-prion protein aggregates. Nat Rev Neurol 6(12):702 33. Lee H-J et al (2008) Assembly-dependent endocytosis and clearance of extracellular a-synuclein. Int J Biochem Cell Biol 40 (9):1835–1849 34. Dunning CJ, George S, Brundin P (2013) What’s to like about the prion-like hypothesis for the spreading of aggregated α-synuclein in Parkinson disease? Prion 7(1):92–97 35. El-Agnaf OM et al (2003) α-Synuclein implicated in Parkinson’s disease is present in extracellular biological fluids, including human plasma. FASEB J 17(13):1945–1947

36. Mougenot A-L et al (2012) Prion-like acceleration of a synucleinopathy in a transgenic mouse model. Neurobiol Aging 33 (9):2225–2228 37. Luk KC et al (2012) Pathological α-synuclein transmission initiates Parkinson-like neurodegeneration in nontransgenic mice. Science 338 (6109):949–953 38. Masuda-Suzukake M et al (2013) Prion-like spreading of pathological α-synuclein in brain. Brain 136(4):1128–1138 39. Woerman AL et al (2018) α-synuclein: multiple system atrophy prions. Cold Spring Harb Perspect Med 8(7):a024588 40. Katorcha E et al (2017) Cross-seeding of prions by aggregated α-synuclein leads to transmissible spongiform encephalopathy. PLoS Pathog 13(8):e1006563 41. Luk KC et al (2012) Intracerebral inoculation of pathological α-synuclein initiates a rapidly progressive neurodegenerative α-synucleinopathy in mice. J Exp Med 209 (5):975–986 42. Dehay B et al (2015) Targeting α-synuclein for treatment of Parkinson’s disease: mechanistic and therapeutic considerations. Lancet Neurol 14(8):855–866 43. Oliveras-Salva´ M et al (2013) rAAV2/7 vectormediated overexpression of alpha-synuclein in mouse substantia nigra induces protein aggregation and progressive dose-dependent neurodegeneration. Mol Neurodegener 8(1):44 44. Polinski NK et al (2018) Best practices for generating and using alpha-synuclein pre-formed fibrils to model Parkinson’s disease in rodents. J Parkinson’s Dis 8(2):303–322 45. Volpicelli-Daley LA et al (2016) How can rAAV-α-synuclein and the fibril α-synuclein models advance our understanding of Parkinson’s disease? J Neurochem 139:131–155 46. Volpicelli-Daley LA et al (2011) Exogenous α-synuclein fibrils induce Lewy body pathology leading to synaptic dysfunction and neuron death. Neuron 72(1):57–71 47. Okuzumi A et al (2018) Rapid dissemination of alpha-synuclein seeds through neural circuits in an in-vivo prion-like seeding experiment. Acta Neuropathol Commun 6(1):96 48. Mammadova N et al (2019) Accelerated accumulation of retinal α-synuclein (pSer129) and tau, neuroinflammation, and autophagic dysregulation in a seeded mouse model of Parkinson’s disease. Neurobiol Dis 121:1–16 49. Specht CG, Schoepfer R (2001) Deletion of the alpha-synuclein locus in a subpopulation of C57BL/6J inbred mice. BMC Neurosci 2 (1):11

Mouse Model of Parkinson’s Disease 50. Sargent D et al (2017) ‘prion-like’propagation of the synucleinopathy of M83 transgenic mice depends on the mouse genotype and type of inoculum. J Neurochem 143(1):126–135 51. Be´temps D et al (2015) Detection of diseaseassociated α-synuclein by enhanced ELISA in the brain of transgenic mice overexpressing human A53T mutated α-synuclein. J Vis Exp 99:e52752 52. Mougenot A-LJ et al (2011) Transmission of prion strains in a transgenic mouse model

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Chapter 6 Promoting Pro-Endocrine Differentiation and Graft Maturation Following Surgical Resection of the Mouse Pancreas Mugdha V. Joglekar, Rohan R. Patil, Sarang N. Satoor, Wilson K. M. Wong, Mahesh S. Karandikar, and Anandwardhan A. Hardikar Abstract Type 1 diabetes (T1D) is an autoimmune disease, where insulin-producing β-cells in the pancreas are inappropriately recognized and destroyed by immune cells. Islet transplantation is the most successful cellbased therapy for T1D individuals who experience frequent and severe life-threatening hypoglycemia. However, this therapy is extremely restricted owing to the limited availability of donor pancreas. In recent years, significant progress has been made in generating β-cells from stem/progenitor cells using different approaches of in vitro differentiation. The insulin production from such in vitro generated β-cells is still far less than that observed in islet β-cells. We employed a novel strategy to improve the efficiency of progenitor cell differentiation by performing partial mouse pancreas resection after transplanting in vitro generated insulin-producing cells under the kidney capsule of these mice. Pancreas resection (pancreatectomy) has been shown to induce regenerative pathways, leading to regeneration of almost the entire resected pancreas over 3–5 weeks in mice. We found that in our method, regenerating mouse pancreas promotes better graft differentiation/maturation and insulin production from transplanted cells. In this chapter, we detail the protocols used for transplantation of in vitro differentiated cells in immunocompromised mice, partial pancreatectomy in host (NOD scid) mice, and assessment of graft function. We believe that our protocols provide a solid platform for further studies aimed at understanding growth/differentiation molecules secreted from regenerating pancreas that promote graft maturation. Key words Pancreatectomy, Diabetes, β-cells, Insulin, Pancreatic regeneration, Transplantation

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Introduction According to the World Health Organization report, the prevalence of adults affected with diabetes has quadrupled since 1980 [1] and according to another report, it is estimated to increase to 693 million by 2045, contributing to an ever increasing social, financial, and health care system burden [2]. There are two major forms of diabetes, where hyperglycemia results either due to loss (type 1 diabetes: T1D) or dysfunction (type 2 diabetes: T2D) of

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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pancreatic insulin-producing β-cells. Even with latest treatments for glucose, lipid control and blood pressure the complications of diabetes are common and impose increased personal suffering and economic burden [3, 4]. T1D is an autoimmune disease, where self-reactive (auto)immune cells destroy insulin-producing cells in the pancreas [5]. Islet transplantation is the most successful cell-based therapy for individuals who present with frequent and severe hypoglycemia [6]. Clinical studies involving islet transplantation provided proofof-principle that restoring adequate β cell mass in individuals with T1D can maintain normal blood glucose levels without the need of exogenous insulin injections for up to 5 years [7–10]. However, the major challenges in islet transplantation include limited availability of donor pancreas and undesired side effects of immunosuppressive medications administered to prevent graft rejection. There is an urgent need to generate more numbers of insulin-producing and glucose-responsive β-cells in the laboratory to meet the increasing demand of transplantation in individuals with T1D. In recent years, there is significant progress in generating insulin-producing cells from stem/progenitor cells [11] or endodermal cells [12]. The insulin-production from these in vitro differentiated cells is however much lower than the actual pancreatic islets as most in vitro differentiation protocols do not always present the full differentiation potential. To overcome this limitation, we employed a novel strategy of performing pancreas resection following transplantation of in vitro differentiated stem/progenitor cells in mice [13, 14]. Our results demonstrate that regenerating mouse pancreas promotes better graft function and enhances insulin production from transplanted cells, probably as a result of growth/ differentiation promoting factors secreted by regenerating pancreas. Pancreas regeneration is observed in animal models of tissue injury, where both exocrine (acinar, endothelial, and duct cells) and endocrine (islet cells) pancreatic compartments are seen to grow back after damage [15, 16]. To investigate islet cell proliferation and regeneration, three different injury models are extensively studied: surgical resection of pancreas (pancreatectomy), pancreatic duct ligation, and chemically induced islet destruction. Our group uses a partial pancreatectomy protocol that involves removing around 70% of the mouse pancreas. Using this method, we demonstrated that pancreatic regeneration occurs under normal conditions as well as in diabetes [17]. We further unravelled that pancreas regeneration in mice is not similar to that of mouse embryonic pancreatic development but involves microRNA-mediated regulation of neurogenin 3 [18, 19]. We also observe improved insulin production from transplanted cells following pancreatectomy [13, 14], thereby making it an important model to understand the underlying factors and mechanisms involved in β-cell regeneration.

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In this chapter, we detail the protocols used for cell transplantation, partial pancreatectomy, and functional assessment of the graft following transplantation in mice. We believe that our protocols will also be very useful to drive research into the area of identifying factors from regenerating pancreas that promote graft maturation and insulin secretion.

2

Materials

2.1 Cell Transplantation

1. NOD scid mice (preferably males). 2. 1.7 mL centrifuge tubes. 3. 0.2 mL sterile pipette tips. 4. Sterile drapes and gloves. 5. Sterile surgical instruments including fine tip forceps, scissors, and scalpel blades. 6. Isoflurane (see Note 1) with appropriate vaporizer, oxygen cylinder, and charcoal filter canister. 7. Weighing balance. 8. Saline: 0.9% sodium chloride in water. 9. Syringes. 10. Disinfectants such as ethanol, betadine, and chlorhexidine. 11. Electric shavers, razor blades, or hair removal cream. 12. 1 mg/mL stock solution of buprenorphine in sterile saline solution or 5 mg/mL stock of commercially available diclofenac sodium in sterile saline solution can also be used. 13. 5-0 Absorbable sutures. 14. Wound clips. 15. Topical antiseptic ointment such as soframycin. 16. 10–20 mg/mL stock of commercially available gentamicin, ampicillin, and cloxacillin in sterile saline. 17. Heating mat/pad. 18. Heating lamp.

2.2

Pancreatectomy

1. NOD scid animals who received cell transplant earlier. 2. Sterile drapes and gloves. 3. Sterile surgical instruments including fine tip forceps, scissors, and scalpel blades. 4. Isoflurane (see Note 1) with appropriate vaporizer, oxygen cylinder, and charcoal filter canister. 5. Weighing balance.

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6. Saline. 7. Surface disinfectant such as ethanol. 8. Razor blades or hair removal cream. 9. 0.1 mg/mL Buprenorphine in sterile saline solution. 10. 10–20 mg/mL stock of commercially available gentamicin, ampicillin, and cloxacillin in sterile saline. 11. 5-0 Absorbable sutures. 12. Wound clips. 13. Topical antiseptic ointment such as soframycin. 14. Heating mat/pad. 15. Heating lamp. 2.3

Insulin ELISA

1. 25 mg/mL stock solution of glucose in saline. 2. Syringes. 3. 1.7 mL centrifuge tubes. 4. Human insulin ELISA kit (Mercodia). 5. Magnetic stirrer. 6. Vortex mixer. 7. Microplate shaker (700–900 cycles per minute, orbital movement). 8. Glassware including beakers and measuring cylinders. 9. MilliQ water. 10. Blotting paper. 11. 1 and 0.2 mL pipette tips. 12. Spectrophotometer.

3

Methods Prepare cells for transplantation in a sterile biosafety class II cabinet. Perform all surgical procedures and cell manipulations within a class II biosafety cabinet in designated animal house. UV sterilize the biosafety cabinet prior to use. Use aseptic drapes, gloves, and autoclaved/baked or disposable surgical instruments at all times.

3.1 Cell Transplantation

One can select cells of their choice for transplantation. We used in vitro-differentiated umbilical cord blood mononuclear cells [13] and human bone-marrow-derived mesenchymal stem cells [14] in our study. Details of generating such differentiated cells can be found in respective articles and are beyond the scope of this protocol. We use immunocompromised NOD scid animals for transplanting human cells, thereby avoiding need of any immunosuppressive drugs and also the fear of graft rejection (see

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Fig. 1 A stepwise illustration of the transplant procedure. (a) Prepare the cells for transplantation with ~15 microliters of whole blood from the host (recipient) mouse, which is pipetted around the cell pellet to engulf the cells within a clot. Allow the clot to form. (b) Pull out the left kidney and stabilize it above the incision by carefully inserting a P20 pipette tip underneath. (c) Make an incision in the capsule (shown by the arrowhead) to gently pick up the membrane with a fine forceps. Place the clot containing the cell pellet at the entry of this pocket once you have made this pocket by following the directions in the text. (d) Gently massage the exterior of the capsule to have the graft containing clot into the base of this pocket. Return the kidney to the peritoneal cavity and follow the directions as specified in text. Make sure to hydrate the kidney/capsule at all times

Note 2). In diabetes research, insulin-producing cells have been transplanted in multiple sites such as omentum, subcutaneous space, portal vein, muscle, liver, kidney capsule, and splenic capsule [20, 21] as well as in encapsulated micro- and macro-vesicles [22]. We have optimized our methods for transplanting cells under the kidney capsule. All procedures and protocols need to be approved by local ethics committee before commencing the experiments. A stepwise illustration of this protocol is provided in Fig. 1. 1. Harvest the cells to be transplanted in a 1.7 mL centrifuge tube by gently spinning them at 400
g for 3 min. 2. Remove as much supernatant as possible, leaving just a few microliters so as not to dry the pelleted cells. 3. Keep the cells on wet ice until the animals are prepared for surgery. 4. Fast the NOD scid animals (around 8 weeks of age) overnight before the surgery (see Note 3). 5. Administer an injection of buprenorphine (0.1 mg/kg) prior to surgery to ensure pain relief. 6. Place the animal on a heat mat with a double layer of gauze above the heat mat throughout the surgery to ensure maintenance of body temperature (without any burns). 7. Anesthetize the mice using 3–5% isoflurane in oxygen at 2.5 mL/min initially and later reduce the rate of isoflurane to 2% once the mouse is anesthetized. 8. Make sure that animals are under proper anesthesia before initiating surgery (see Notes 4 and 5). 9. Clean the end of tail thoroughly with disinfectant and gently rub the tail to enhance circulation in that area. With a scalpel snip the tail end just to get 2–3 drops of blood (10–20 μL).

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Quickly aspirate this blood with a P20 pipette and dispense it slowly around the cell pellet. Allow the blood to clot with the cell pellet engulfed within the blood clot. Ensure that the cell pellet is engulfed within the clot by trying to lift the clot with a blunt forceps or with a sterile pipette tip. Leave the tube containing the cell pellet clot on ice (see Fig. 1a). 10. Keep the animal in lateral position (left side up) and thoroughly clean the lower abdomen area with disinfectant swabs (ethanol and betadine). Shave the area without causing any cuts to the skin (see Note 6). 11. Use a sterile surgical scalpel to make a small midline skin incision of 1 cm length in the shaved area of the lower abdomen, just above the left kidney. 12. Make a similar but smaller cut (not more than 40 mm) on the peritoneal wall and gently pull out the kidney from abdominal cavity. Place the kidney on the peritoneal cut (see Note 7) and stabilize it on top of the incision by placing a pipette tip underneath the kidney so as to prevent it from falling back into the peritoneal cavity (see Fig. 1b). 13. Keep a few syringes filled with saline and ensure that the kidney remains moist at all times by bathing it intermittently (as required) with saline. Since the next steps involve making a pocket in this outermost layer/capsule (renal fascia), it is important to ensure that the capsule does not dry out. 14. Using a sterile scalpel, make a very small (2–3 mm), superficial cut on kidney capsule from the cranial pole extending towards the caudal pole. 15. Use a fine jewelers’ forceps to pick up the membrane and then add a few drops of saline (using a needleless syringe) under the kidney capsule/fascia. 16. Use a pipette tip to gently press down on the kidney while holding the capsule with the fine forceps so as to create a pocket under the capsule (see Note 8). 17. Use a curved blunt forceps or pipette tip to gently lift the blood +cell clot and place it just at the opening of the pocket made under the kidney capsule. Keep bathing the kidney with saline (see Fig. 1c). 18. Gently push the clot inside the pocket and then using the side of a pipette tip, gently massage the graft/clot from outside of the capsule so as to push the clot towards the bottom of this pocket. This will ensure that the transplanted cells will remain under the kidney capsule and would not be flushed away in the abdominal cavity. Ensure that the kidney capsule is moist at all times so as to prevent the capsule from rupturing (see Fig. 1d).

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19. Remove the pipette tip that was inserted under the kidney (in step 12 above) and then gently place the kidney back inside the peritoneal cavity by guiding it through the incision. 20. Close the peritoneum with absorbable sutures without damaging any internal organs and also without making any cuts on the membrane. 21. Close the skin cut with 2–4 wound clips and apply antiseptic cream (soframycin). Turn off the isoflurane supply. 22. Place mice in a clean cage with easy access to food and water. 23. Keep the cage in an area where a heat gradient (warm to room temperature) is generated by using either a heating pad or lamp. Mice are kept in the warm area until they wake up completely from the anesthesia and would move to the cooler side of the cage once recovered (see Note 9). 24. Administer an intraperitoneal injection of gentamycin (3 mg/ kg body weight), ampicillin, and cloxacillin (20 mg/kg body weight). 25. Monitor the operated animal at least twice every day for two days post-surgery and administer buprenorphine analgesic (0.1 mg/kg) daily if any signs of pain or discomfort are seen in the animals. 26. Follow up the operated animals daily for 1 week, and thrice a week for the next 6 weeks post-surgery for wound inflammation, infection, bleeding or discharge, reduced movement, abnormal breathing, abnormal grooming, hunched posture, other signs of pain (squinting of eyes, ears flattened), and abnormal behavior (see Note 10). 27. Remove the clips or sutures 8–10 days post-surgery, when the wound is healed. 28. Randomize some of the transplanted animals for partial pancreatectomy while maintain the others without partial pancreatectomy as controls. 3.2 Partial Pancreatectomy

1. Perform the partial pancreatectomy procedure after 2 weeks of cell transplantation surgery, when the operated animals display all signs of recovery and normal movement and feeding patterns. A stepwise illustration is shown in Fig. 2. 2. Perform pre-operative steps as detailed above in Subheading 3.1 (steps 4–8). 3. Place the animal in lateral position with left side up and shave the area just below the rib cage after cleaning with disinfectant swabs (see Fig. 2a, b). 4. Use the sterile surgical scalpel/blade to make a small skin incision of 0.5 cm to 1 cm length in the shaved area (see Fig. 2c).

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Fig. 2 A stepwise presentation of surgical pancreatic resection. (a) Anesthetize the mouse, keep in the left lateral position, and clean the area just below the rib cage. (b) Shave the clean area and make a small cut on the skin and to the peritoneal wall (c). (d) Gently pull out the spleen and the pancreas attached to the spleen. (e) Pull away the pancreas from the spleen with a blunt forceps without causing any bleeding. (f) Cut majority of splenic pancreas (90%) up to the main pancreatic duct. (g) Place the spleen back inside the body only when there is no bleeding. (h) Close the peritoneum with absorbable 5-0 sutures and close the skin cut with two wound clips (i). Originally published in our article in Developmental Biology [18] and reproduced with permission from the Elsevier

5. Make a similar but smaller cut (not more than 40 mm) on the peritoneal wall and gently pull out the spleen and the pancreas attached to the spleen (see Note 11) (see Fig. 2d). 6. Gently place a curved blunt forceps under the pancreas and another under the spleen so as to pull away the pancreas from the spleen. Continue to do so until you get to the major pancreatic duct (see Fig. 2e). At this point, make a cut leaving behind some of the ductal portion of the pancreas. The resected pancreas would constitute around 90% of the splenic pancreas (see Fig. 2f). If you observe bleeding, place a sterile gauze pad and apply pressure to stop the bleed. If you observe significant bleeding in your hands, it is best to use one of the smaller battery-operated cautery devices for making this cut.

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7. Place the spleen inside the body cavity only when you are sure that there is no bleeding (see Fig. 2g). This is a very critical step to check. 8. Close the peritoneum with absorbable 5-0 sutures without damaging any internal organs and also without making any cuts on the membrane (see Fig. 2h). 9. Close the skin cut with two wound clips (see Fig. 2i) and apply antiseptic cream. Turn off the isoflurane supply. 10. Follow the steps 22–27 from Subheading 3.1 above to ensure animals are recovered from anesthesia, surgery. and are being monitored daily and then weekly for any signs of discomfort, pain, and infection. 11. Monitor the animals for another 4–10 weeks before assessing their graft function. Please note that pancreas regeneration takes around 4–6 weeks (depending on the age of mice). 3.3

Insulin ELISA

Functional assessment of the transplanted graft can be performed by measuring circulating human insulin, immunostaining the kidney tissue to detect the presence of insulin/C-peptide protein as well as performing real-time PCR to detect various human-specific endocrine and islet hormone transcripts in the kidney tissue. We describe here measurement of circulating human insulin using ELISA. 1. Fast the animals overnight before initiating functional assessment. 2. Inject 2 g/kg body weight glucose intraperitoneally. 3. Collect blood sample using cardiac puncture method 30 min after the glucose administration and just prior to euthanasia. 4. Place the blood tube on ice once it clots (leave blood at room temperature for 15 min to allow blood to clot) and separate the serum after centrifugation of clotted blood at 2000
g for 15 min. If plasma is desired, proceed to centrifuge the blood tube immediately using appropriate (EDTA/Heparin) collection tubes. 5. Aspirate the upper straw colored serum in a fresh centrifuge tube and store at 80  C until further use. 6. Bring all test samples and reagents of the ELISA kit to room temperature before starting the insulin measurement assay. 7. Prepare reagents including wash buffer 1
by diluting the concentrate with MilliQ water. 8. Dilute the enzyme conjugate to 1
. 9. Prepare a plate map to include controls, calibrators, and samples in at least duplicates. 10. Dilute the samples if necessary (see Note 12).

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11. Add 25 μL each of above into appropriate wells and add 100 μL of enzyme conjugate 1
to each well. 12. Incubate at room temperature on plate shaker (slow speed) for 1 h. 13. Wash six times with 700 μL of Wash buffer 1
per well, invert and tap the plate firmly against the absorbent paper after each wash (see Note 13). 14. Add 200 μL of substrate TMB (3,30 ,5,50 -Tetramethylbenzidine) into each well and incubate the plate on lab bench for 15 min at room temperature (see Note 14). 15. After incubation, add 50 μL stop solution to each well. Place the plate on the shaker to ensure mixing. 16. Read the plate on spectrophotometer within 30 min at optical density 450 nm. 17. Prepare a calibrator curve as per manufacturer’s instruction for each assay run and extrapolate sample concentrations using GraphPad Prism or similar softwares.

4

Notes 1. Different anesthetic agents are used based on the practices of one’s institute/University. We use isoflurane, but ketamine and xylazine are also used routinely in other places where vaporizers are not available/used. 2. Other animal models of immunodeficiency can also be used as per the availability, experimental need, and institutional ethics committee recommendations. 3. Fasting reduces the peristaltic bowel movements and minimizes discomfort to the animal post-surgery. We find that animals that are not fasted for at least 6 h have dilated gut that could adversely affect the success of these surgeries. 4. Assess proper anesthetization by observing gradual loss of voluntary movement, steady breathing, and muscle relaxation. One can test the loss of reflexes by toe pinching or applying a cotton wisp to the eyes to check blinking reflex. 5. If the surgical procedure is time consuming, then apply ophthalmic ointment to prevent dryness of the eyes while animals are under anesthesia. 6. Make sure that after shaving the skin, you can locate the laterally placed, reddish brown colored kidney. 7. Make sure that the cut on intraperitoneum is just enough to pull the kidney out so that it can then rest on the peritoneal membrane without prolapsing back in the cavity.

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8. While making the pocket under kidney capsule, care should be taken as to not puncture the thin capsule membrane. 9. Make sure that the mouse has completely recovered from anesthesia and is freely moving in the cage. 10. Humane endpoints will be rapid weight loss >20%, severe depression, surgical complications unresponsive to medical treatment. 11. In mice, pancreas extends from tip of the spleen to all throughout the length of mesentery. Splenic pancreas constitutes around 80–90% of total pancreas whereas mesenteric pancreas is around 10%. When spleen is pulled out, the splenic pancreas also gets pulled out with it as shown in Fig. 2d. 12. Serum and plasma samples are normally not required to be diluted; however, samples with concentration above highest calibrator, e.g., Calibrator 5, should be diluted in lowest calibrator, e.g., Calibrator 0. 13. One can use an automatic plate washer with overflow wash function, if available. In this case, invert and tap the plate firmly against the absorbent paper after the final wash and do not include soak step in wash procedure. 14. Be careful not to contaminate the substrate TMB with enzyme conjugate solution.

Acknowledgments The support provided to MVJ through the Australian Diabetes Society (ADS) Skip Martin fellowship and the JDRF International post-doctoral fellowship, WKMW through the University of Sydney post-graduate and JDRF post-graduate top-up awards and to AAH through the JDRF Australia Career Development Award as well as the visiting professorship through the Danish Diabetes Academy is highly acknowledged. The methodologies presented herein were trained and utilized by research staff and fellows in the Hardikar Lab; especially in projects involving Dr. Vishal Parekh and Dr. Smruti Phadnis, which are acknowleged [13, 14]. References 1. Collaboration NCDRF (2016) Worldwide trends in diabetes since 1980: a pooled analysis of 751 population-based studies with 4.4 million participants. Lancet 387 (10027):1513–1530. https://doi.org/10. 1016/S0140-6736(16)00618-8 2. Cho NH, Shaw JE, Karuranga S, Huang Y, da Rocha Fernandes JD, Ohlrogge AW, Malanda

B (2018) IDF Diabetes Atlas: Global estimates of diabetes prevalence for 2017 and projections for 2045. Diabetes Res Clin Pract 138:271–281. https://doi.org/10.1016/j.dia bres.2018.02.023 3. Bjork S (2001) The cost of diabetes and diabetes care. Diabetes Res Clin Pract 54(Suppl 1): S13–S18

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4. Willey VJ, Kong S, Wu B, Raval A, Hobbs T, Windsheimer A, Deshpande G, Tunceli O, Sakurada B, Bouchard JR (2018) Estimating the real-world cost of diabetes mellitus in the United States during an 8-year period using 2 cost methodologies. Am Health Drug Benefits 11(6):310–318 5. Atkinson MA, Eisenbarth GS, Michels AW (2014) Type 1 diabetes. Lancet 383 (9911):69–82. https://doi.org/10.1016/ S0140-6736(13)60591-7 6. Shapiro AM, Pokrywczynska M, Ricordi C (2017) Clinical pancreatic islet transplantation. Nat Rev Endocrinol 13(5):268–277. https:// doi.org/10.1038/nrendo.2016.178 7. Shapiro J, Ryan E, Warnock GL, Kneteman NM, Lakey J, Korbutt GS, Rajotte RV (2001) Could fewer islet cells be transplanted in type 1 diabetes? Insulin independence should be dominant force in islet transplantation. BMJ 322(7290):861 8. Harlan DM, Rother KI (2004) Islet transplantation as a treatment for diabetes. N Engl J Med 350(20):2104.; author reply 2104. https://doi.org/10.1056/ NEJM200405133502022 9. Matsumoto S, Okitsu T, Iwanaga Y, Noguchi H, Nagata H, Yonekawa Y, Yamada Y, Fukuda K, Tsukiyama K, Suzuki H, Kawasaki Y, Shimodaira M, Matsuoka K, Shibata T, Kasai Y, Maekawa T, Shapiro J, Tanaka K (2005) Insulin independence after living-donor distal pancreatectomy and islet allotransplantation. Lancet 365 (9471):1642–1644. https://doi.org/10. 1016/s0140-6736(05)66383-0 10. Dy EC, Harlan DM, Rother KI (2006) Assessment of islet function following islet and pancreas transplantation. Curr Diab Rep 6 (4):316–322 11. Path G, Perakakis N, Mantzoros CS, Seufert J (2019) Stem cells in the treatment of diabetes mellitus - Focus on mesenchymal stem cells. Metabolism 90:1–15. https://doi.org/10. 1016/j.metabol.2018.10.005 12. McKimpson WM, Accili D (2019) Reprogramming cells to make insulin. J Endocr Soc 3 (6):1214–1226. https://doi.org/10.1210/js. 2019-00040 13. Parekh VS, Joglekar MV, Hardikar AA (2009) Differentiation of human umbilical cord

blood-derived mononuclear cells to endocrine pancreatic lineage. Differentiation 78 (4):232–240. https://doi.org/10.1016/j. diff.2009.07.004S0301-4681(09)00085-1 14. Phadnis SM, Joglekar MV, Dalvi MP, Muthyala S, Nair PD, Ghaskadbi SM, Bhonde RR, Hardikar AA (2011) Human bone marrow-derived mesenchymal cells differentiate and mature into endocrine pancreatic lineage in vivo. Cytotherapy 13(3):279–293. https://doi.org/10.3109/14653249.2010. 523108 15. Zhou Q, Melton DA (2018) Pancreas regeneration. Nature 557(7705):351–358. https:// doi.org/10.1038/s41586-018-0088-0 16. Hardikar AA (2004) Generating new pancreas from old. Trends Endocrinol Metab 15 (5):198–203. https://doi.org/10.1016/j. tem.2004.05.001 17. Hardikar AA, Karandikar MS, Bhonde RR (1999) Effect of partial pancreatectomy on diabetic status in BALB/c mice. J Endocrinol 162 (2):189–195. https://doi.org/10.1677/joe. 0.1620189 18. Joglekar MV, Parekh VS, Mehta S, Bhonde RR, Hardikar AA (2007) MicroRNA profiling of developing and regenerating pancreas reveal post-transcriptional regulation of neurogenin3. Dev Biol 311(2):603–612. https:// doi.org/10.1016/j.ydbio.2007.09.008 19. Joglekar MV, Parekh VS, Hardikar AA (2007) New pancreas from old: microregulators of pancreas regeneration. Trends Endocrinol Metab 18(10):393–400. https://doi.org/10. 1016/j.tem.2007.10.001 20. Stokes RA, Cheng K, Lalwani A, Swarbrick MM, Thomas HE, Loudovaris T, Kay TW, Hawthorne WJ, O’Connell PJ, Gunton JE (2017) Transplantation sites for human and murine islets. Diabetologia 60 (10):1961–1971. https://doi.org/10.1007/ s00125-017-4362-8 21. van der Windt DJ, Echeverri GJ, Ijzermans JN, Cooper DK (2008) The choice of anatomical site for islet transplantation. Cell Transplant 17 (9):1005–1014 22. Vaithilingam V, Bal S, Tuch BE (2017) Encapsulated Islet transplantation: where do we stand? Rev Diabet Stud 14(1):51–78. https://doi.org/10.1900/RDS.2017.14.51

Chapter 7 Color-Coded Imaging of Cancer and Stromal-Cell Interaction in the Pancreatic-Cancer Tumor Microenvironment (TME) Atsushi Suetsugu and Robert M. Hoffman Abstract The tumor microenvironment (TME) contains stromal cells in a complex interaction with cancer cells. This relationship has become better understood with the use of fluorescent proteins for in vivo imaging, originally developed by our laboratories. Spectrally-distinct fluorescent proteins can be used for colorcoded imaging of the complex interaction of the tumor microenvironment in the living state using cancer cells expressing a fluorescent protein of one color and host mice expressing another-color fluorescent protein. Cancer cells engineered in vitro to express a fluorescent protein were orthotopically implanted into transgenic mice expressing a fluorescent protein of a different color. Confocal microscopy was then used for color-coded imaging of the TME. Color-coded imaging of the TME has enabled us to discover that stromal cells are necessary for metastasis. Patient-derived orthotopic xenograft (PDOX) tumors were labeled by first passaging them orthotopically through transgenic nude mice expressing either green, red, or cyan fluorescent protein in order to label the stromal cells of the tumor (Yang et al., Cancer Res 64:8651–8656, 2004; Yang et al. J Cell Biochem 106: 279–284, 2009). The colored stromal cells become stably associated with the PDOX tumors through multiple passages in transgenic colored nude mice or non-colored nude mice. The fluorescent protein-expressing stromal cells included cancer-associated fibroblasts and tumor-associated macrophages. Color-coded imaging enabled the visualization of apparent fusion of cancer and stromal cells. Color-coded imaging is a powerful tool visualizing the interaction of cancer and stromal cells during cancer progression and treatment. Key words Tumor microenvironment, Niche, Cancer-associated fibroblasts, GFP, RFP, CFP, Colorcoded imaging

1

Introduction In vivo imaging with fluorescent proteins was pioneered by our laboratories and has been particularly useful for studying tumor growth, invasion, cancer-cell trafficking, metastasis, angiogenesis, and other aspects of tumor progression [1–5]. Multicolored proteins have allowed the color coding of cancer cells growing in vivo therapy distinguishing between highly and poorly metastatic cells, cancer stem and non-stem cells, and gene transfer between cancer cells [3, 4, 6–13].

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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The tumor microenvironment (TME) contains stromal cells in a complex interaction with cancer cells. This relationship has become better understood with the use of different-color fluorescent proteins for in vivo imaging, originally developed by our laboratories. Spectrally-distinct fluorescent proteins can be used for color-coded imaging of the complex interaction of the tumor microenvironment in the living state using cancer cells expressing a fluorescent protein of one color and host mice expressing anothercolor fluorescent protein. Cancer cells engineered in vitro to express a fluorescent protein were orthotopically implanted into transgenic mice expressing a fluorescent protein of a different color [14, 15]. Confocal microscopy was then used for color-coded imaging of the TME. Color-coded imaging of the TME has enabled us to discover that stromal cells are necessary for metastasis and that therapy can selectively target stromal cells. Color-coded imaging also demonstrated that cancer and stromal cells can fuse and become highly malignant. This chapter focuses on color-coded imaging of cancer and stromal cells in the tumor microenvironment (TME), associated with tumor growth and progression. Colorcoded imaging of the TME provides visual targets for immunooncology and is very timely and has the potential to unlock the secrets of the TME that influence cancer growth and progression.

2 2.1

Materials Animals

1. Transgenic C57B6-GFP mice (Osaka University, Osaka). 2. NOD/SCID mice were purchased from Charles River (Wilmington, MA, USA). 3. Green fluorescent protein (GFP)-expressing athymic nu/nu nude mice (AntiCancer Inc. San Diego, CA). 4. Red fluorescent protein (RFP)-expressing athymic nu/nu nude mice (AntiCancer Inc. San Diego, CA). 5. Cyan fluorescent protein (CFP)-expressing athymic nu/nu nude mice (AntiCancer Inc. San Diego, CA).

2.2 Animal Care and Treatment

1. Animal diets were obtained from Harlan Teklad (Madison, WI, USA). 2. High efficacy particulate arrestance (HEPA)-filtered rack. 3. Ampicillin (5.0%, w/v; Sigma, St. Louis, MO, USA). 4. 8–0 nylon surgical sutures (Ethilon; Ethicon Inc., NJ, USA). 5. 6–0 nylon surgical sutures (Ethilon; Ethicon Inc., NJ, USA). 6. Ketamine mixture (0.02 mL solution of 20 mg/kg ketamine, 15.2 mg/kg xylazine, and 0.48 mg/kg acepromazine maleate).

Color-Coded Imaging of the TME

2.3

Cells and Culture

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1. Cancer cell lines were obtained from the American Type Culture Collection (Manassas, VA). 2. Colon 26 labeled with RFP (AntiCancer inc. San Diego, CA). 3. RPMI 1640 medium (Gibco Life Technologies, Grand Island, NY). 4. FBS (Gemini Biological Products, Calabasas, CA). 5. 10% heat-inactivated FBS (Gemini Biological Products, Calabasas, CA).

2.4

Imaging System

1. The OV100 variable magnification Small Animal Imaging System (Olympus Corp., Tokyo). 2. FV1000 confocal fluorescence microscope (Olympus Corp., Tokyo). 3. MT-20 light Germany).

source

(Olympus

Biosystems,

Planegg,

4. DP70 CCD camera (Olympus). 5. Semiconductor laser at 473 nm for GFP excitation. 6. Tunable Mai Tai HP femtosecond laser emitting at 700–1020 nm (Newport Spectra-Physics, Irvine, CA).

3 3.1

Methods Animal Care

1. Breed and maintain transgenic GFP and RFP nude mice and non-transgenic nude mice in a HEPA-filtered environment. 2. Sterilize cages, food, and bedding by autoclaving [14]. 3. Add Ampicillin to the autoclaved drinking water. 4. Perform surgical procedures and imaging by anesthetizing animal by intramuscular injection of 0.02 mL of a solution of 50% ketamine, 38% xylazine, and 12% acepromazine maleate. 5. All animal studies were conducted in accordance with the principles of and procedures outlined in the NIH Guide for Care and Use of Laboratory Animals [16].

3.2

Cell Culture

1. Maintain human or mouse cancer cell lines in RPMI 1640 medium supplemented with 10% FBS, 2 mmol/L glutamine. 2. Culture the cell line at 37
C in a 5% CO2 incubator.

3.3 Establishment of a PDOX Model of Patient Tumors

1. Obtain tumor tissue from patients at surgery with informed consent. 2. Cut tumor tissue into 3 mm3 fragments and transplant subcutaneously into nude or SCID mice [17, 18].

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3.4 Orthotopic Tumor Transplantations in Transgenic Fluorescent ProteinExpressing Nude Mice

1. Transplant intact pancreatic-cancer-tissue fragments orthotopically into four 6-week-old transgenic GFP or RFP nude mice using surgical orthotopic implantation (SOI). 2. Make a small 6–10 mm transverse incision in the left flank of the mouse through the skin and peritoneum. 3. Expose the tail of the pancreas through this incision and suture a single 3 mm [19, 20] tumor fragment from harvested subcutaneous tumors to the tail of the pancreas using 8–0 nylon surgical sutures. 4. Close the incision after completion of surgery in one layer using 6–0 nylon surgical sutures [19–21].

3.5 Fluorescence Imaging

1. Perform whole-tumor macroimaging using the OV100 Small Animal Imaging System [23]. 2. Obtain bright-field and GFP- and RFP-fluorescence images. The OV100 contains an MT-20 light source (Olympus Biosystems, Planegg, Germany) and DP70 CCD camera (Olympus). The OV100 was used for subcellular imaging in live mice. The optics of the OV100 fluorescence imaging system have been specially developed for macroimaging as well as microimaging with high light-gathering capacity. The instrument incorporates a unique combination of high numerical aperture and long working distance.

3.6 Confocal Microscopy

1. Use confocal two-photon microscopy for two- (2D, x, y) and three-dimensional (3D, x, y, z) high-resolution imaging. 2. Use a continuous wave (cw) semiconductor laser at 473 nm for GFP excitation and a tunable Mai Tai HP femtosecond laser emitting at 700–1020 nm for deep tissue imaging of autofluorescence and GFP. 3. Obtain fluorescence images using the 20/0.50 UPlan FLN and 40/1.3 Oil Olympus UPLAN FLN objectives [24].

3.7 Histological Analysis

1. Perform sectioning of the primary tumor, liver metastases, and disseminated peritoneal metastases at a thickness of 8-mm using frozen sections for fluorescence imaging and paraffinembedded tissue for staining using hematoxylin and eosin for microscopic analysis [25].

3.8

1. The GFP stromal cells from the GFP host mouse migrated into the orthotopic pancreatic tumor causing the tumors to fluoresce bright green [25]. Both GFP cancer-associated fibroblasts (CAFs) and tumor-associated microphages (TAMs) were observed in the primary tumor. Histological examination at 110 days of tumor growth revealed pancreatic tubular adenocarcinoma [25] (see Fig.1).

Results

3.8.1 GFP Host Stromal cells Infiltrate a Pancreatic Cancer PDOX

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H.E. stain

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Fig. 1 (a) Image of liver metastasis. Yellow arrows indicate tumor-associated macrophages (TAMs). White arrows indicate cancer-associated fibroblasts. Image was obtained with an Olympus FV1000 confocal microscope (Bar ¼ 20 μm). (b) Liver metastasis stained with hematoxylin and eosin. Blue arrows indicate pancreatic tubular adenocarcinoma. Yellow arrows indicate stromal cells. Red arrows indicate hepatocytes (Bar ¼ 100 μm) 3.8.2 GFP Host Stromal Cells Infiltrate Peritoneal Disseminated Metastases of Pancreatic Cancer PDOX

1. The GFP stromal cells from the GFP host mouse formed a capsule around the disseminated peritoneal metastases. Both GFP-labeled CAFs and TAMs were observed in the disseminated peritoneal metastases. Histological examination at 110 days of tumor growth demonstrated pancreatic tubular adenocarcinoma, similar to the primary tumor [25] (see Fig. 2) (see Note1).

3.8.3 RFP Host Stromal cells Infiltrate Pancreatic Cancer PDOX

1. The RFP stromal cells from the RFP host mice formed a capsule around the tumor and infiltrated into the central part of the tumor as well. RFP-expressing TAMs could be visualized in the tumor [16] (see Fig. 3) (see Note 2).

3.8.4 GFP Host Stromal Cells Infiltrate Pancreatic PDOX Labeled with RFP Stroma to Form a TwoColor Stroma Model

1. Tumors were grown in RFP transgenic nude mice and orthotopically in GFP nude mice, after which the human pancreatic cancer PDOX contained both RFP and GFP stromal cells. The RFP stromal cells still persisted after passage to GFP transgenic mice. Under confocal microscopy with the FV1000, RFP and GFP stromal cells were clearly visualized in the tumor including GFP and RFP CAFs and TAMs in the central part of the tumor [16] (see Fig.4) (see Note 3).

3.8.5 CFP Host Stromal Cells Infiltrate Pancreatic Cancer PDOX Previously Grown in RFP and GFP Transgenic Mice to Form a Three-Color Stroma Model

1. Pancreatic cancer previously grown in RFP and GFP transgenic mice was orthotopically implanted in 6-week-old nude CFP mice. The tumors were excised and observed with the FV1000 confocal microscope. RFP-, GFP-, and CFP-expressing stromal cells were observed in the human pancreatic cancer patient tumor. The RFP stroma persisted after two passages and GFP

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Fig. 2 (a) Color-coded image of disseminated peritoneal metastasis. Red arrow indicates disseminated peritoneal metastasis with green fluorescent protein (GFP) stroma. Bar ¼ 10 mm. (b) Image of disseminated peritoneal metastasis. Yellow arrows indicate tumor-associated macrophages (TAMs). White arrows indicate cancer-associated fibroblasts (CAFs). Image was obtained with an Olympus FV1000 confocal laser microscope. Bar ¼ 20 μm. (c) Disseminated peritoneal metastasis stained with H&E. Blue arrows indicate pancreatic tubular adenocarcinoma. Bar ¼ 100 μm

stroma persisted after one passage in CFP mice. RFP TAMs and CAFs and GFP blood vessels still persisted in the human pancreatic cancer patient tumor after one and two passages, respectively [26] (see Fig. 5) (see Note 4). 3.8.6 Noninvasive Imaging of Pancreatic Cancer PDOX with Labeled Stromal Cells

1. Noninvasive imaging at days 21, 30, and 74 demonstrated extensive orthotopic growth of the pancreatic cancer PDOX, labeled with RFP and GFP stroma, on the nude mouse pancreas [27] (see Fig. 6) (see Note 5).

3.8.7 Color-Coded Imaging of Stromal-Cell and Cancer-Cell Response to Therapy

1. An inhibitor of transforming growth factor-β (TGF-β) was shown to target stroma in an orthotopic mouse model of pancreatic cancer with a color-coded TME. The BxPC-3 human pancreatic adenocarcinoma cell line expressing GFP was used in an orthotopic model in transgenic nude mice ubiquitously expressing RFP. The area of RFP fluorescence from the stromal cells relative to the area of GFP fluorescence of the cancer cells was significantly reduced by a TGF-β

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Fig. 3 (a) Patient-derived orthotopic xenograft (PDOX) model of human pancreatic cancer patient tumor transplanted to RFP transgenic nude mouse. Yellow arrow indicates host RFP nude mouse pancreas. Blue arrow indicates tumor with infiltrating RFP stroma cells (Bar ¼ 10 mm). Image obtained with the Olympus OV100. (b) Human pancreatic tumor excised from RFP nude mouse with RFP stroma cells. The image is of a cross-section of the tumor. Blue arrow indicates RFP stromal cells (Bar ¼ 10 mm). Image obtained with the Olympus OV100. (c) Visualization of RFP tumor-associated macrophages (TAMs) in the human pancreatic cancer patient tumor (F2). High-magnification image taken with the Olympus FV1000 confocal microscope. Yellow arrows indicate RFP TAMs (Bar ¼ 50 μm)

inhibitor indicating targeting of the stroma by the inhibitor. Color-coded imaging in an orthotopic pancreatic cancer cell line mouse model thus readily enabled detection of the selective targeting of stromal cells by a TGF-β inhibitor [22] (see Fig. 7) (see Notes 6 and 7). 3.8.8 Fusion of Cancer and Stromal Cells

1. Red fluorescent protein (RFP-expressing) mouse colon cancer-26 cells were initially injected subcutaneously in green fluorescent protein (GFP) nude mice. The resulting subcutaneous tumors were harvested and cultured. The cultured

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subcutaneous tumors contained RFP colon cancer cells, GFP stromal cells, and recombinant cancer-stromal cells expressing yellow fluorescence. (Fig. 8) (see Note 8) After 14 days culture, the cells were injected into the spleen. After splenic injection, colon cancer 26 metastases were observed in the liver, ascites, and bone marrow. Using the Olympus FV1000 confocal microscope, the cells cultured from tumors and metastasis in each site were visualized. RFP colon cancer cells, GFP stromal cells derived from host GFP nude mice, and recombinant yellow fluorescent cells were observed in stromal cells were observed. (Fig. 8) Color-coded imaging demonstrated the dynamics of colon cancer and stromal cells at different metastatic sites including the formation of recombinant cancerstromal cells [28].

Color-Coded Imaging of the TME

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Fig. 5 Pancreatic cancer patient-derived orthotopic xenograft (PDOX) growing in a cyan fluorescent protein (CFP)-expressing host. RFP cancer-associated fibroblasts (CAFs) (yellow arrow) and GFP TAMs (green arrows) present in the tumor. White arrow indicates CFP CAFs. (Bar ¼ 30 μm). Image was obtained with an Olympus FV1000 confocal microscope

Fig. 6 Noninvasive imaging of fluorescent tumor from a patient with pancreatic cancer growing in a PDOX model in non-transgenic nude mice. The PDOX was previously grown in transgenic RFP and transgenic GFP nude mice where the PDOX acquired RFP and GFP stroma, respectively. Whole-body noninvasive imaging with the OV100 of human pancreatic cancer PDOX in transgenic nude mice. Mice were noninvasively imaged at day-21 (left panel), day-30 (middle panel), and day-74 (right panel)

2. High-resolution color-coded fluorescence imaging is a powerful technique for differentially labeling cancer and stromal cells in the tumor microenvironment (TME) with spectrally distinct fluorescent proteins, allowing observations of the behavior of each cell type and their interaction within tumors, thereby better defining the specific role of cancer and stromal cells in tumor progression, and how to target each cell type with novel therapeutics, including immun-oncology therapeutics which target both cancer and stromal cells, particularly T-lymphocytes [29].

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Fig. 7 Color-coded imaging of resected tumors and frozen sections from TGF-β inhibitors-treated and control tumors. Bright-field and fluorescence images of whole tumors were obtained with the OV100 Small Animal Imaging System. Representative images of frozen tumor sections in both groups imaged with the FV-1000 confocal microscope

4

Notes 1. After dissecting the orthotopic transplanted mouse, immediately observe tissue with OV100 so as not to attenuate the fluorescent protein. 2. Before transplantion into NOD-SCID mice, make sure to remove connective tissue to make it only the pancreatic-cancer part of the patient pancreatic cancer specimen. 3. When observing tissues with the FV1000, it is important to fix the mouse so that it does not move.

Color-Coded Imaging of the TME

RFP filter

GFP filter

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Merge

day14

a

20μm

RFP filter

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GFP filter

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b Fig. 8 (a) High-magnification images of cells cultured from harvested mouse colon-26 RFP tumors grown GFP mice on day 14. White arrows indicate spindle-shaped yellow fluorescent recombinant cells expressing yellow fluorescence (Bar ¼ 20 μm). (b) High-magnification images of cells cultured from ascites. Many colon-26-RFP cells and a few GFP stromal cells were observed (Bar ¼ 100 μm). White arrows indicate recombinant yellow fluorescent cancer-stromal cells (Bar ¼ 30 μm). Images were captured with the Olympus FV1000 confocal microscope

4. Set the gain so that the wavelengths do not overlap (especially GFP and CFP). 5. When observing the mouse with OV100, fix the mouse on the table. 6. When observing the structure with the FV1000, the structure is kept horizontal, and a glass plate is brought into close contact with the structure for observation. 7. When observing a deep part of tissue, use oil with 20/0.50 UPLAN FLN and 40/1.3 Oil Olympus UPLAN FLN objectives 6. 8. Set the gain well when checking cell fusion.

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References 1. Chishima T, Miyagi Y, Wang X, Yamaoka H, Shimada H, Moossa AR, Hoffman RM (1997) Cancer invasion and micrometastasis visualized in live tissue by green fluorescent protein expression. Cancer Res 57:2042–2047 2. Hoffman RM, Yang M (2006) Subcellular imaging in the live mouse. Nat Protocol 1:775–782 3. Hoffman RM, Yang M (2006) Color-coded fluorescence imaging of tumor host interactions. Nat Protoco1 1(12):928–935 4. Hoffman RM (2005) The multiple uses of fluorescent proteins to visualize cancer in vivo. Nat Rev Cancer 5:796–806 5. Hoffman RM (2012) In: Walker JM (ed) In vivo cellular imaging using fluorescent proteins: methods and protocols. methods in molecular biology, vol 872. Humana Press, New York, NY 6. Yamamoto N, Yang M, Jiang P, Xu M, Tsuchiya H, Tomita K, Moossa AR, Hoffman RM (2003) Determination of clonality of metastasis by cell-specific color-coded fluorescent-protein imaging. Cancer Res 63:7785–7790 7. Yang M, Li L, Jiang P, Moossa AR, Penman S, Hoffman RM (2003) Dual-color fluorescence imaging distinguishes tumor cells from induced host angiogenic vessels and stromal cells. Proc Natl Acad Sci U S A 100:14259–14262 8. Yamauchi K, Tome Y, Yamamoto N, Hayashi K, Kimura H, Tsuchiya H, Tomita K, Bouvet M, Hoffman RM (2012) Color-coded real-time subcellular fluorescence imaging of the interaction between cancer and host cells in live mice. Anticancer Res 32:39–43 9. Amoh Y, Hamada Y, Katsuoka K, Hoffman RM (2012) In vivo imaging of nuclearcytoplasmic deformation and partition during cancer cell death due to immune rejection. J Cell Biochem 133:465–472 10. Glinskii AB, Smith BA, Jiang P, Li X-M, Yang M, Hoffman RM, Glinsky GV (2003) Viable circulating metastatic cells produced in orthotopic but not ectopic prostate cancer models. Cancer Res 63:4239–4243 11. Tome Y, Tsuchiya H, Hayashi K, Yamauchi K, Sugimoto N, Kanaya F, Tomita K, Hoffman RM (2009) In vivo gene transfer between interacting human osteosarcoma cell lines is associated with acquisition of enhanced metastatic potential. J Cell Biochem 108:362–367 12. Suetsugu A, Osawa Y, Nagaki M, Moriwaki H, Saji S, Bouvet M, Hoffman RM (2010)

Simultaneous color-coded imaging to distinguish cancer “stem-like” and non-stem cells in the same tumor. J Cell Biochem 111:1035–1041 13. Hassanein MK, Suetsugu A, Saji S, Moriwaki H, Bouvet M, Moossa AR, Hoffman RM (2011) Stem-like and non-stem human pancreatic cancer cells distinguished by morphology and metastatic behavior. J Cell Biochem 112:3549–3554 14. Yang M, Reynoso J, Jiang P, Li L, Moossa AR, Hoffman RM (2004) Transgenic nude mouse with ubiquitous green fluorescent protein expression as a host for human tumors. Cancer Res 64:8651–8656 15. Yang M, Reynoso J, Bouvet M, Hoffman RM (2009) A transgenic red fluorescent proteinexpressing nude mouse for color-coded imaging of the tumor microenvironment. J Cell Biochem 106:279–284 16. Suetsugu A, Katz M, Fleming J, Truty M, Thomas R, Moriwaki H, Bouvet M, Saji S, Hoffman RM (2012) Multi-color palette of fluorescent proteins for imaging the tumor microenvironment of orthotopic tumorgraft mouse models of clinical pancreatic cancer specimens. J Cell Biochem 113:2290–2295 17. Kim MP, Evans DB, Wang H, Abbruzzese JL, Fleming JB, Gallick GE (2009) Generation of orthotopic and heterotopic human pancreatic cancer xenografts in immunodeficient mice. Nat Protoc 4:1670–1680 18. Kim MP, Truty MJ, Choi W, Kang Y, ChopinLally X, Gallick GE, Wang H, McConkey DJ, Hwang R, Logsdon C, Abbruzzesse J, Fleming JB (2012) Molecular profiling of direct xenograft tumors established from human pancreatic adenocarcinoma after neoadjuvant therapy. Ann Surg Oncol 19(Suppl 3):S395–S403 19. Fu X, Guadagni F, Hoffman RM (1992) A metastatic nude-mouse model of human pancreatic cancer constructed orthotopically from histologically intact patient specimens. Proc Natl Acad Sci U S A 89:5645–5649 20. Hoffman RM (1999) Orthotopic metastatic mouse models for anticancer drug discovery and evaluation: a bridge to the clinic. Investig New Drugs 17:343–359 21. Hiroshima Y, Maawy A, Zhan Y, Murakami T, Momiyama M, Mori R, Matsuyama R, Chishima T, Tanaka K, Ichikawa Y, Endo I, Hoffman RM (2015) Fluorescence-guided surgery, but not bright-light surgery, prevents local recurrence in a pancreatic cancer patientderived orthotopic xenograft (PDOX) model

Color-Coded Imaging of the TME resistant to neo- adjuvant chemotherapy (NAC). Pancreatology 15:295–301 22. Murakami T, Deng SJ, Eilber FC, Zhao M, Zhang Y, Zhang N, Singh A, Russell T, Deng S, Reynoso J, Quan C, Hiroshima Y, Matsuyama R, Chishima T, Tanaka K, Bouvet M, Chawla S, Endo I, Hoffman RM (2016) Tumor targeting Salmonella typhimurium A1-R in combination with doxorubicin eradicate soft tissue sarcoma in a patientderived orthotopic xenograft PDOX model. Oncotarget 7:12783–12790 23. Yamauchi K, Yang M, Jiang P, Xu M, Yamamoto N, Tsuchiya H, Tomita K, Moossa AR, Bouvet M, Hoffman RM (2006) Development of real-time subcellular dynamic multicolor imaging of cancer-cell trafficking in live mice with a variable-magnification wholemouse imaging system. Cancer Res 66:4208–4214 24. Uchugonova A, Duong J, Zhang N, Ko¨nig K, Hoffman RM (2011) The bulge area is the origin of nestin-expressing pluripotent stem cells of the hair follicle. J Cell Biochem 112:2046–2050 25. Suetsugu A, Katz M, Fleming J, Truty M, Thomas R, Saji S, Moriwaki H, Bouvet M, Hoffman RM (2012) Imageable fluorescent metastasis resulting in transgenic GFP mice orthotopically implanted with human-patient

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primary pancreatic cancer specimens. Anticancer Res 32:1175–1180 26. Suetsugu A, Hassanein MK, Reynoso J, Osawa Y, Nagaki M, Moriwaki H, Saji S, Bouvet M, Hoffman RM (2012) The cyan fluorescent protein nude mouse as a host for multicolor-coded imaging models of primary and metastatic tumor microenvironments. Anticancer Res 32:31–38 27. Suetsugu A, Katz M, Fleming J, Truty M, Thomas R, Saji S, Moriwaki H, Bouvet M, Hoffman RM (2012) Non-invasive luorescent-protein imaging of orthotopic pancreatic-cancer- patient tumorgraft progression in nude mice. Anticancer Res 32:3063–3068 28. Nakamura M, Suetsugu A, Hasegawa K, Satake T, Kunisada T, Shimizu M, Saji S, Moriwaki H, Hoffman RM (2018) Colorcoded imaging distinguishes cancer cells, stromal cells, and recombinant cancer-stromal cells in the tumor microenvironment during metastasis. Anticancer Res 38:4417–4423 29. Takahashi H, Sakakura K, Arisaka Y, Tokue A, Kaira K, Tada H, Higuchi T, Okamoto A, Tsushima Y, Chikamatsu K (2019) Clinical and biological significance of PD-L1 expression within the tumor microenvironment of oral squamous cell carcinoma. Anticancer Res 39:3039–3046

Chapter 8 Generating Ins2+/
/miR-133aTg Mice to Model miRNA-Driven Cardioprotection of Human Diabetic Heart Hamid R. Shahshahan, Tyler N. Kambis, Sumit Kar, Santosh K. Yadav, and Paras K. Mishra Abstract Diabetes mellitus (DM) is caused either due to insulin deficiency (T1DM) or insulin resistance (T2DM). DM increases the risk of heart failure by diabetic cardiomyopathy (DMCM), a cardiac muscle disorder that leads to a progressive decline in diastolic function, and ultimately systolic dysfunction. Mouse models of T1DM and T2DM exhibit clinical signs of DMCM. Growing evidence implicates microRNA (miRNA), an endogenous, non-coding, regulatory RNA, in the pathogenesis and signaling of DMCM. Therefore, inhibiting deleterious miRNAs and mimicking cardioprotective miRNAs could provide a potential therapeutic intervention for DMCM. miRNA-133a (miR-133a) is a highly abundant miRNA in the human heart. It is a cardioprotective miRNA, which is downregulated in the DM heart. It has anti-hypertrophic and anti-fibrotic effects. miR-133a mimic treatment after the onset of early DMCM can reverse histological and clinical signs of the disease in mice. We hypothesized that overexpression of cardiac-specific miR-133a in Ins2+/
Akita (T1DM) mice can prevent progression of DMCM. Here, we describe a method to create and validate cardiac-specific Ins2+/
/miR-133aTg mice to determine whether cardiac-specific miR-133a overexpression prevents development of DMCM. These strategies demonstrate the value of genetic modeling of human disease such as DMCM and evaluate the potential of miRNA as a therapeutic intervention. Key words Tissue-specific transgenic mice, Cardiac physiology, Diabetes models, Akita

1

Introduction Diabetic cardiomyopathy (DMCM) is caused due to detrimental effects of type 1 and type 2 diabetes mellitus (T1DM and T2DM) on cardiac muscles [1]. Hormones, cytokines, hyperglycemia, fatty acids, and adrenergic tone produced in the DM heart leads to diastolic and systolic dysfunction, and ultimately heart failure, independent of the effects of atherosclerosis, valvular disease, or hypertension [2]. Importantly, tight glycemic control of glucose does not entirely reduce the risk of heart failure in DM

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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patients [3–5]. Therefore, it is necessary to prevent DMCM after the onset of DM and treat DMCM to reduce the risk of heart failure. Several murine models exist for DM, which develop the hallmark symptoms of DMCM. For T1DM, streptozotocin (STZ)induced and Ins2+/
(Akita mice) models destroy pancreatic beta cells through chemically and genetically induced cell death, respectively [6, 7]. T2DM can be modeled by the db/db mouse, which induces hyperinsulinemia, hyperglycemia, and hyperlipidemia through mutation of leptin receptor gene [6, 8, 9]. All of these models develop many of the main characteristics of DMCM, including db/db mice develop systolic dysfunction at the age of 12–14 weeks [6, 10]. Ins2+/
Akita mouse has spontaneous mutation in the insulin gene, which causes misfolding of pro-insulin and ultimately leads to pancreatic beta cell death and hyperglycemia at the age of four week [11, 12]. These mice show diastolic dysfunction [13] and reduced ejection fraction at the age of 14-week [14]. Additional reports show molecular evidence of DMCM including increased cardiac hypertrophy, fibrosis, mitochondrial dysfunction, and inflammation [10, 13, 15–17]. Moreover, while T2DM is typically diagnosed by increased circulating levels of insulin due to insulin resistance , chronic insulin secretion by beta cells ultimately leads to beta cell death and they requiring insulin injections similar to T1DM patients [18]. Interestingly, Akita mice also develop insulin resistance [19]. Thus, Akita mice could be an appropriate model for double DM with the phenotypes of both T1DM and latestage T2DM. Non-coding, regulatory RNAs, including microRNAs, have been implicated in the development of DMCM and thus are promising diagnostic and therapeutic targets of DMCM [20]. They regulate mitochondrial function, autophagy, fibrosis, cell death, and oxidative stress in the heart [21]. miR-133a is highly abundant in the human heart and prevents cardiac remodeling [22, 23]. It is reduced in DM patients and rodents [14, 24]. We and others have shown that miR-133a can ameliorate hypertrophy, fibrosis, and systolic function in different diabetic models and cardiomyocytes subjected to high glucose concentrations [11, 17, 25]. However, it has not been determined if miR-133a mimic treatment can prevent DMCM if used at the time of onset of DM. We sought to develop a model to investigate the preventative potential of miR-133a on DMCM by crossbreeding Akita with cardiac-specific miR-133a transgenic (miR-133aTg) mice (see Fig. 1). Here, we describe the methods used to create and validate Akita/miR-133aTg mice (see Figs. 2 and 3).

miRNA Cardioprotection of Diabetic Hearts F0

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Fig. 1 Schematic presentation of crossbreeding between Akita and miR-133aTg mice. The F1 generation includes WT, Akita, miR-133aTg, and the novel Akita/miR-133aTg mice

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Fig. 2 Genotyping WT, Akita, miR-133aTg, and Akita/miR-133aTg mice. (a) Agarose gel electrophoresis of PCR products for Insulin 2 gene after restriction digestion, showing a single band for the WT gene and two bands for the heterozygous insulin gene. (b) Agarose gel electrophoresis of PCR products using miR-133aTg primers shows one band for miR-133aTg gene and no band for WT gene

2

Materials 1. Mice: Akita (Ins2+/
), stock number 003548, The Jackson Laboratory. miR-133aTg mice were gifted by Dr. Scot Matkovich, Washington University at St. Louis, USA. Both strains have C57BL/6 J background. 2. Tissue collection: Ear puncher, forceps, PCR tubes. 3. DNA extraction: Sodium hydroxide, EDTA, Tris Hydrochloride (Thermo Fisher Scientific), Thermal cycler (Bio-Rad Laboratories), Microvolume spectrophotometer (Thermo Fisher Scientific).

Hamid R. Shahshahan et al.

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Fig. 3 Validation of Akita/miR-133aTg phenotypes. (a) Relative gene expression of miR-133a normalized to U6 endogenous control. RNA was obtained from the left ventricle of WT, Akita, Akita/miR-133aTg, and miR-133aTg mice. (b) Blood glucose levels of WT, Akita, Akita/miR-133aTg, and miR-133aTg mice. Data are presented as mean  SEM. *, P < 0.05; ** P < 0.01; *** P < 0.001. N ¼ 5–7 per group. Each point represents one mouse. One-way analysis of variance (ANOVA) followed by Tukey’s test was used for statistical significance

4. PCR: Maxima hotstart green PCR master mix (Thermo Fisher Scientific), Nuclease-free water, Primers for Ins2 and miR-133aTg, Thermal cycler (Bio-Rad Laboratories). 5. Restriction digest: Fnu4HI and buffer G (Thermo Fisher Scientific), Nuclease-free water, Incubator. 6. Electrophoresis: Agarose (Sigma-Aldrich); 10 TBE buffer; Ethidium bromide; DNA ladder (Thermo Fisher Scientific), Gel electrophoresis system (Thermo Fisher Scientific), Gel imaging system (Bio-Rad Laboratories). 7. Glucose measurement: Glucometer (Roche), Glucose strips, Syringe needle, Ethanol.

3

Methods

3.1 Crossbreeding Ins2+/
and miR-133aTg Mice

Insulin 2 gene in mouse is orthologous to human insulin gene. In Akita mice, a point mutation in this gene leads to lower insulin production and T1DM [12]. Insulin 2 homozygous mutants do not survive very long. Therefore, Insulin 2 heterozygous mice are used. miR-133aTg mice have an extra miR-133a gene cloned into the α-myosin heavy-chain promoter, which expresses this gene only in the heart [26]. After confirmation of mouse strains by genotyping, a female miR-133aTg was crossbred to a male Akita (Ins2+/
) mouse in F0 generation. In F1 generation, four types of offspring were found:

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wild type, Akita, miR-133aTg, and Akita/miR-133aTg. These mice were validated by genotyping; blood glucose levels, and cardiac levels of miR-133a. 3.2

Genotyping

3.2.1 DNA Extraction

1. Any DNA isolation kit or method that yields purified DNA can be used, but the following protocol is generally faster and more cost-effective, especially for a large number of mice [27]. The resulting DNA has high quality for the polymerase chain reaction (PCR). 2. Collect a small piece of tissue from the ear, which was cut by ear puncher, in a PCR tube. Any other tissue also can be used for DNA extraction (see Note 1). 3. Add 75 μL of lysis reagent (25 mM NaOH/0.2 mM EDTA) to the tube (see Note 2). 4. Incubate the tubes in a thermal cycler at 98  C for an hour. 5. After the one hour, hold samples at 4  C until the next step. 6. Add 75 μL of neutralizing reagent (40 mM Tris–HCl) (pH 5.5). 7. Centrifuge the tubes at 1972  g for 3 min. 8. Collect the supernatant for PCR and measure the DNA concentration with a microvolume spectrophotometer (see Note 3).

3.2.2 Polymerase Chain Reaction (PCR)

1. The Akita genotyping protocol explained here (see Note 4) requires two strands of primers: forward, 5´-TGC TGA TGC CCT GGC CTG CT-30 ; and reverse, 50 -TGG TCC CAC ATA TGC ACA TG-30 . 2. The PCR protocol for Insulin 2 is: 94  C, 3 min; 94  C, 20 s; 64  C, 30 s; decrease temperature
0.5  C per cycle; 72  C, 35 s; repeat steps 2–4 for 12 cycles, 94  C, 20 s; 58  C, 30 s; 72  C, 35 s; repeat steps 5–7 for 25 cycles; 72  C, 2 min; 4  C, hold. 3. The two strands of primers for miR-133aTg genotyping are: forward, 50 -GAA GCC TAG CCC ACA CCA GAA ATG-30 ; and reverse, 50 -TGA GTA AGT TGA GAT GTA ATT CAT GCT CA-30 . 4. The PCR protocol for miR-133aTg is: 94  C, 5 min; 94  C, 1 min; 53  C, 1 min; 72  C, 1 min, repeat steps 2–4, 30 times; 72  C, 5 min; 4  C, hold.

3.2.3 Restriction Digest

1. Only for Akita genotyping, a restriction digestion step on PCR products is necessary before electrophoresis. 2. For a 10 μL reaction, mix 4 μL PCR product with 0.6 μL Fnu4HI (SatI), 1 μL 10 buffer G (provided by the enzyme), and nuclease-free water to final volume (see Note 5).

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3. Mix gently and spin down for a few seconds. 4. Incubate at 37  C for at least 3 h (see Note 6). 5. If gel electrophoresis is not performed immediately, samples should be stored at
20  C. 3.2.4 Agarose Gel Electrophoresis

1. Separate the PCR products by electrophoresis using a 2% agarose gel and ethidium bromide (0.003%) and visualize the bands under UV light. 2. For a 100 mL 2% agarose gel, dissolve 2 g of agarose in 100 mL 1 TBE buffer by heating the mixture in a microwave oven up to boiling point. 3. Stir the solution occasionally and make sure agarose is completely dissolved. 4. Allow the solution to cool down to about 60  C and then add 3 μL ethidium bromide and mix thoroughly (see Note 7). 5. Pour the solution in the gel tray, insert the comb, and let it solidify for at least 30 min. 6. When the gel is ready, add 1 TBE buffer to the chamber and load the samples and a DNA ladder into the wells. 7. Run the gel at 90 V for 1.5 h or until the samples are at a sufficient resolving distance.

3.2.5 Analysis of Gel-Band

1. For the Insulin 2 gene primer, wild type mice will have one band at 140 base pairs (see Note 8). 2. The mutant band shows up at 280 bp; therefore, Akita mouse, which is heterozygous, has two bands at 140 and 280 bp. 3. The miR-133aTg band shows up at 670 bp. 4. Wild type mice do not show any bands for miR-133a transgene primer (see Fig. 2).

3.3 Blood Glucose Measurement

1. To minimize blood glucose level variation (see Note 9), fast the mice in the morning and measure glucose level after 6–8 h at the afternoon (see Note 10). 2. Clean the tail with 70% ethanol and give it a few seconds to dry. 3. Using a clean needle, prick the tail superficially at somewhere close to the tip. 4. Gently squeeze the surrounding area to get a few microliters of blood. 5. Add the blood to the tip of a glucose strip inserted in a glucometer and wait for a few seconds to read the result. 6. Press the bleeding spot with a clean paper/gauze for about 10 sec to stop bleeding before returning the mouse to the animal cage.

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7. Generally, Akita male mice have higher blood glucose levels than females. In wild type mice, levels can vary from nearly 100–200 mg/dL. In adult Akita males, glucose levels can easily reach up to 500 mg/dL. Glucose levels in Female Akita may vary from normal range to over 300 mg/dL. 3.4 Real-Time PCR Measurement of miR-133a

1. Before isolating RNA, make sure the work environment is nuclease-free by cleaning with RNase inhibitor. 2. Isolate RNAs using a kit designed for small RNAs. The recommended kit is the mirVana miRNA isolation kit. The optional miRNA enrichment step is not necessary. 3. Obtain RNA concentrations by Nanodrop and make sure that a high quality of RNA (260/280 > 1.8) is used. 4. Convert RNA into cDNA using the TaqMan® microRNA reverse transcription kit. Make sure to use a starting RNA concentration of 8 ng. 5. For more accurate pipetting, create 1 Master mix including 100 mM dNTPs, Multiscribe Reverse Transcriptase, 50 U/μ L, 10 Reverse transcription buffer, RNase Inhibitor, 20 U/μ L, Nuclease-Free water, and the target miRNA RT primer. Make sure the RT primer is not purple in color, as this is the Taqman RNA assay used for QPCR. 6. For QPCR, use 1 μL of cDNA product from the prior step, 1 μL 20 miRNA assay (protected from the light), 10 μL 2 Universal PCR Master Mix (No AmpErase UNG), and 9 μL nuclease-free water per replicate. Applied Biosystems recommends performing four replicates of each reaction. 7. Thermal cycling parameters should match those listed in the Taqman® protocol. For the anneal/extend stop, modify the temperature to be 55  C. 8. Quantify values of relative gene expression by normalizing against a miRNA control such as U6.

4

Notes 1. Clean the ear puncher and forceps after each sampling to minimize the chance of cross-contamination. 2. Briefly vortex and then spin down the tubes to make sure the samples are in contact with the solution. Make sure tissues are not stuck on the wall or in the cap of the tubes before the next step. 3. Using too much DNA template can lead to having no products after the final PCR cycle; therefore, measuring DNA concentration and using an equal quantity of DNA for all samples is

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essential, particularly for miR-133aTg genotyping where mice with no bands after electrophoresis are considered wild type. 4. Besides the restriction enzyme digest protocol explained here, there are two other methods available for Akita genotyping according to The Jackson Laboratory: endpoint analysis and pyrosequencing. 5. Prepare a restriction digest master mix (Fnu4HI, buffer G, and water) according to the number of samples to minimize variation among your samples. 6. The amount of enzyme for each reaction, total volume, and incubation time may need to be optimized for the experimental setup. 7. Safer alternatives to ethidium bromide can be used for detection of DNA. 8. Always use negative and positive controls for each genotyping. 9. Keeping the measurement times consistent in different days, especially if the mice are not fasted, helps to minimize variation in glucose levels. 10. Removing food from the old cage is not enough for fasting because small pieces of food can still be present in the bedding. Transfer mice to a new cage with water and no food.

Acknowledgments This study was supported, in part, by the National Institutes of Health grants HL-113281 and HL-116205 to Paras K. Mishra. We greatly appreciate Dr. Scot Matkovich from the Washington University, St. Louis for his kind gift of cardiac-specific miR-133aTg mice. References 1. Boudina S, Abel ED (2010) Diabetic cardiomyopathy, causes and effects. Rev Endocr Metab Disord 11(1):31–39 2. Bugger H, Abel ED (2014) Molecular mechanisms of diabetic cardiomyopathy. Diabetologia 57:660–671 3. Kannel WB, Hjortland M, Castelli WP (1974) Role of diabetes in congestive heart failure: the Framingham study. Am J Cardiol 34(1):29–34 4. Kasznicki J, Drzewoski J (2014) Heart failure in the diabetic population-pathophysiology, diagnosis and management. Arch Med Sci 10 (3):546–556 5. Nichols GA, Gullion CM, Koro CE et al (2004) The incidence of congestive heart

failure in type 2 diabetes: an update. Diabetes Care 27:1879–1884 6. Badole SL, Jangam GB (2014) Animal models of diabetic cardiomyopathy. Elsevier Inc., Amsterdam 7. Wang J, Takeuchi T, Tanaka S et al (1999) A mutation in the insulin 2 gene induces diabetes with severe pancreatic β-cell dysfunction in the Mody mouse. J Clin Invest 103(1):27–37 8. Ross SA, Gulve EA, Wang M (2004) Chemistry and biochemistry of type 2 diabetes. Chem Rev 104(3):1255–1282 9. Coleman DL (1978) Obese and diabetes: two mutant genes causing diabetes-obesity syndromes in mice. Diabetologia 14(3):141–148

miRNA Cardioprotection of Diabetic Hearts 10. Fuentes-Antras J, Picatoste B, GomezHernandez A et al (2015) Updating experimental models of diabetic cardiomyopathy. J Diabetes Res 2015:656795 11. Nandi SS, Shahshahan HR, Shang Q et al (2018) MiR-133a mimic alleviates T1DMinduced systolic dysfunction in Akita: an MRI-based study. Front Physiol 9:1–11 12. Yoshioka M, Kayo T, Ikeda T, Koizuni A (1997) A novel locus, Mody4, Distal to D7Mit189 on chromosome 7 determines early-onset NIDDM in nonobese C57BL/6 (Akita) mutant mice. Diabetes 46(5):887–894 13. Basu R, Oudit GY, Wang X et al (2009) Type 1 diabetic cardiomyopathy in the Akita (Ins2WT/C96Y) mouse model is characterized by lipotoxicity and diastolic dysfunction with preserved systolic function. Am J Physiol Circ Physiol 297(6):H2096–H2108 14. Kesherwani V, Shahshahan HR, Mishra PK (2017) Cardiac transcriptome profiling of diabetic Akita mice using microarray and next generation sequencing. PLoS One 12(8):1–17 15. Chavali V, Tyagi SC, Mishra PK (2013) Predictors and prevention of diabetic cardiomyopathy. Diabetes Metab Syndr Obes 6:151–160 16. Chavali V, Tyagi SC, Mishra PK (2014) Differential Expression of Dicer, miRNAs, and Inflammatory Markers in Diabetic Ins2+/
Akita Hearts. Cell Biochem Biophys 68 (1):25–35 17. Nandi SS, Zheng H, Sharma NM et al (2016) Lack of MIR-133a decreases contractility of diabetic hearts: A role for novel cross talk between tyrosine aminotransferase and tyrosine hydroxylase. Diabetes 65(10):3075–3090 18. Rosengren A, Jing X, Eliasson L, Renstro¨m E (2008) Why treatment fails in type 2 diabetes. PLoS Med 5(10):1426–1427

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19. Hong EG, Dae YJ, Hwi JK et al (2007) Nonobese, insulin-deficient Ins2Akita mice develop type 2 diabetes phenotypes including insulin resistance and cardiac remodeling. Am J Physiol Endocrinol Metab 293(6): E1687–E1696 20. Mishra PK, Tyagi N, Kumar M, Tyagi SC (2009) MicroRNAs as a therapeutic target for cardiovascular diseases. J Cell Mol Med 13 (4):778–789 21. Guo R, Nair S (2017) Role of microRNA in diabetic cardiomyopathy: From mechanism to intervention. Biochim Biophys Acta Mol basis Dis 1863(8):2070–2077 22. Leptidis S, el Azzouzi H, Lok SI et al (2013) A deep sequencing approach to uncover the miRNOME in the human heart. PLoS One 8(2): e57800 23. Care` A, Catalucci D, Felicetti F et al (2007) MicroRNA-133 controls cardiac hypertrophy. Nat Med 13(5):613–618 24. VanderKaay S, Letts L, Jung B, Moll SE (2020) Doing what’s right: a grounded theory of ethical decision-making in occupational therapy. Scand J Occup Ther 27(2):98–111 25. Chen S, Puthanveetil P, Feng B et al (2014) Cardiac miR-133a overexpression prevents early cardiac fibrosis in diabetes. J Cell Mol Med 18(3):415–421 26. Matkovich SJ, Wang W, Tu Y et al (2010) MicroRNA-133a protects against myocardial fibrosis and modulates electrical repolarization without affecting hypertrophy in pressureoverloaded adult hearts. Circ Res 106 (1):166–175 27. Truett GE, Heeger P, Mynatt RL et al (2000) Preparation of PCR-quality mouse genomic dna with hot sodium hydroxide and tris (HotSHOT). BioTechniques 29(1):52–54. https:// doi.org/10.2144/00291bm09

Chapter 9 Generating a Podocyte-Specific Neonatal F Receptor (FcRn) Knockout Mouse Judith Blaine Abstract Proteinuria is a widely used marker of renal disease and is strongly associated with renal and cardiovascular outcomes. The molecular mechanisms underlying filtration of serum proteins through the glomerular filtration barrier (GFB) remain to be determined. Since the GFB is a complex structure, studies of albumin or IgG trafficking in cultured cells in vitro may not fully recapitulate these processes in vivo. In other epithelial cells including renal proximal tubular cells, the neonatal Fc receptor (FcRn) is required to divert albumin and IgG from the degradative pathway which allows these proteins to be recycled or transcytosed. To examine the role of podocyte FcRn in albumin and IgG trafficking in vivo, we detail the creation of a podocyte-specific FcRn knockout mouse and describe methods for examining intraglomerular detection of albumin and IgG in these mice. Key words Neonatal Fc receptor, Podocyte, Knockout mice, Albuminuria

1

Introduction Multiple studies have shown that proteinuria is an independent marker of kidney disease progression [1, 2]. While in clinical practice, proteinuria is widely used as one of the measures of renal dysfunction, the exact mechanisms of how serum proteins such as albumin and IgG are handled by the kidney remain to be fully determined. Both the glomerulus and the proximal tubules are involved in the renal handling of serum proteins. The primary barrier to filtration of large plasma proteins such as albumin and IgG into the urine is the glomerular filtration barrier (GFB) which consists of three layers—a fenestrated endothelium, the glomerular basement membrane, and the podocyte [3]. The podocyte is a specialized epithelial cell that plays a key role in preventing passage of significant amounts of albumin or IgG through the GFB. Each podocyte consists of a large cell body and multiple processes which ramify to form smaller processes. The slit diaphragm, which extends between the foot processes of neighboring podocytes, precludes

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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filtration of proteins ~70 kDa or larger through the GFB. The precise amount of albumin filtered through the GFB is a hotly contested topic [4, 5]. By even the most conservative estimates, at least 4 g albumin per day pass through the GFB [6]. The amount of IgG that passes through the glomerular filtration barrier is unknown. The neonatal Fc receptor (FcRn) is a protein that has homology to major histocompatibility complex class I and binds albumin and IgG at acidic pH but has minimal affinity for these proteins at neutral pH [7]. Within the adult kidney, FcRn is expressed in podocytes, vascular endothelial cells, and the proximal tubule [7, 8]. In proximal tubule cells and the vasculature, the neonatal Fc receptor (FcRn) prevents albumin and IgG from being degraded, thereby allowing albumin to be recycled or transcytosed [9–12]. Previous work has shown that global knockout of FcRn results in the intraglomerular accumulation of IgG in mice [13]. However, since FcRn is required for recycling of albumin and IgG through endothelial cells back to the vasculature, global FcRn KO mice are significantly hypoalbuminemic and hypogammaglobulinemic compared to wild type (WT) mice [14]. Thus, podocytes in the global KO mice are exposed to significantly less albumin and IgG than WT mice which might alter handling of these proteins. In order to study albumin and IgG trafficking through the GFB in mice with normal plasma levels of these proteins, we created podocyte-specific FcRn knockout mice.

2

Materials

2.1

Mice

2.2

Genotyping

FcRn floxed mice [15] were a kind gift of Dr. Sally Ward (UT Southwestern). Podocin-Cre mice (stock number 008205) were obtained from Jackson Laboratories (Bar Harbor, Maine). Both the FcRn floxed mice and the podocin-Cre mice have been bred onto a C57BL6 background. 1. Primers for determining if the mouse is homozygously floxed (FcRn fl/fl;+/+) or has the desired FcRn knockout in podocytes (FcRn fl/fl;Pod-Cre/+) were obtained from Integrated DNA Technologies (Coralville, IA) and are as follows: Primer set 1: For detection of the floxed allele: Forward primer (50 TTA TGT GGA GAT GGA AAA CAA CCA TGT ACA 30 ) and reverse primer (50 CCA TGG CTT TCT CTC AGC GGC GAT GAC 30 ). Primer set 2: For detection of the deleted (Cre) allele: Forward primer (50 GCG GTC TGG CAG TAA AAA CTA 30 ) and reverse primer (50 GTC AAA CAG CAT TGC TGT CAC 30 ).

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2. DNA isolation from mouse ear samples: Viagen DirectPCR kit (Thermo Fisher Scientific, Waltham MA, proteinase-K (Thermo Fisher Scientific). 3. PCR: GoTaq™ Hot Start Polymerase: Green Master Mix, 2 (Promega, Madison WI), BioRad T-100 Thermal Cycler, PCR tubes (VWR, Radnor PA). 2.3 Phenotype Characterization

1. Albuwell M kit (Exocell, Philadelphia PA), Quantichrom™ Creatinine Assay Kit (Bioassay Systems, Hayward CA), Quantichrom™ Urea Assay Kit (Bioassay Systems), 96-well plates, 1.5 mL microcentrifuge tubes coated with heparin sodium 30,000 units per 30 mL, Marathon 16 Km centrifuge (Thermo Fisher Scientific, Waltham, MA).

2.4

1. Tissue-Tek optimal cutting temperature compound (VWR), 5%, 10%, and 25% sucrose in phosphate-buffered saline (PBS), Fisherbrand™ 12-550-15 Super frost Plus Microscope Slides (Thermo Fisher Scientific), Fisherbrand™ Cover Glasses: Rectangles No. 1.5 (Fisher Scientific). Phosphatebuffered saline (PBS), PBS + 0.1% Triton-X 100 (PBS-X), 4% Periodate-Lysine-Paraformaldehyde (PLP) (0.075 M lysine, 0.37 M sodium phosphate (pH 7.2), 2% formaldehyde, and 0.01 M NaIO4), normal goat serum, ImmEdge™ Hydrophobic Pap Pen (Vector Laboratories, Burlingame, CA), Vectashield® Mounting Media (Vector Laboratories), nail polish.

Imaging

2. Primary antibodies: anti-mouse IgG (GW20083F, SigmaAldrich. St. Louis MO), anti-mouse albumin (ab106582, Abcam, Cambridge, MA). Secondary antibodies: Alexa 488-conjugated goat anti-chicken IgG (Invitrogen), Alexa 633-conjugated phalloidin (Invitrogen). 3. Zeiss 780 laser-scanning confocal/multiphoton-excitation fluorescence microscope with a 34-Channel GaAsP QUASAR Detection Unit and non-descanned detectors (Zeiss, Thornwood NY), Zeiss C-Apochromat 40/1.2NA Korr FCS M27 water-immersion lens objective. 4. Image analysis: Image J software (NIH, Bethesda, Maryland).

3

Methods

3.1 Creation of Podocyte-Specific FcRn Knockout Mice

1. To create podocyte-specific FcRn KO mice, male podocin-Cre mice are bred with female FcRn floxed mice (see Notes 1–3). Ear samples are used to obtain DNA for genotyping (see Note 4). DNA is isolated from each ear sample by digesting the sample in 100 μL Viagen Direct PCR and 2 μL Proteinase K and placing the sample at 55  C overnight. The following morning the samples are placed at 85  C for 45 min. The DNA can be stored at 4  C prior to being used for PCR.

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2. For each mouse to be genotyped, two different PCR reactions are run. In one reaction, primer set 1 (Flox PCR) is used and in the other reaction primer set 2 (Cre PCR) is used. PCR reactions are set up as follows: (a) For the Flox PCR (Primer Set 1): To each PCR tube add the following: 6.9 μL Nanopure water, 12.5 μL GoTaq Hotstart Polymerase Green Master Mix 2X, 0.3 μL primer 1 (0.5 μM final concentration), 0.3 μL primer 2 (0.5 μM final concentration), 5 μL diluted DNA (dilute DNA isolated from mouse ear 1:10). (b) Reactions for Flox are run on a Biorad T-100 Thermal Cycler PCR machine using the following settings: 95  C for 4 min then 30 cycles of 95  C for 30 s, 55  C for 30 s, 72  C for 60 s. At the completion of these cycles, 72  C for 5 min. (c) PCR reactions for Cre (Primer set 2) are set up as follows: To each PCR tube add the following: 6.9 μL Nanopure water, 12.5 μL GoTaq Hotstart Green Master Mix 2X, 0.3 μL primer 1 (0.5 μM final concentration), 0.3 μL primer 2 (0.5 μM final concentration), 5 μL diluted DNA (1:10). (d) Reactions for Cre are run on a Biorad T-100 Thermal Cycler PCR machine using the following settings: 95  C for 4 min then 35 cycles of 94  C for 30 s, 51.7  C for 60 s, 72  C for 60 s. At the completion of these cycles, 72  C for 2 min. 3. PCR products are run on a 2% agarose gel. Typical results are shown in Fig. 1. For the Flox PCR, a homozygous FcRn floxed mouse will show a band at 400 bp while a wild type mouse will show a band at 300 bp. A heterozygous mouse will show a band at both 300 bp and 400 bp. For the Cre PCR, mice in which the podocin-Cre gene has been expressed will show a band at 200 bp. Thus, control (FcRn fl/fl:+/+) mice are homozygous for the FcRn floxed gene whereas podocytespecific FcRn KO (FcRn fl/fl; Pod-Cre) mice are double transgenic resulting in no FcRn expression in podocytes. In Fig. 1, mouse 3 is a control mouse (FcRn fl/fl;+/+) whereas mice 4 and 5 are podocyte-specific FcRn knockout mice (FcRn fl/ fl;Pod-Cre/+). Figure 2a shows FcRn expression in a glomerulus from a control and a podocyte-specific FcRn knockout mice. In the KO, FcRn is absent from peripherally located glomerular cells which is where podocytes are located. Figure 2b shows that FcRn expression is significantly reduced in podocytes isolated from podocyte-specific FcRn knockout mice compared to controls.

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B

A Flox genotyping

1 2 3 4 5 Flox

400 bp 300 bp

Cre genotyping

Cre

1 2 3 4 5 200 bp

Fig. 1 Agarose gels of PCR performed for genotyping of the Flox gene (a) and Cre gene (b). Separate PCR and gels are performed on the same sample for each animal due to the unknown gene expression pattern prior to genotyping. (a) Gel on the left (Flox genotype) is a 2% agarose gel looking at expression of the floxed FcRn gene. Lanes labeled 1 and 2 are animals with FcRn heterozygous flox expression. Lane 3 is a previously known homozygous floxed FcRn mouse used as a positive control. Lanes 4 and 5 are newly identified homozygous FcRn floxed mice. (b) Cre genotyping: 2% agarose gel looking at expression of the podocin-cre gene. Lane labeled 1–5 correspond to samples from mice labeled 1–5 in the gel on left (a: Flox genotyping). Lanes 1, 2, 4, and 5 demonstrate expression of podocin-Cre gene. Taken together, these genotyping gels show that mouse 3 is a control mouse (FcRn fl/fl;+/+) while mice 4 and 5 are podocyte-specific FcRn knockout mice (FcRn fl/fl; Pod-Cre/+) 3.2 Phenotypic Analysis of Control and Podocyte-Specific FcRn Knockout Mice

1. For phenotypic characterization of control or podocyte-specific FcRn KO mice, we measure albuminuria and blood urea nitrogen (BUN). Urine is collected by holding the mouse over a 96-well plate and collecting urine from the plate wells. Urine can be stored at 80  C for further analysis. Blood is collected either by submandibular bleed or via cardiac puncture at the time of sacrifice. 2. After collection into a heparinized 1.5 mL centrifuge tube, blood is spun at 1000 rpm (123  g) for 5 min in a Fisher Marathon 16 Km centrifuge and then the serum is pipetted off and stored at 80  C for further analysis. 3. Albuminuria is measured using the Albuwell M kit according to the manufacturer’s directions. BUN is measured using the Quantichrom Urea Assay Kit according to the manufacturer’s directions. Urine creatinine is measured using the Quantichrom Creatinine Assay Kit. Urinary albumin is normalized to urinary creatinine to account for differences in concentration of the urine. Albuminuria and BUN results are shown for 3-month and 6-month-old podocyte-specific FcRn KO mice in Fig. 3.

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A

FcRn fl/fl;+/+

FcRn fl/fl;Pod-Cre/+

* FcRn Actin

B

FcRn fl/fl;+/+

FcRn Actin

FcRn Actin

*

FcRn fl/fl;Pod-Cre/+

FcRn Actin

Fig. 2 FcRn expression in control and podocyte-specific FcRn KO mice. (a) In control (FcRn fl/fl:+/+) mice FcRn (green) is located in cells peripherally located in the glomerulus in the expected location of podocytes (arrows) as well as in endothelial cells. In the podocyte-specific FcRn KO (FcRn fl/fl; Pod-Cre/+) mice, there is no FcRn expression in peripherally located cells. FcRn is expressed in endothelial cells in the glomerulus (arrowheads) as well as in larger blood vessels (asterisk). Actin is stained blue. Scale bar: 10 μm. (b) Podocytes isolated from control mice have significantly increased FcRn expression compared to podocyte-specific FcRn KO mice. *, Scale bar: 10 μm 3.3 Immunolocalization of Intraglomerular Albumin and IgG in Control and Podocyte-Specific FcRn KO Mice

1. For the in vivo immunolocalization studies, the kidneys are cleared of blood by perfusion of phosphate-buffered saline (PBS). After removal, the kidneys are kept in PLP for 24 h. Following fixation, the kidneys are infused with 5% sucrose for 2 h, 10% sucrose for 2 h, and 25% sucrose overnight then embedded in OCT and frozen in liquid nitrogen. 2. Sections are cut from OCT blocks on a cryostat at 3 μm and placed on glass slides. We place no more than 3 sections per slide. 3. For immunofluorescence studies, a circle is drawn around each tissue section using a Pap pen. The sections are permeabilized in PBS-X at room temperature for 10 min. Sections are blocked in 10% serum (use the serum of the animal the secondary antibody was made in which in this case is goat) and 1% BSA. Block for 30 min at room temperature. 4. After aspiration of the blocking solution, primary antibodies are applied. Primary antibodies are diluted in 10% blocking solution in PBS-X and are as follows: For IgG staining: antimouse IgG made in chicken (1:250) and for albumin staining:

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Fig. 3 There is no significant difference in albuminuria or BUN levels in control and podocyte-specific FcRn KO mice at 3 or 6 months. (a and b) urinary albumin normalized to creatinine in 3-month and 6-month-old control or podocyte-specific FcRn KO mice. Figure is modified from [16]. (c and d) BUN levels creatinine in 3-month and 6-month-old control or podocyte-specific FcRn KO mice

anti-mouse albumin (1:250) (see Note 5). Depending on the size of the Pap circle, approximately 100 μL antibody solution is needed per section. Primary antibody incubations are done at overnight at 4  C in a humidified chamber. 5. The following day sections are washed in PBS-X three times for 5 min each. After washing, the sections are incubated at room temperature for 60 min with secondary antibodies: Alexa 488-conjugated goat anti-chicken IgG (1:250) and Alexa 633-conjugated phalloidin (1:200) to stain actin. The sections must be protected from light during incubation. 6. After incubation in secondary antibodies, sections are washed 3 times in PBS-X (5 min each) and then once in PBS. Sections are mounted using Vectashield and coverslipped. The edges of

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the coverslip are sealed with nail polish. The sections need to be protected from light during washes and after application of the coverslip. 3.4

Imaging

1. Images are acquired on a confocal microscope Zeiss 780 laserscanning confocal/multiphoton-excitation fluorescence microscope with a 34-Channel GaAsP QUASAR Detection Unit and non-descanned detectors. Imaging settings are initially set to avoid saturation, minimize contributions from tubular autofluorescence, and maximize the signal-to-noise ratio (see Note 6). To allow comparison between images, settings are kept constant between different samples. The objective used for imaging is a Zeiss C-Apochromat 40/1.2NA Korr FCS M27 water-immersion lens objective. The laser used is a 30 mW Argon Laser using excitation at 488 nm and HeNe 5 mW (633 nm). Image acquisition is performed using Zeiss ZEN 2012 software. 2. Post-acquisition images are analyzed using Image J software (NIH). For each glomerulus, the fluorescence intensity of albumin or IgG is normalized to the glomerular area. 20–25 glomeruli are analyzed per mouse. Results of imaging experiments are shown in Fig. 4. Podocyte-specific FcRn KO mice show an increase in intraglomerular IgG accumulation over time whereas there is no significant difference in intraglomerular albumin accumulation between control and KO mice.

4

Notes 1. For breeding, we always carry the Podocin-Cre gene in the male mice. That is we breed male podocin-Cre mice with female floxed FcRn mice. 2. The podocin-Cre/FcRn floxed breeding pairs tend to produce small litters but the mice are viable and survive for at least 12–18 months. 3. Even though the homozygously FcRn floxed mice have been bred onto a C57BL6 background, these mice are grey in appearance. Due the presence of homozygous flox alleles, the podocyte-specific FcRn flox mice are also grey in color. 4. We use ear samples rather than tail samples for genotyping as ears samples tend to give higher DNA yields. 5. While we have been able to costain for intraglomerular albumin and other proteins of interest, we have been unable to costain for IgG and any other protein. This may be due to blocking/ interference from the anti-IgG antibody.

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Fig. 4 Podocyte-specific FcRn KO mice have a significant increase in intraglomerular IgG accumulation over time but no change in albumin accumulation. (a), Intraglomerular IgG: At 3 months of age, there is no significant difference in intraglomerular IgG accumulation in control (FcRn fl/fl;+/+) versus podocyte-specific FcRn KO (FcRn fl/fl;Pod-Cre/+) mice (n ¼ 3 mice per group). By 6 months of age, there is a statistically significant increase in intraglomerular IgG accumulation in the knockout compared to controls (n ¼ 6 control and 6 KO mice, * p < 0.05). Intraglomerular IgG accumulation is even more significantly increased by 12 months of age in the podocyte-specific FcRn KO (n ¼ 6 control and 6 KO mice, **p < 0.0001). Scale bar 20 μm. The figure is taken from [16]. (b) Intraglomerular Albumin: There is minimal albumin accumulation in both control and podocyte-specific FcRn KO mouse glomeruli, and there is no significant difference between control and KO animals at 3, 6, or 12 months. By 12 months of age, both control and podocyte-specific FcRn KO mice have significantly less intraglomerular albumin than 3-month-old control or KO animals, *p < 0.01, **p < 0.0001. Scale bar 20 μm. NS ¼ not significant. Number of mice per group is the same as in (a). The figure is taken from [16]

6. For imaging experiments, we never use the 488 nm laser at more than 5% power as cellular autofluorescence increases significantly above this laser power.

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Acknowledgments This work was funded by a Norman Coplon Satellite Healthcare grant and NIH R01DK104264 to JB. References 1. Bakris GL (2008) Slowing nephropathy progression: focus on proteinuria reduction. Clin J Am Soc Nephrol 3:S3–S10 2. Hemmelgarn BR, Manns BJ, Lloyd A, James MT, Klarenbach S, Quinn RR, Wiebe N, Tonelli M (2010) Relation between kidney function, proteinuria, and adverse outcomes. JAMA 303:423–429 3. Scott RP, Quaggin SE (2015) Review series: the cell biology of renal filtration. J Cell Biol 209:199–210 4. Peti-Peterdi J (2009) Independent two-photon measurements of albumin GSC give low values. Am J Physiol Ren Physiol 296:F1255–F1257 5. Russo LM, Sandoval RM, McKee M, Osicka TM, Collins AB, Brown D, Molitoris BA, Comper WD (2007) The normal kidney filters nephrotic levels of albumin retrieved by proximal tubule cells: retrieval is disrupted in nephrotic states. Kidney Int 71:504–513 6. Castrop H, Schiessl IM (2016) Novel routes of albumin passage across the glomerular filtration barrier. Acta Physiol (Oxford, England) 219(3):544–553 7. Kuo T, Baker K, Yoshida M, Qiao S-W, Aveson V, Lencer W, Blumberg R (2010) Neonatal Fc receptor: from immunity to therapeutics. J Clin Immunol 30:777–789 8. Dickinson BL (2016) Unraveling the immunopathogenesis of glomerular disease. Clin Immunol 169:89–97 9. Andersen JT, Sandlie I (2009) The Versatile MHC class I-related FcRn protects IgG and albumin from degradation: implications for development of new diagnostics and therapeutics. Drug Metab Pharmacokinet 24:318–332 10. Chaudhury C, Mehnaz S, Robinson JM, Hayton WL, Pearl DK, Roopenian DC, Anderson CL (2003) The major histocompatibility

complex–related Fc receptor for IgG (FcRn) binds albumin and prolongs its lifespan. J Exp Med 197:315–322 11. Kim J, Bronson CL, Hayton WL, Radmacher MD, Roopenian DC, Robinson JM, Anderson CL (2006) Albumin turnover: FcRn-mediated recycling saves as much albumin from degradation as the liver produces. Am J Physiol Gastrointest Liver Physiol 290:G352–G360 12. Tenten V, Menzel S, Kunter U, Sicking E-M, van Roeyen CRC, Sanden SK, Kaldenbach M, Boor P, Fuss A, Uhlig S, Lanzmich R, Willemsen B, Dijkman H, Grepl M, Wild K, Kriz W, Smeets B, Floege J, Moeller MJ (2013) Albumin is recycled from the primary urine by tubular transcytosis. J Am Soc Nephrol 24 (12):1966–1980 13. Akilesh S, Huber TB, Wu H, Wang G, Br H, Kopp JB, Miner JH, Roopenian DC, Unanue ER, Shaw AS (2008) Podocytes use FcRn to clear IgG from the glomerular basement membrane. Proc Natl Acad Sci USA 105:967–972 14. Roopenian DC, Christianson GJ, Sproule TJ, Brown AC, Akilesh S, Jung N, Petkova S, Avanessian L, Choi EY, Shaffer DJ, Eden PA, Anderson CL (2003) The MHC class I-like IgG receptor controls perinatal IgG transport, IgG homeostasis, and fate of IgG-Fc-coupled drugs. J Immunol 170:3528–3533 15. Montoyo HP, Vaccaro C, Hafner M, Ober RJ, Mueller W, Ward ES (2009) Conditional deletion of the MHC class I-related receptor FcRn reveals the sites of IgG homeostasis in mice. Proc Natl Acad Sci 106:2788–2793 16. Dylewski J, Dobrinskikh E, Lewis L, Tonsawan P, Miyazaki M, Jat PS, Blaine J (2019) Differential trafficking of albumin and IgG facilitated by the neonatal Fc receptor in podocytes in vitro and in vivo. PLoS One 14: e0209732

Chapter 10 Mouse Models of Colitis-Associated Colon Cancer Santhakumar Manicassamy, Puttur D. Prasad, and Daniel Swafford Abstract Crohn’s disease (CD) and ulcerative colitis are two main clinically defined forms of chronic inflammatory bowel disease (IBD). Chronic intestinal inflammation is inextricably linked to colitis-associated colon carcinogenesis (CAC). Patients with ulcerative colitis (UC) and Crohn’s disease (CD) have an increased risk of colon cancer. Our understanding of IBD and IBD-associated colon carcinogenesis depends largely on rodent models. AOM-DSS-induced colitis-associated colon cancer in mice is the most widely used and accepted model that can recapitulate the human IBD-associated colon cancer. Here, we have provided detailed protocols of this mouse model of experimentally induced chronic intestinal inflammationassociated colon cancer. We will also discuss the protocols for the isolation and analysis of inflammatory immune cells from the colon. Key words IBD-associated colon cancer, AOM, DSS, Chronic colitis, CAC, Flow cytometry and intestinal immune cells

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Introduction Inflammatory bowel disease (IBD) is an important public health problem in western society, affecting more than two million Americans. Crohn’s disease (CD) and Ulcerative colitis are two main clinically defined forms of chronic inflammatory bowel disease (IBD) with multifactorial etiologies [1, 2]. Accumulating evidence suggests that chronic intestinal inflammation is inextricably linked to colitis-associated colon carcinogenesis (CAC) [3]. Patients with ulcerative colitis (UC) and Crohn’s disease (CD) have an increased risk of colon cancer, which is dependent on the duration, extent, and severity of inflammatory disease [4, 5]. Patients with IBD suffer from abdominal pain, diarrhea, weight loss, and rectal bleeding [1]. Biopsies from IBD patients with active disease show infiltration of mononuclear and polymorphonuclear leukocytes; epithelial cell erosion, loss of goblet cells, crypt abscesses, and epithelial cell hyperplasia [6]. Although the pathogenesis of IBD and CAC is largely unknown, accumulating evidence suggests three critical interacting elements involved in the pathogenesis of CAC—the

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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gut mucosal immune response, intestinal microflora, and host genetic susceptibility factor and environment. Immune cells in the intestine have emerged as principal effectors in the pathogenesis of IBD and CAC [7–9]. Thus, our understanding of IBD and IBD-associated colon cancer depends largely on animal models that can recapitulate the human diseases. In general, azoxymethane/dextran sodium sulphate (AOM-DSS)-induced colon tumorigenesis is the most widely accepted mouse model of CAC. In this model system, DSS-mediated injury induces intestinal inflammation that contributes to colon carcinogenesis caused by azoxymethane (AOM). This model is also useful in evaluating the efficacy of prolonged prophylactic or therapeutic treatment of colitis and CAC. In addition to AOM, there are other carcinogens such as MAM acetate, DMH, etc. that can induce colon tumors to varying extent [10]. These models are also useful in delineating the signaling pathways and evaluating the effects of nutritional components on gut inflammation and CAC [11]. In addition, this model serves as an excellent platform for experimental applications and preclinical testing of novel therapeutic studies [11]. Spontaneous models of chronic colitis are based on genetic defects in immune regulation or targeted gene deletion that are involved in immune regulation (IL-10-deficient mice). Several recent excellent review articles and other manuscripts provide a more detailed description of different animal models of intestinal inflammation and CAC [1, 9–14]. This unit provides a detailed protocol of the most commonly used mouse model of CAC, AOM-DSS-induced colon tumorigenesis.

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Materials

2.1 AOM-DSSInduced InflammationAssociated Colon Cancer

1. Animals: Male or Female C57BL/6 mice (8–12 weeks old) purchased from Jackson Laboratories. Mice are maintained in a barrier SPF room with a 12 h dark/12 h light cycle. Other strains can also be used but susceptibility and severity may vary. All animal experiments must be done in accordance with institutional and national guidelines and regulations (see Note 1). 2. 3% (w/v) Dextran Sulfate Sodium salt (DSS; MW: 36,000–50,000 Da; MP Biomedicals). Weigh 15 g DSS powder and dissolve in 500 mL of autoclaved drinking water. Store until use at 4  C (see Note 2). 3. Azoxymethane (AOM, 1 mg/mL) (Sigma). Dissolve 10 mg of AOM in 10 mL of sterile phosphate-buffered saline (PBS). Aliquot about 500 μL into 1.5 mL sterile microcentrifuge tubes and store until use at 20  C (see Note 3). 4. Phosphate-buffered saline 1 (PBS).

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5. 70% Ethyl alcohol (Pharmco-AAPER): add 70 mL of absolute alcohol to 30 mL sterile distilled water. 6. Digital caliper (Fisher Scientific). 7. Tissue path Macrosette cassettes (Fisher Scientific). 8. Dissection tools. 9. 1 and 10 mL syringes (BD Biosciences). 10. 281/2 Needles. 11. o-Dianisidine dihydrochloride (Sigma-Aldrich). 1 mg/mL of dianisidine in PBS (see Note 4). 12. Hexadecyltrimethylammonium bromide (Sigma-Aldrich). Dissolve 0.5 g Hexadecyltrimethylammonium bromide in 1 mL of 50 mM PBS (pH 6.0). 13. Fluorescein isothiocyanate (FITC)-dextran (average molecular weight, 4 kDa; Sigma-Aldrich). 14. Biotek EL808 Absorbance plate reader (Biotek). 15. Microtest plate 96-well flat bottom (Fisher). 16. 30% H2O2 (Sigma-Aldrich). Add 4 ul of 30% H2O2 to 96 ul of H2O. 17. Fetal bovine serum (Atlanta Biologicals). 18. 0.5 M EDTA (pH 8.0) (cellgro). 19. Trypan blue (Invitrogen). 20. DNase I (Roche): Prepare stock solution by dissolving 1 mg of DNase I in 50 mL of HBSS. Store in 20 freezer. 21. Collagenase VIII (Sigma-Aldrich): Prepare stock solution of 100 U/mL in complete RPMI medium. Store in 20  C freezer. 22. Collagenase IV (Worthington): Prepare stock solution of 4000 U/mL in complete RPMI medium. Store in 20  C freezer. 23. Percoll (GE Healthcare): Prepare 100% percoll by mixing 90 mL of percoll with 10 mL of 10 PBS. Prepare 40% percoll by mixing 40 mL of 100% percoll with 60 mL of 1 PBS. 24. 15 and 50 mL conical tubes (BD Biosciences). 25. 100 μm, 70 μm, and 40 μm cell strainer (BD Biosciences). 26. Brefeldin A and monensin (BD Pharmingen). 27. PMA and Ionomycin (Sigma Aldrich). 28. FACS Buffer: 1 PBS Containing 0.5% FBS. 29. CMF/HEPES: HBSS (Ca2+ and Mg2+ free) containing 2 mM HEPES and 2% FBS.

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Methods

3.1 AOM-DSSInduced Colon Cancer 3.1.1 Treating Mice with AOM

1. Acclimatize the C57BL/6 mice for 1–2 weeks before the experiment. Utilize age- and gender-matched 8–12 weeks old mice. 2. One week before the DSS treatment (Day 7), weigh each mouse at least two times, and record accurate weight to ensure uniform dosing of AOM (see Fig. 1). Calculate the total volume of AOM required to achieve a dose of 10 mg/kg body weight. 3. After calculating the required AOM dosage, using a 281/ 2 tuberculin syringe inject each mouse intraperitoneally with the appropriate volume of AOM. For example, a 20 g mouse would receive a 200 μL injection of 1 mg/mL AOM solution (see Note 5). Weigh and monitor the mice for the next 48 h.

3.1.2 Treating Mice with DSS

Start first DSS cycle (Day 0–6) 1. After one week of AOM injection, replace the drinking water with 3.0% DSS water in appropriate mouse cages (see Note 6). Estimate 5 mL DSS solution per mouse per day (see Note 7). Control mice receive the same drinking water without DSS.

2. Replace the remaining DSS solution with regular water on day 6. This is followed by 2 weeks of recovery in regular water. 3. Monitor body weight and disease activity index every other day until the experiment is terminated. These are important parameters indicating the colitis severity between experimental groups (see Note 8). Start second DSS cycle (Day 21–27). 4. After the first recovery period, replace the regular water with 3% DSS water and leave the mice with DSS water from day 21–27. This is followed by 2 weeks of recovery in regular water (second recovery period) (see Fig. 1) Start third DSS cycle (Day 42–48). 5. After the second recovery period, replace the regular water with 3% DSS water and leave the mice with DSS water from day 42–46. This is followed by 2 weeks of recovery in regular water (third recovery period) (see Fig. 1) 3.2 Sacrificing Animals and Tissue Harvesting

To assess the histopathology and changes in gene expression, animals can be sacrificed at the end of the experiments (day 62) by the institution’s approved method (see Note 9). Using standard dissecting scissors, make a large incision in its ventral (belly) side. Locate and carefully make incisions at the connections of the ileum/cecum and rectum/anus to remove the entire colon. Additional tissues such as mesenteric lymph node and spleen can be

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Fig. 1 Treatment scheme of the AOM-DSS protocol for induction of colitisassociated colon cancer. Mice are injected intraperitoneally with azoxymethane (10 mg/kg body weight) and then subjected to three cycles of DSS treatment, wherein one cycle constituted mice being given 3% DSS (36–50 kDa) in their drinking water for 6 days followed by 14 days of normal drinking water. At day 9 and 62, colons of mice are analyzed for inflammation and cancer, respectively

harvested at this time. Determine colon length (digital caliper) and weight. Remove the fecal contents by rinsing the colon with sterile PBS. 3.3 Preparing the Colon for Histological Assessment

Colon tissue can be fixed with 10% buffered formalin for 24–48 h and tissue sections may then be stained for Hematoxylin and eosin or ki67 to assess histological damage and proliferation in the colon.

3.4 Assessment of Colitis Induction in Treated Mice

Assessments of disease activity index and intestinal permeability can be made depending on experimental goals. 1. Disease activity index (DAI): DAI is used to evaluate the grade and extent of the intestinal inflammation using a previously established scoring system [15–17]. Feeding mice with DSS in drinking water leads to the development of inflammation in the colon with signs of weight loss, watery diarrhea, fecal bleeding, and infiltration of inflammatory granulocytes [17, 18]. To assess the extent of colitis, body weight, stool consistence, and blood in the stool are monitored daily (see Table 1) (see Note 8). The score of DAI ranges from 0 to 12 (total score). 2. Measurement of intestinal permeability: Intestinal permeability can be measured during the recovery period after the DSS treatment. FITC-labeled dextran is used as an indicator to examine the permeability of the intestines. Mice is given fluorescein isothiocyanate (FITC)-dextran by oral gavage at a dose of 0.5 mg/g of body weight. Four hours later, mice are bled, and FITC-dextran was quantified in the serum via a fluorescence spectrophotometer. 3. Histological grading of colitis: Histological grading on the severity of inflammation is performed with the H&E staining

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Table 1 Disease activity index (DAI) Score

Weight loss

Stool consistency

Blood in the stool

0

None

Normal

Normal

1

1–5%

2

5–10%

Very soft

Slight bleeding

3

10–20%

4

>20%

Watery diarrhea

Gross bleeding

Table 2 Histology grading (degree of inflammation) Score

Degree of inflammation

0

No inflammation

1

Mild inflammation or prominent lymphoid aggregates

2

Moderate inflammation (mucosa)

3

Moderate inflammation associated with crypt loss (submucosa)

4

Severe inflammation with crypt loss and ulceration (submucosa)

Table 3 Histology grading (degree of crypt damage) Score

Crypt damage

0

No destruction

1

1–33% of crypts destroyed

2

34–66% of crypts destroyed

3

67–100% of crypts destroyed

method and histological scoring of colonic tissue using previously published results [18, 19]. Administration of DSS in mice results in severe macroscopic and histological damage in the colon. The degree of inflammation was scored as follows: (0) no inflammation, (1) mild inflammation or prominent lymphoid aggregates, (2) moderate inflammation, (3) moderate inflammation associated with crypt loss, and (4) severe inflammation with crypt loss and ulceration. Crypt destruction was graded as follows: (0) no destruction, (1) 1%–33% of crypts destroyed, (2) 34%–66% of crypts destroyed, and (3) 67%– 100% of crypts destroyed. The individual scores from inflammation and crypt damage were summed to derive histological

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score for colonic inflammation (maximum score 7) (see Tables 2 and 3). 3.5 Assessment of Inflammation

Assessment of inflammation can be performed at the end of the First, second, or third DSS cycle depending on experimental goals. 1. Ex vivo colon culture: Cut open the colon longitudinally and wash four times in HBSS supplemented with penicillin and streptomycin. Cut the colon laterally into 1-cm segments and culture each piece in 24-well flat-bottom culture plates in serum-free RPMI 1640 medium supplemented with penicillin and streptomycin, l-glutamine, and nonessential amino acids. After 24–48 h, collect the supernatant and centrifuge at 13,000  g for 10 min at 4  C and store at 20  C for assaying pro-inflammatory and anti-inflammatory cytokines by ELISA (see Fig. 2). 2. Tissue myeloperoxidase (MPO) assay: MPO activity can be used as a marker of inflammation. MPO is a marker for neutrophils and helps in assessing the neutrophil influx into inflamed tissue [20]. Take 50 mg of colon tissue in a 2 mL microcentrifuge tube containing 1 mL of PBS for 30 s. Spin the homogenate at 20,400  g for 5 min. Decant the supernatant without disturbing the pellet. Suspend the pellet in 1 mL of 50 mM PBS (pH 6.0) containing 0.5% hexadecyltrimethylammonium bromide. The sample is freeze-thawed three times, sonicated (90 s), and centrifuged at 20,400  g for 5 min. Incubate the sample in water for 120 min at 60  C. Centrifuge the sample at 20,400  g for 5 min. MPO is assayed in the clear supernatant in a 96-well plate in triplicates. Add 10 μL of supernatant (in triplicates) to 200 μL of 1 mg/mL of dianisidine dihydrochloride and 5  10–4% H2O2. The change in optical density is measured at 450 nm three times at 30 sec intervals using a spectrophotometer. Use Human neutrophil MPO (Sigma) as a standard. One unit of MPO activity is defined as the amount that degraded 1.0 μmol of peroxide/min at 25  C (see Note 10). 6. Isolation of leucocytes from colon: Cut open the colon longitudinally and clean fecal contents. Wash in HBSS three times. Cut the colon laterally into small (1 cm) pieces and place into a 50 mL conical tube containing 10–15 mL of CMF/HEPES and agitate gently. Filter the content using a single mesh wire strainer. Repeat this step for 6 times to remove the mucus and rest of fecal content. Transfer the washed pieces into another 50 mL conical tube. Add 20 mL complete RPMI/collagenase  and incubate for 30 min at 37 C. After 15 min, pipette the tissue up and down. Pour the content of each tube through a fresh 100 μm cell strainer into a 50 mL conical tube and save

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Fig. 2 Colonic inflammatory cytokine levels in response to AOM-DSS treatment. Severity of CAC is associated with higher levels of inflammatory cytokines in the colonic environment. (a) RNA was extracted from the colons of untreated and AOM-DSS-treated mice. Expression of pro-inflammatory genes was quantified by qPCR. (b) Excised colon samples of untreated and AOM-DSS-treated mice were cultured for 48 h ex vivo, and the pro-inflammatory cytokine levels in the culture supernatants were quantified by ELISA. The error bars indicate mean  SEM of 5–6 mice/group

it. Transfer pieces back into a tube with fresh RPMI/collagenase and incubate again as described above. Pool the filtrate in the 50 mL conical tubes and spin 10 min at 252  g. Discard the supernatant. Wash the pellet once with RPMI and spin 10 min at 252  g (see Note 11). Suspend cells in 7 mL 40% Percoll in sterile PBS, underlay with 5 mL 80% Percoll and spin 20 min at 363  g at room temperature with no brake. Collect cells from the interface in a 15 mL tube and fill will RPMI-10. Vortex and spin for 10 min at 252  g. Discard the supernatant and wash one more time with RPMI (see Note 12). Cells can be used for enrichment of specific subsets of immune cells, gene expression analysis, and intracellular staining for cytokines. 3. Immune cell stimulation and Intracellular staining for cytokines: Stimulate isolated lymphocytes with PMA (50 ng/mL) and ionomycin (500 ng/mL) for 6 h in the presence of Brefeldin A and monensin for 6 h. The cells are centrifuged at 252  g for 5 min. Remove the supernatant and resuspend the pellet in 0.5 mL of FACS buffer. 100 μL cell suspension (1–3 million or at least 300–500,000 cells) is added to each well in a 96-well plate or 1.5 mL tube for each leukocyte and isotype control (depending on the number of leukocyte markers stained). The suspension is centrifuged and the supernatant discarded. Then, the cells can be surface stained for CD4+ T cells (CD45+ I-Ab–CD4+), CD8+ T cells (CD45+ I-Ab– CD8+), DCs (CD45+ I-Ab+ CD11c+ CD64 ), or macrophages (CD45+ I-Ab+ CD11b+ CD64+) followed by intracellular staining of IFN-γ, IL-17A, TNF-α, and IL-10 for CD4+ and CD8+ T cells, and IL-6, pro-IL-1β, TNF-α, and IL-10 for DCs and macrophages (see Note 13). The samples are analyzed

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immediately by flow cytometry or stored in the dark at 4 C for later analysis (24–48 h). 3.6 Assessment of CAC Following AOM/ DSS Treatment

1. Endoscopy (optional). Endoscopy can be performed at the end of the second and third cycle to confirm the tumor growth in vivo prior to sacrifice (see Note 14). 2. Measurement of tumor burden and size. Place the cleaned and longitudinally cut and opened colon with the luminal side facing upwards under the dissection microscope. Identify the tumors along the length of the colon and record the number of tumors. Tumor size can be measured using digital calipers (see Note 15). 3. RNA isolation and Real-time PCR. Depending on the experimental goals, if desired, tumor tissue or adjacent tissue can be harvested for RNA or protein analysis using a scalpel. Total mRNA can be isolated from colon or cells using the Total RNA Kit according to the manufacturer’s protocol. cDNA is generated using an RNA to cDNA Ecodry Premix Kit according to the manufacturer’s protocol. Subsequently, quantitative realtime PCR is done using SYBR Green Master Mix with genespecific primers [21] and a MyiQ5 ICycler, and with gene expression normalized relative to Gapdh.

3.7 Representative Results and Discussion

Assessments of weight loss, DAI, intestinal permeability, and histological grading are the most important parameters for determining the severity of colitis and CAC. Mice treated with AOM-DSS may have significantly worsened disease compared to the control mice with water. AOM-DSS-treated animals will show a marked increase in weight loss (see Fig. 3a) and marked reduction in colon length compared to the controls (see Fig. 3b). MPO activity can be used as a surrogate marker for colitis severity. MPO activity, a hallmark of colonic inflammation, is markedly increased in the colons of mice after AOM-DSS treatment (see Fig. 3c). An increase in FITCdextran in the serum of AOM-DSS-treated mice is observed after oral gavage (see Fig. 3d), indicating severe impairment in the integrity of the intestinal epithelial barrier. Consistent with enhanced gut inflammation, mice will have increased tumor burden in response to AOM-DSS treatment (see Fig. 4a). This is associated with a significant increase in tumor numbers and tumor load in the distal colon (see Fig. 4b, c). Histopathological analysis of colonic sections from AOM-DSS-treated mice would reveal a marked increase in damage to mucosa with epithelial erosion, frequent ulceration, and loss of crypt structure (see Fig. 4d). In line with this observation, mice treated with AOM-DSS will show a marked increase and histological score compared to the controls treated with water only (see Fig. 4e). Furthermore, immunohistochemistry with Ki67

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Fig. 3 Assessment of colonic inflammation in response to AOM-DSS treatment. (a) Change in body weight and (b) colon length of mice treated with AOM-DSS. (c) Myeloperoxidase activity in colon on day 9. (d) Mice were fed with FITC-dextran on day 9, and 4 h later FITC-dextran was quantified in serum. The error bars indicate mean  SEM of 5–6 mice/group

Fig. 4 Assessment of CAC following AOM-DSS treatment. (a) Representative images of colon polyps as induced by AOM-DSS in mice. (b) Average polyp numbers and (c) polyp size (mm) from mice treated with AOM-DSS. (d) Representative colon sections from mice treated with AOM-DSS were microscopically analyzed for tissue damage and infiltration of inflammatory cells after staining with hematoxylin and eosin (H&E) staining. Scale bar, 200 μm. (e) Histopathological score (degree of inflammation plus crypt damage) of colons was graded following analysis of H&E-stained cross-sections of colons of AOM-DSS-treated mice. (f) Representative colon sections from mice treated with AOM-DSS and assessed for proliferating cells after staining with Ki67 antibody. Scale bar, 200 μm. (g) Ki67-positive cells were manually counted in colon sections (f) of mice treated with AOM-DSS. The error bars indicate mean  SEM of 5–6 mice/group

would reveal that increased tumor numbers in the colons is associated with markedly increased cell proliferation (see Fig. 4f, g). The higher levels of pro-inflammatory cytokines such as IL-6, IL-1β, and TNF-α present in the gut microenvironment promotes colonic inflammation and CAC [22, 23]. Consistent with the tumor

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burden, the colons from mice treated with AOM-DSS would express significantly higher levels of pro-inflammatory cytokines such as IL-1β, IL-6, and TNF-α compared to the colons from WT mice treated with water (see Fig. 2a, b). Dysfunctional APCs play an important role in the pathogenesis of IBD and colitisassociated colorectal cancer (CAC) and are the major producers of pro-inflammatory factors. Moreover, APCs control both T cell differentiation and expansion through the secretion of various inflammatory and anti-inflammatory cytokines. A delicate balance between regulatory T cells versus pathological effector T cells underlies disease progression in many inflammatory diseases, including IBD and CAC. Recent evidences suggest that IL-17A-producing CD4+ T (Th17) cells play an important role in the pathogenesis of IBD and CAC [24]. In contrast, regulatory T cells such as Foxp3+ Tregs and IL-10-producing Tr1 cells play a pivotal role in controlling inflammatory conditions and maintaining immune tolerance in the intestine [25, 26]. IL-6 and IL-1β induce the differentiation of naı¨ve CD4+ T cells to Th17 cells. Thus, phenotypic and functional characterization of these immune cells in the colon can be performed using this model system. Another major advantage is that this model helps in understanding the role of both innate and adaptive immune cells in both induction and progression of inflammation and inflammation-associated colon cancer. In summary, the AOM-DSS-induced model of CAC closely recapitulates histological, pathological, and molecular features of IBD-associated colon cancer in humans.

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Notes 1. Conditions need to be standardized for each particular strain, animal facility, and environmental variability such as microflora, diet, and other environmental conditions. Thus, we recommend using control and experimental mice that are littermates. This will control for any environmental differences, including microbiota variability. Otherwise, we strongly recommended that the control and experimental mice be housed in the same room. Based on our experience, mice from both sexes are susceptible to AOM/DSS-induced CAC. 2. Prepare DSS with animal facility water. Mice do not drink distilled water. It is important to use DSS of specified molecular weight. Prepared DSS solution can be kept refrigerated for up to 1 week. DSS from different manufacturers and different lots may yield different results. We recommend using DSS from the same lot and the same manufacturer. 3. o-Dianisidine dihydrochloride solution should be prepared fresh when performing the assay.

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4. AOM concentration is an important variable to adjust, and this depends on the mouse strain and animal facility [10]. We recommend performing a trial experiment with different dosage of AOM between 7.5 and 12.5 mg/kg doses. 5. DSS concentration is an important variable to adjust, and this depends on the mouse strain and animal facility. If a particular strain is very sensitive to 3.0% DSS, then we recommend using a reduced concentration range between 1 and 2% DSS. Thus, dosage and duration of DSS treatment need to be optimized for each strain. 6. DSS should also be prepared fresh for each cycle of administration and the solution should be replaced every 2–3 days during the treatment period. 7. AOM is a volatile genotoxic agent and should be handled carefully according to its MSDS. Stock solutions should be  prepared in a chemical hood and can be kept in 20 C up to 1 year. Stock solutions of AOM should not be used after repeated freeze-thaw cycles. 8. Body weight should be used as an indicator of overall animal health. If an animal loses more than 15% of their initial body weight, they may require early euthanasia (follow all appropriate IACUC protocols). 9. Scarification of AOM-DSS-treated mice can be delayed further (weeks to months) for larger tumors. 10. MPO activity assay should be performed within the first week of tissue collection. Activity decreases over time. 11. RPMI/collagenase should be prepared fresh from the stock when performing the isolation. Each batch of collagenase should be tested for toxicity and activity. 12. If the cell suspension clumps together, increase the concentration of DNase I. 13. FACS buffer should be prepared fresh every time when performing the assay. 14. Only a few mice should be evaluated using endoscopy, as it has the potential to disrupt tumors in the colon. This step is generally not necessary. 15. Tumors are also easily visible when placed on a dark surface. Alternatively, 1% Alcian blue dye can be applied to identify the tumors. Digital photography of the gross colon specimens will also be helpful for precise analysis of tumor burden (see Fig. 3a).

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Acknowledgments Funding provided by NIH (R01DK097271, R01DK123360) and Augusta University institutional development funds. References 1. Sands BE (2007) Inflammatory bowel disease: past, present, and future. J Gastroenterol 42 (1):16–25 2. Fiocchi C (1998) Inflammatory bowel disease: etiology and pathogenesis. Gastroenterology 115(1):182–205 3. Terzic J, Grivennikov S, Karin E, Karin M (2010) Inflammation and colon cancer. Gastroenterology 138(6):2101–2114. e5 4. Kim ER, Chang DK (2014) Colorectal cancer in inflammatory bowel disease: the risk, pathogenesis, prevention and diagnosis. World J Gastroenterol 20(29):9872–9881 5. Dyson JK, Rutter MD (2012) Colorectal cancer in inflammatory bowel disease: what is the real magnitude of the risk? World J Gastroenterol 18(29):3839–3848 6. Price AB, Morson BC (1975) Inflammatory bowel disease: the surgical pathology of Crohn’s disease and ulcerative colitis. Hum Pathol 6(1):7–29 7. Duchmann R, Kaiser I, Hermann E, Mayet W, Ewe K, Meyer zum Buschenfelde KH (1995) Tolerance exists towards resident intestinal flora but is broken in active inflammatory bowel disease (IBD). Clin Exp Immunol 102 (3):448–455 8. Powrie F (1995) T cells in inflammatory bowel disease: protective and pathogenic roles. Immunity 3(2):171–174 9. Strober W, Fuss IJ, Blumberg RS (2002) The immunology of mucosal models of inflammation. Annu Rev Immunol 20:495–549 10. Rosenberg DW, Giardina C, Tanaka T (2009) Mouse models for the study of colon carcinogenesis. Carcinogenesis 30(2):183–196 11. De Robertis M, Massi E, Poeta ML, Carotti S, Morini S, Cecchetelli L, Signori E, Fazio VM (2011) The AOM/DSS murine model for the study of colon carcinogenesis: from pathways to diagnosis and therapy studies. J Carcinog 10:9 12. Perse M, Cerar A (2012) Dextran sodium sulphate colitis mouse model: traps and tricks. J Biomed Biotechnol 2012:718617 13. Sussman DA, Santaolalla R, Strobel S, Dheer R, Abreu MT (2012) Cancer in inflammatory bowel disease: lessons from animal

models. Curr Opin Gastroenterol 28 (4):327–333 14. Ostanin DV, Bao J, Koboziev I, Gray L, Robinson-Jackson SA, Kosloski-Davidson M, Price VH, Grisham MB (2009) T cell transfer model of chronic colitis: concepts, considerations, and tricks of the trade. Am J Physiol Gastrointest Liver Physiol 296(2):G135–G146 15. Axelsson LG, Landstrom E, Goldschmidt TJ, Gronberg A, Bylund-Fellenius AC (1996) Dextran sulfate sodium (DSS) induced experimental colitis in immunodeficient mice: effects in CD4(+) -cell depleted, athymic and NK-cell depleted SCID mice. Inflamm Res 45 (4):181–191 16. Egger B, Bajaj-Elliott M, MacDonald TT, Inglin R, Eysselein VE, Buchler MW (2000) Characterisation of acute murine dextran sodium sulphate colitis: cytokine profile and dose dependency. Digestion 62(4):240–248 17. Laukoetter MG, Nava P, Lee WY, Severson EA, Capaldo CT, Babbin BA, Williams IR, Koval M, Peatman E, Campbell JA et al (2007) JAM-A regulates permeability and inflammation in the intestine in vivo. J Exp Med 204(13):3067–3076 18. Azuma YT, Matsuo Y, Kuwamura M, Yancopoulos GD, Valenzuela DM, Murphy AJ, Nakajima H, Karow M, Takeuchi T (2010) Interleukin-19 protects mice from innatemediated colonic inflammation. Inflamm Bowel Dis 16(6):1017–1028 19. Okayasu I, Hatakeyama S, Yamada M, Ohkusa T, Inagaki Y, Nakaya R (1990) A novel method in the induction of reliable experimental acute and chronic ulcerative colitis in mice. Gastroenterology 98(3):694–702 20. Krawisz JE, Sharon P, Stenson WF (1984) Quantitative assay for acute intestinal inflammation based on myeloperoxidase activity. Assessment of inflammation in rat and hamster models. Gastroenterology 87(6):1344–1350 21. Suryawanshi A, Manoharan I, Hong Y, Swafford D, Majumdar T, Taketo MM, Manicassamy B, Koni PA, Thangaraju M, Sun Z et al (2015) Canonical wnt signaling in dendritic cells regulates Th1/Th17 responses and suppresses autoimmune neuroinflammation. J Immunol 194(7):3295–3304

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22. Francescone R, Hou V, Grivennikov SI (2015) Cytokines, IBD, and colitis-associated cancer. Inflamm Bowel Dis 21(2):409–418 23. Saleh M, Trinchieri G (2011) Innate immune mechanisms of colitis and colitis-associated colorectal cancer. Nat Rev Immunol 11 (1):9–20 24. De Simone V, Pallone F, Monteleone G, Stolfi C (2013) Role of TH17 cytokines in the control of colorectal cancer. Onco Targets Ther 2 (12):e26617

25. Zheng T, Zhang B, Chen C, Ma J, Meng D, Huang J, Hu R, Liu X, Otsu K, Liu AC et al (2018) Protein kinase p38alpha signaling in dendritic cells regulates colon inflammation and tumorigenesis. Proc Natl Acad Sci U S A 115(52):E12313–E12E22 26. Ahn J, Son S, Oliveira SC, Barber GN (2017) STING-dependent signaling underlies IL-10 controlled inflammatory colitis. Cell Rep 21 (13):3873–3884

Chapter 11 Generation of Colon Cancer Model Based on Colonoscopy Injection Suhwan Chang Abstract Colorectal cancer animal model is a very useful tool to explore the tumor initiation and development. In the past year, many methods have been used for building up the mouse model including the subcutaneous injection and cecal wall injection or implantation. But this model cannot reflect the native stromal environment of the colon mucosa. Recently, the in vivo murine endoscopy has been developed allowing high-resolution imaging of the colon. Endoscopy orthotopic injection tumor cell line becomes a low cost, fast tumor growth simple technique. In this chapter, we describe detailed protocols for rapidly and efficiently building up colon cancer tumors model by using the colonoscopy-guided mucosal injection. This model can be used to explore drug testing, gene function assessment, and cancer metastasis. Key words Colon cancer, Mouse model, Colonoscopy, Cell injection

1

Introduction Colorectal cancer (CRC) is the third most common type of cancer with a global incidence of over 1.3 million cases per year [1]. Despite recent therapeutic progress, the prognosis of CRC is still poor, and it remains one of the reasons for cancer-related death in the world. However, observational studies in CRC populations provide limited insights into disease biology. Compared with the in vitro studies, the CRC small animal models can offer the ability to study tumor initiation, tumor–stromal interactions, and pathophysiologic mechanisms of CRC development and more accurate results for the preclinical assessment of novel therapeutic modalities [2]. Many murine CRC models have been reported as before [3, 4]. The main approaches include spontaneous tumors developing by mutant mice, treatment with a wide range of carcinogenic agents, and ectopic or orthotopic implantation of tumor cells. Each kind of models has its advantage and disadvantage. More and more researchers focus on the orthotopic model and PDX (patients-

Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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derived xenograft) model. The PDX models offer advantages which have been shown to stably retain molecular, genetic, and histopathological features of their originating tumors. But for the CRC model, the PDX model application in research remains severely limited by high cost, the complexity of the surgical procedure, and procedure-related inflammation and mortality [5]. Additionally, most of the used CRC PDX model is generated by subcutaneous or colon serosal implantation of fresh, surgically derived human tumor fragment. But the tumor implanted serosal colon grows outside of the colon, which is not a real orthotopic model as it cannot mimic the luminal tumor environment. In spite of the developing tumors formed after transplantation of cell lines often poorly reflect the histopathological features of human CRC, it is still the most commonly used mouse model in basic and translational oncology [6]. Low cost, fast tumor growth and simple technique make the application of these models widely used [5]. Moreover, endoscopy orthotopic injection tumor cell line enables researchers to determine the site of tumor growth and more accurate simulation of the tumor’s microenvironment [6, 7]. Therefore, orthotopic implantation models are invaluable for the evaluation of new diagnostic approaches. Recently, the in vivo murine endoscopy has been developed allowing high-resolution imaging of the colon in living mice and enabling researchers to directly visualize and score pathologic tissue changes in the colon [2, 8–11]. It has become a new powerful tool for CRC orthotopic mouse model building up and standard endoscopy can easily be performed by a trained researcher.

2

Materials

2.1

Cell Line

Mouse-derived colon cancer cell line (such as CT-26) or humanderived colon cancer cell line (such as HCT-116) can be chosen based on the experimental needs.

2.2

Animals

Ideal recipient mice for colonoscopy-guided mucosal injection is 6to 8-weeks old. Male mice or female mice are all acceptable. Mouse strains are chosen according to the experimental design. Orthotopic engraftment of human-derived tumor cell line is generally performed in nude mice or NOD scid gamma (NSG) mice. C57BL/6 mouse usually is used for mouse-derived cell line mucosal injection.

2.3

Equipment

2.3.1 Cell Culture and Cell Preparation for Injection

1. Cell culture media (RPMI-10% FBS supplemented with antibiotics) and PBS. 2. Culture plates, 100 mm. 3. Pipettes and pipettes tips, 10, 200, and 1000 μL. 4. Tubes, 1.5 mL. 5. Biosafety level 2 (BSL2) cabinet.

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6. Cell culture incubator set to 5% CO2, 37  C. 7. Inverted-phase microscope. 8. Centrifuge. 2.3.2 ColonoscopyGuided Mucosal Injection

1. Oral gavage needle. 2. Heating pad; PBS or osmotic solution. 3. Eye ointment and lidocaine glue. 4. Sterile isotonic saline or PBS. 5. Mucosal injection needle (30–33 gauge, 16-inches long, 45-degree bevel) and syringe. 6. Optical colonoscopy system (Karl Storz) including Hopkins Telescope. 7. Professional Image Enhancement System; HD camera system (Karl Storz) with examination sheath and operating sheath. 8. Cold light source.

3

Methods

3.1 Cell Preparation Procedure

1. Before 5 days of the tumor implantation, thaw the tumor cell lines and maintain 5% CO2 at 37 uC in high glucose RPMI medium supplemented with 10% fetal bovine serum and 1% penicillin streptomycin antibiotic solution. 2. According to the injection number of the mouse, calculate the cells as 1–3  105 contain in 50 μL suspension for one mouse. Prepare cells and keep the cells in ice before the injection.

3.2 Colonoscopy Preparation and Staff Member

1. Attach the air pump, light source cable, and camera head to the colonoscope. 2. Sterilize the endoscope, clean the telescope lens with lens paper, and set the focus so that objects at a distance of 3–5 mm give a crisp picture. 3. Two people are required for the mucosal injection procedure.

3.3 Animal Preparation Procedure

1. Place the mouse in a supine position on a heating pad and anesthetize mouse with 2.0% isoflurane. 2. Adjust the isoflurane level so that the mouse is breathing 1 breath per second (most used was 0.2 L/min). 3. Apply a small amount of eye ointment to the eyes of the mouse to protect the corneas. 4. Flush the colon with the intrarectal application of PBS (2–4 mL) to stimulate bowel cleansing by using an oral gavage needle (giving gentle abdominal pressure would be helpful to force stool pellets out).

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3.4 Implantation Procedure

1. Apply lidocaine glue to the anus of the mouse. 2. Carefully push the endoscope into the rectum as far as possible under visual control to ensure colonic mucosa was healthy and unaltered before the procedure starts. Usually, with a rigid scope, it can be inserted up to 3.5 cm and then the colon makes a flexure and is difficult (dangerous) to pass this part. 3. The ideal injection location should be proximally 2–3 cm from the anal verge. This site has enough space and allows us to give additional injections if need. 4. After determining the injection location, one investigator controls the endoscopy and the second investigator inserts the needle. A flexible 30 G injection needle was introduced through the working channel of the endoscope and colonic mucosa was gently penetrated using the attached 30 G needle with its bevel facing the lumen. 5. The best way is the needle parallel to the colonoscope with a 30 angle relative to the colon wall. 6. Keep the endoscope close to the colon wall so that you can see the needle entering the mucosa. 7. For tumor cell injection, a small bubble formation with overlying mucosa indicated the successful implantation. The second investigator should quickly inject the remaining volume and the bulb becomes big and fill most of the colon lumen. 8. Keep the needle in the bubble for at least 10 s to prevent leakage of the sample from the injection site. 9. Finally, remove the endoscope and put the mouse back into the cage until it recovers. During the procedure, do not forget to record the useful image by DV or picture.

3.5 Tumor Follow-Up with Colonoscopy

1. For most mouse models, tumors can be visualized with colonoscopy 1–2 weeks after tumor cells injection according to the injection cells number (see Fig. 1). 2. The tumor can be evaluated by two criteria. The first one is the tumor initiation rate and the second is tumor size judgment. Recommend using a tumor scoring in vivo in mice according to tumor occupying the lumen which has been published [9].

4

Additional Tips 1. Gently clear the intestinal feces by injection PBS and with massage. If the feces did not discharge, repeat injection PBS or wait 15–20 min. 2. Carefully control the air pump. Adjust the air pump pressure by putting the scope into water. One small bubble per second is

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Fig. 1 (a) The procedure of endoscopy-guided tumor cells injection. You can see the bulb formation (blue arrow) and the needle (black arrow) located in the right side. (b) Tumor located in the right side and occupied 50% of colon lumen (yellow arrow). (c, d) The histology image by H&E staining and the tumor located between the mucosa and muscle layer

ideal. If the air pressure too high, it will lead to excessive intestinal filling and harm the colon. Also, this will increase the perforation rate during the operation process. 3. If the first injection fails, adjust the needle in a new location at least 0.5 cm away from the prior injection site. Do not repeat puncture at the same site many times. 4. If a colonic stent is placed after tumor growth, the ideal injection site should be proximally 2–2.5 cm from the anal verge. Because we need to leave enough space for stent placement. 5. The injection needle tip should always be visible. Otherwise, the injection failure rate and perforation rate will increase.

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6. For the follow-up colonoscopy, do not try to pass the tumor especially when the tumor is very large. It is very easy to cause perforation.

Acknowledgments This research was supported by an internal grant of the University of Ulsan, College of Medicine (Physician Scientist Program) and a grant from Asan Institute for Life Sciences (grant No.2019 IP0571). References 1. Siegel RL, Miller KD, Jemal A (2018) Cancer statistics, 2018. CA Cancer J Clin 68:7–30 2. Roper J, Tammela T, Akkad A, Almeqdadi M, ¨ H (2018) Santos SB, Jacks T, Yilmaz O Colonoscopy-based colorectal cancer modeling in mice with CRISPR-Cas9 genome editing and organoid transplantation. Nat Protoc 13:217–234 3. Rosenberg DW, Giardina C, Tanaka T (2009) Mouse models for the study of colon carcinogenesis. Carcinogenesis 30:183–196 4. Taketo MM, Edelmann W (2009) Mouse models of colon cancer. Gastroenterology 136:780–798 5. Gengenbacher N, Singhal M, Augustin HG (2017) Preclinical mouse solid tumour models: status quo, challenges and perspectives. Nat Rev Cancer 17:751–765 6. Achyut BR, Yang L (2011) Transforming growth factor-beta in the gastrointestinal and hepatic tumor microenvironment. Gastroenterology 141:1167–1178 7. Suganuma M, Watanabe T, Yamaguchi K, Takahashi A, Fujiki H (2012) Human gastric

cancer development with TNF-α-inducing protein secreted from helicobacter pylori. Cancer Lett 322:133–138 8. Zigmond E, Halpern Z, Elinav E, Brazowski E, Jung S, Varol C (2011) Utilization of murine colonoscopy for orthotopic implantation of colorectal cancer. PLoS One 6:e28858 9. Becker C, Fantini MC, Neurath MF (2006) High resolution colonoscopy in live mice. Nat Protoc 1:2900–2904 10. Neurath MF, Wittkopf N, Wlodarski A, Waldner M, Neufert C, Wirtz S, Gu¨nther C, Becker C (2010) Assessment of tumor development and wound healing using endoscopic techniques in mice. Gastroenterology 139:1837–1843. e1831 11. Nakanishi M, Me´noret A, Belinsky GS, Giardina C, Godman CA, Vella AT, Rosenberg DW (2007) Utilizing endoscopic technology to reveal real-time proteomic alterations in response to chemoprevention. Proteomics Clin Appl 1:1660–1666

Chapter 12 Generation of Transgenic Fluorescent Reporter Lines for Studying Hematopoietic Development in the Mouse Jeffrey Barminko, Andrei M. Vacaru, and Margaret H. Baron Abstract Hematopoiesis in the mouse and other mammals occurs in several waves and arises from distinct anatomic sites. Transgenic mice expressing fluorescent reporter proteins at various points in the hematopoietic hierarchy, from hematopoietic stem cell to more restricted progenitors to each of the final differentiated cell types, have provided valuable tools for tagging, tracking, and isolating these cells. In this chapter, we discuss general considerations in designing a transgene, survey available fluorescent probes, and describe methods for confirming and analyzing transgene expression in the hematopoietic tissues of the embryo, fetus, and postnatal/adult animal. Key words Hematopoiesis, Transgenic mice, Knock-in, Green fluorescent protein, Fluorescent reporter

1

Introduction Since the discovery and cloning of wild type green fluorescent protein (wtGFP) from the bioluminescent jellyfish Aequorea victoria [1] and the subsequent creation of spectral variants [2–4], fluorescent protein FPs have become indispensable for imaging cellular differentiation and function at high resolution and in real time (reviewed in refs. [3, 4]). Gene-specific regulatory elements can be used to drive targeted expression of FP reporters, with spatial- and/or temporal specificity, virtually in any cell type of a transgenic animal. The hematopoietic system has benefitted enormously from this approach, which made it possible to explore the emergence, expansion, migration, and differentiation of progenitors for the erythroid, myeloid, and lymphoid lineages. It is now possible to label and track the development of distinct hematopoietic cell types in vivo and to isolate these cells directly, using

Jeffrey Barminko and Andrei M. Vacaru are Co-first authors. Shree Ram Singh et al. (eds.), Mouse Genetics: Methods and Protocols, Methods in Molecular Biology, vol. 2224, https://doi.org/10.1007/978-1-0716-1008-4_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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fluorescence-activated cell sorting (FACS). In this chapter, which has been updated from an earlier version [5], we discuss the steps involved in the generation and analysis of transgenic lines in which fluorescent reporters are expressed in hematopoietic lineages of the mouse, the most genetically tractable model for mammalian development. 1.1 Ontogeny of the Mouse Hematopoietic System

Hematopoiesis is a precisely orchestrated, stepwise process that leads to the formation of all lineages of the blood [6]. Primitive erythroid cells (EryP) are the first hematopoietic lineage to be detected in the mouse embryo (reviewed in ref. [7]). They are generated late in gastrulation, in the blood islands of the yolk sac (YS), along with macrophages and megakaryocytes [8]. The first definitive hematopoietic cells, comprising erythroid, megakaryocytic, and myeloid lineages, also arise in the YS, shortly after the appearance of EryP (reviewed in refs. [7, 8]). Hematopoietic stem cells (HSCs) form in the aorta-gonad-mesonephros (AGM) region of the embryo, in the large arteries and placenta, and, very likely, in the YS (for a review, see ref. [7]). HSCs do not differentiate in these sites but instead seed the fetal liver (FL), where they expand and produce progenitors that give rise to definitive erythro-myeloid and lymphoid lineages. Late in gestation, HCSs migrate from the fetal liver to the bone marrow, which becomes the main blood production center in the postnatal animal [9]. The general hierarchy of hematopoietic development is shown as a “snapshot” in ref. [10].

1.2 General Considerations in Designing a Transgene

Transgenic mouse lines expressing fluorescent proteins (FPs) are invaluable tools for studying the development of the hematopoietic system. Careful design is necessary to achieve the desired expression of the reporter protein. The main components of a fluorescent reporter transgene are the promoters (and, usually, other upstream regulatory sequences), sequences encoding the fluorescent protein, and splice/polyadenylation signals (see Fig. 1). A cartoon outlining the most commonly used approach for creation of a transgenic mouse line is presented in Fig. 2.

1.2.1 Regulatory Elements

The promoter is the region of a gene from which mRNA transcription is initiated and is essential for controlling both the spatial and temporal expression of a transgene. A number of hematopoieticspecific promoters have been used successfully to drive expression of fluorescent reporters in different hematopoietic lineages (see Table 1). The transgene construct should include a translational start codon (ATG), a Kozak sequence either upstream from or coupled to the start codon [11], and a translational stop codon (see Fig. 1). Posttranscriptional regulatory elements may be included to enhance mRNA stability (e.g., see ref. [12]).

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Fig. 1 Basic design of a fluorescent transgenic construct. “R” marks the restriction enzyme sites used for the removal of the bacterial backbone before transgene injection. s/pA represents the splicing and polyadenylation signals. A stop codon (denoted by “X”) should also be included in the construct design. The enhancer may be positioned upstream or downstream from the promoter and sequences encoding the FP and regulatory signals. For additional details, see text

Additional regulatory elements should accompany the promoter to drive the desired transgene expression pattern. The most commonly used regulatory elements are enhancers or other upstream regulatory elements [13], an intron, which provides splice donor and acceptor sites, and a polyadenylation signal (see Fig. 1). The intron may be taken from the same gene as the promoter. Low transgene expression may be significantly increased through the use of a generic intron [14]. It is well documented that prokaryotic sequences in the vector perturb the frequency and extent of transgene expression [15– 17]. Therefore, restriction enzyme sites flanking the transgene should be included to allow the removal of the vector backbone before microinjection. Following microinjection into the male pronucleus of a fertilized egg (see Fig. 2), the transgene is inserted randomly, and often in multiple copies, into the genome [17]. The neighboring chromatin may influence expression of the transgene, leading to undesired effects such as ectopic expression or even silencing [17, 18]. To avoid these effects, chromatin insulators can be used. These DNA elements, together with the proteins that bind to them, impair interactions with neighboring chromatin [19]. Enhancers are used to stimulate transcription and may do so in an

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Fig. 2 Basic steps in the generation of a fluorescent transgenic reporter mouse line. Pronuclear injection of the transgene is shown in this cartoon, but transgenic mouse lines can also be generated by blastocyst injection or embryo aggregation with genetically modified ES cells (see text)

orientation- and position-independent manner [13]. Locus control regions (LCRs) from globin or CD4 genes have been used as enhancers for transgene expression in erythroid- or T cells, respectively [13, 20–22]. For example, a minimal human ε-globin promoter combined with a truncated human β-globin LCR, a regulatory element that controls the erythroid-specific expression of all cis-linked globin genes [23], have been used to generate mouse lines expressing GFP or CFP in the primitive erythroid lineage [24–26] (see Fig. 3a). We recommend that expression of a newly designed transgene be tested in cultured cells before moving forward with the generation of the transgenic mouse line. This precaution helps to ensure that the transcriptional regulatory elements are functional and that the reporter sequence is translated to a functional protein. 1.2.2 Insertion of Exogenous DNA into the Genome

The most commonly used approaches for generation of fluorescent reporter mice are microinjection of DNA (plasmid or BAC; see below) into a fertilized egg (see Fig. 2) and targeted insertion (“knock-in”). Insertion of a transgene into the mouse genome

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Table 1 Transgenic mouse lines expressing fluorescent reporter proteins in the hematopoietic system Gene

Reporter

Lineage labeled

Reference

Gata2

GFP (KI)

HSC

[77]

Hoxb4

YFP (KI)

HSC

[78]

Ly-6A

GFP (Tg)

HSC

[79]

a

Bmi1

GFP (KI)

HSC

[80]

Abcg2

IRES-GFP (KI)

HSC, erythroid

[81]

c-Kit

GFP (Tg)

HSC, progenitors

[82]

c-Kit

GFP (CKO)

HSC, progenitors

[83]

Runx1

GFP (Tg)

HSC, progenitors

[84]

Pu.1

IRES-GFP (KI)

HSC, lymphoid and myeloid progenitors

[85]

CD41

farnesyl-YFP (Tg)

HSC, progenitors, megakaryocytes, platelets

[86]

Gfi1B

GFP (KI)

HSC, erythroid and myeloid progenitors

[87]

Etv2

GFP (Tg)

Hematopoietic and endothelial progenitors

[88]

Etv2

EYFP (Tg)

Hematopoietic and endothelial progenitors

[89]

Gata1

GFP (Tg)

Hemangioblast, EryP, EryD, megakaryocytes

[90]

EpoR

GFP-Cre (KI)

Erythroid progenitors, endothelial

[91]

Lysozyme EGFP (KI) M

Myelomonocytic cells, including macrophages and granulocytes

[92]

MafB

GFP (Tg)

Myelomonocytic lineages of hematopoietic cells, peritoneal macrophages

[93]

c-fms

EGFP (Tg)

Macrophages, dendritic cells, myeloid cells

[94]

β-globin

ECFP (Tg)

MEP, EryD

[54]

miR-144/ EGFP (KI) 451

EryD

[28]

Eklf

GFP (Tg)

EryD

[95]

ε-globin

KGFP (Tg); H2B-GFP (Tg); H2B-CFP

EryP

[24–26]

γ-globin β-globin

EGFP DsRed

EryP EryD

[96]

Langerin

IRES-EGFP (KI)

Langerhans cells

[97]

TCRb

GFP (Tg)

Lymphoid progenitors

[98]

Runx1

IRES-GFP (KI)

Lymphoid, myeloid, lower levels in erythroid

[31]

CD2

EGFP (Tg)

T lymphoid cells

[99]

Rag2

GFP (KI)

T and B lymphoid cells

[100] (continued)

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Table 1 (continued) Gene

Reporter

Lineage labeled

Reference

FoxP3

GFP (KI)

T regulatory lymphoid cells

[101]

Ror(gT)

EGFP (KI)

T helper 17 lymphoid cells

[102]

Pax5

EGFP (KI)

Pre-B, B lymphoid cells

[103]

Rag1

GFP (KI)

B lymphoid cells

[104]

Blimp1

IRES-EGFP (KI)

B lymphoid cells, plasma cells

[105]

CX3CR1

GFP (KI)

Macrophages, monocytes, NK cells, dendritic cells, microglia

[106]

CD45

YFP (KI)

Widespread hematopoietic

[107]

Abbreviations: Tg transgenic, KI knock-in, CKO conditional knock-out, GFP green fluorescent protein, IRES internal ribosomal entry site, EGFP enhanced GFP, YFP yellow fluorescent protein, CFP cyan fluorescent protein, HSC hematopoietic stem cell, EryP primitive erythroid, EryD definitive erythroid, MEP megakaryocyte-erythroid progenitor, NK natural killer cells a GFP expression is highest in hematopoietic stem cells (HSCs) and is downregulated during lineage commitment and differentiation

can also be achieved by viral infection of the egg or injection of genetically modified embryonic stem (ES) cells into blastocysts [27]. In the “knock-in” method, DNA sequences are engineered to integrate directly into a defined locus within the genome, using site-specific recombination [17]. The knock-in approach avoids problems related to random insertion, as expression of exogenous sequences is controlled by the endogenous regulatory elements of the target gene. An example of a knock-in transgenic mouse is the mirR-144/451-GFP line, in which GFP is expressed in adult and fetal liver erythroid cells [28, 29] (see Fig. 3). A concern for the knock-in approach is that it may result in haploinsufficiency, influencing not only expression of the FP reporter but also the phenotype of the resulting animal. For example, loss of one Runx1 allele affected the distribution of HSCs in the embryo [30]. This problem was overcome by linking the sequences encoding the FP to the endogenous gene through an internal ribosomal entry site (IRES) [31] to create a dicistronic fusion mRNA [32]. Conventional transgenes are often too small to accurately reproduce the endogenous expression of the promoter/enhancer elements. Bacterial artificial chromosomes (BAC) accommodate large DNA sequences, allowing cloning of all or most of the endogenous regulatory elements required to recapitulate the normal pattern of gene expression when linked to other sequences [33] such as those encoding an FP.

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Fig. 3 Tagging of primitive and definitive erythroid lineages using GFP reporters. (a) GFP expression in the primitive erythroid cells of an embryonic stage (E)8.5 ε-globin-H2B-GFP embryo [25] (left panel, whole embryo, scale bar 200 μm; right panel, magnified view of yolk sac, scale bar, 50 μm). Embryos were photographed on a Zeiss Lumar V12 stereomicroscope equipped with epifluorescence illumination and a NeoLumar S 1.5
FWD 30 mm objective. (b) Wet preparation of green fluorescent erythroid cells from the bone marrow (BM) and peripheral blood (PB) of an adult miR-144/451-GFP knock-in mouse [28]. The cells were photographed on a Zeiss Axio Observer Z1 inverted microscope with epifluorescence illumination and a Plan-Apochromat 20
/0.8 objective. Scale bar, 20 μm

CRISPR/Cas9 technology has been used to insert EGFP into the genome of fertilized mouse eggs. Initially, the knock-in frequency obtained using CRISPR/Cas9 was too low to be considered practical. More recently, however, a new approach using a two-part guide RNA annealed with Cas9 protein to form ribonucleoprotein (RNP) complexes has circumvented this limitation [34]. The RNP can be injected into the zygote with a targeting guide RNA to generate site-specific recombination events. CRISPR/Cas9 technology has been used to create lines expressing a fluorescent protein in a developmental stage- and tissue-specific manner [35].

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1.2.3 Alternative Approaches to Drive Hematopoietic Lineage-Specific Expression of Fluorescent Protein Reporters

Inducible expression of fluorescent reporters can be achieved using the Cre-loxP or FLP-FRT systems. For example, a mouse line in which the fluorescent reporter sequences are preceded by a floxed STOP cassette (e.g., refs. [36–38]) can be mated with a deletor line designed to express Cre recombinase under the control of a promotor active in the cells of interest. Such targeted recombination approaches have been of great utility for lineage tracing studies (for a recent example, see ref. [39]). Depending on the system chosen, recombinase activity may be constitutive or inducible. Inducible recombination activity can be triggered by tamoxifen in the case of a Cre-estrogen receptor (ER) fusion gene or by doxycycline when Cre expression is controlled by a Tetracycline (Tet)-responsive element (reviewed in ref. [40]). The Mx1-Cre deletor line can be used for targeting of definitive hematopoietic lineages. In this system, Cre expression is activated by injection of interferon or synthetic double-stranded RNA polyinosinic-polycytidylic acid (poly I:C) [41].

1.3 Fluorescent Protein Reporters

A wide range of FPs covering nearly the entire visible spectrum can be used for generating transgenic reporter mice (reviewed in refs. [2–4]). Since the discovery of wtGFP (see Subheading 1), investigators have sought to generate other FPs with reduced phototoxicity, improved brightness and phosphostability over broad ranges of pH and temperature [2–4]. Site-directed mutagenesis of wtGFP has been employed not only for the generation of FPs with improved functionality but also with diverse spectral characteristics, for example, cyan FP (CFP), blue FP (BFP), and yellow FP (YFP) [2–4]. FPs with emission peaks in the red and far-red spectra have been especially useful for live-cell or whole-animal imaging, owing to their long wavelength emission and, consequently, reduced phototoxicity [2, 42]. Initially, the tetrameric DsRed was cloned from the nonbioluminescent sea anemone Discosoma striata but was found to be toxic to cells [43]. Subsequently, the monomeric variant mRFP was engineered and could be expressed ubiquitously in mice without deleterious effects on development [44]. Directed mutagenesis of mRFP was used to create several variants, including the orange mTomato and the red mStrawberry and mCherry FPs [42]. Far-red fluorophores such as mPlum, genetically engineered from a blue chromoprotein of the sea anemone Actinia equina [45], offer deep tissue penetration and reduced autofluorescence. Spectral properties of various FPs are discussed in detail in refs. [2–4].

1.3.1 Fluorescent Fusion Proteins

FPs can be engineered for localization to specific subcellular regions [46]. For example, to localize the FP to the nucleus, a nuclear localization signal is incorporated into the construct or the FP is fused to histone H2B sequences. Histone H2B fusion-FPs bind to

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chromatin and are present through all phases of the cell cycle [47, 48]. Unlike cytoplasmic GFP, which is diluted during subsequent cell divisions, H2B-FPs are stably expressed and permit monitoring of both the cell cycle and apoptosis [48]. A histone H2B-GFP expressed under the control of a human epsilon-globin promoter and truncated LCR has been used for labeling the nuclei of primitive erythroid cells [25] (see Fig. 3a). Cell morphology and migration can also be observed by labeling the outer or inner leaflet of the cell membrane with lipid-tagged FP fusions (containing, for example, a glycosylphospatidylinositol (GPI) anchor or a myristoylation sequence, ref. [49]). 1.3.2 Photomodulatable FPs

Photoactivatable fluorescent proteins (PAFP) are stimulated by lights of specific wavelengths, intensities, and durations, allowing for spatiotemporal labeling of live cells, organelles, and molecules [2–4]. There are two types of PAFP functions: photoactivation (PA) and photoswitching. Photoactivators convert from a non-fluorescent to a bright fluorescent state and can be either irreversible or reversible. PA-GFP is a GFP variant that is irreversibly converted to an anionic form, resulting in a 100-fold increase in its emission intensity [50]. Tetrameric kindling FP (KFP) can be reversibly or irreversibly photoactivated, depending on the intensity of the activating light [4, 51]. It converts to a red fluorescent state following exposure to green light and returns to a non-fluorescent state in the absence of stimulation [51]. Photoswitchers change their fluorescent state and emit at a different wavelength (such as cyan-green for PS-CFP or green-red for EosFP, Kaede, and Kikume Green-Red, KikGR) upon exposure to transient but intense light [3].

1.3.3 General Considerations for Choosing Fluorescent Protein Reporters

When choosing an FP reporter protein, the investigator should consider whether it will be used for multi-color analysis in combination with other FPs or in immunofluorescence studies with a fluorophore of a different color. The availability of fluorescent variants allows the researcher to select combinations that minimize spectral overlap. For example, the combinations of GFP/CFP and GFP/YFP exhibit significant emission overlap, whereas CFP/YFP does not [52]. Bright reporters in the red or far-red spectra increase the possibilities of combining different reporters to mark cells of different lineages or to mark different regions of the same cell. Due to the lower phototoxicity of the excitation light required by red or far-red fluorophores, these reporters are more suitable for live imaging studies [53]. While imaging or flow cytometric analysis of double transgenic mice expressing FPs with overlapping spectra can be challenging [52], the judicious choice of excitation light and filters will allow optimal separation of reporter signals, as observed for the simultaneous imaging of erythroid cells expressing ECFP and myeloid cells expressing EGFP [54].

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1.4 Confirmation and Analysis of Transgene Expression

The choice of genetic background should be carefully considered in planning the generation of a transgenic mouse line. For microinjection, zygotes of mixed or outbred background are often used [17]. Microinjection of zygotes from inbred mice is more difficult and embryo viability is lower [17].

1.4.1 Mouse Background

Only a relatively small number among the many available inbred strains (e.g., C57BL/6 or 129/Sv) are routinely used to create transgenic or knock-in mice. In the context of the present discussion, inbred strains would be desirable if the transgenic animals will be mated with knock-out mice known to have a backgrounddependent phenotype or if tissue from the genetically modified animals will be used for HSC or other transplantation studies [55]. Genetic background effects (variable penetrance or expressivity) are caused by modifier genes [55]. ICR (CD1) mice are the most widely used outbred mouse strain. Unlike inbred mice, ICR mice display inter-individual genetic variation. However, ICR mice are inexpensive, have excellent reproductive and maternal characteristics, and yield relatively large litter sizes [17].

1.4.2 Breeding

Once founders are identified (see Subheading 3.2), the colony should be expanded. Female founders should be bred so that they can give birth to at least one litter before being sacrificed. Male founders should be placed in a cage with two nontransgenic females and plugs checked daily. Ideally, the male founder should plug 6–8 females in the first few weeks [17]. The gold standard for assessing transgene integration is germline transmission to the F1 generation. By this metric, founders should transmit the transgene to 50% of their progeny. If transmission is not observed, the founder genotype should be reanalyzed. If the founder is positive for the transgene and germline transmission does not occur, it is likely that the founder is mosaic for the transgene and, therefore, either transmits the transgene through the germline at very low levels or not at all. In certain scenarios, transgenes will integrate at multiple loci resulting in progeny that inherit the transgene at unusually high frequencies [56]. In contrast with mouse lines created by gene targeting, each transgenic founder is distinct because of the random nature of transgene integration. Therefore, the decision to eliminate a transgenic mouse line from a colony will be irreversible. To reduce costs, the investigator may choose to maintain an active colony of a few mating pairs or a small number of males that can be mated periodically to produce a younger generation. This is a relatively inexpensive approach but carries the risk that transgene expression may decrease in later generations or as the animals age; this phenomenon has often been seen for globin transgenes [57]. Transgene silencing may be avoided through preserving the line as frozen embryos or sperm (so that in vitro fertilization can be performed at a later date) [55]. Cryopreservation services are provided by

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some institutional core transgenic mouse facilities and by mouse suppliers such as Taconic Farms and Charles River Laboratory. 1.5 Analysis of Fluorescent Protein Expression Using Microscopy

The fluorescence of embryos, tissues, or cells from transgenic reporter mice can be analyzed using epifluorescence or confocal laser scanning microscopy. Confocal microscopes offer several advantages over epifluorescence microscopes. Whereas in epifluorescent microscopy, the entire field is illuminated by light emitted by a mercury or xenon UV lamp, in confocal microscopy, light emitted by the laser is focused through a pinhole, creating point illumination. Out-of-focus signals are thereby eliminated and resolution is increased [58]. The confocal microscope images thin sections of the specimen that can be combined using the microscope’s software into accurate 3D reconstructions of the sample. Confocal microscopes also have an increased level of sensitivity due to light detectors that can amplify the signals received from the specimen. Another advantage of confocal microscopy is that it is less invasive, resulting in reduced photobleaching [58]. The illumination provided by high-power lasers, combined with their reduced light scattering properties, allows imaging of thick, semitransparent sections, live tissues or embryos [58]. Newer model epifluorescent microscopes use LED light sources that are more suitable for live imaging than are classical mercury lamps. For live imaging of explanted embryos or tissues, temperature and gas composition must be carefully controlled using an environmental chamber. Inverted microscopes are typically used to image live material. For a discussion of imaging mouse embryos using confocal microscopy, see refs. [59–61]. Regardless of the type of microscope used, it is essential to select the appropriate light source and filters for analysis of the fluorescent specimen [58]. The identity of the cells expressing the FP may be determined using immunofluorescence, by staining for cell type-specific markers. The staining can be performed on live or fixed cells in solution or on fixed cells deposited on microscope slides. The fixation and permeabilization method should be carefully optimized for each cell type. An overview of different fixation and permeabilization options is reviewed in ref. [58].

1.6 Analysis of Fluorescent Protein Expression Using Flow Cytometry

Analytical flow cytometry is a fundamental technique for assessing fluorescence in cells from transgenic FP reporter mice. Once tissues have been dispersed into single cell suspensions, fluorescent reporter expression from a transgene, combined with antibody staining for specific cell surface markers, can be analyzed using a flow cytometer to identify the cell surface characteristics of the component cell populations. Fluorescence activated-cell sorting (FACS) is a specialized type of flow cytometry that permits the physical separation of a heterogeneous sample into distinct cell populations based on their fluorescence. The ability to isolate labeled cell populations in a single step provides a valuable tool

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for elucidation of their developmental potential, cell cycle, and other properties, and for culture ex vivo. For detailed protocols and reviews, see ref. [62]. 1.7 Analysis of Hematopoietic Progenitor Potential

2

The lineage potential of hematopoietic progenitors can be ascertained by culturing a subset of the target population in a semisolid medium with the appropriate maturation factors. Progenitor distributions will vary depending on the stage of development and location of hematopoiesis (i.e., yolk sac, fetal liver, or bone marrow). In the yolk sac, the earliest hematopoietic progenitors form primarily primitive erythroid and megakaryocyte colonies. Shortly after these lineages arise, a wave of hematopoietic progenitors with myeloid and lymphoid potential emerges. Definitive (adult type) erythroid progenitors first arise in the yolk sac ~ E9.5 from erythromyeloid progenitors (EMPs) that migrate to the fetal liver by E11.5 [63]. HSC-derived erythroid progenitors are found in the fetal liver at ~E12.5 and eventually migrate to the bone marrow, which remains the major source of hematopoietic cells during postnatal life (reviewed in ref. [64]). In a semisolid medium, the progenitors for these different lineages form colonies that have unique morphological features. The frequency of a particular colony type can be calculated by counting the numbers of that colony present and dividing that value by the total number of colonies in the culture. The use of this culture technique to assess the colony forming potential of cells identified by endogenous fluorescence reporter expression has contributed substantially towards the delineation of the stages and hierarchy of hematopoiesis.

Materials

2.1 DNA Purification for Microinjection

1. Agarose. 2. Ethidium bromide. 3. Injection buffer: 10 mM Tris, pH 7.4, 0.2 mM EDTA. 4. QIAquick Gel Extraction Kit (QIAGEN). 5. Restriction enzymes.

2.2 Isolation of Genomic DNA

1. DirectPCR Lysis Reagent (mouse tail) (Viagen).

2.3 Polymerase Chain Reaction (PCR)

1. Plastic tubes for PCR, 0.2 mL.

2. 10 mg/mL Proteinase K in 20 mM Tris–HCl, pH 8. Aliquots are stored at 20  C.

2. TaKaRa ExTaq DNA polymerase, supplied with 10
ExTaq buffer and dNTP mix. 3. Thermal cycler. 4. For agarose gel electrophoresis: agarose, gel casting tray, comb with desired number of teeth, power supply, ethidium bromide, DNA size ladder.

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1. Dissecting scissors and forceps. 2. Watchmaker’s forceps, Dumont #5 and #55. 3. Sterile 3 mL plastic transfer pipettes. 4. Stereomicroscope with transmitted and reflected light sources.

2.5 Glassware and Plasticware

1. 24-well Nunclon tissue culture plates. 2. Circular coverslips, 12 mm, no. 1. 3. 3 and 5 mL syringes. 4. 20G and 25G syringe needles. 5. 15 and 50 mL polypropylene tubes, 15 and 50 mL.

2.6 Embryo dissection and Cell Preparation for Flow Cytometry

1. Phosphate-buffered saline (PBS; pH 7.4). 2. Iscove’s Modified Dulbecco’s Medium (GIBCO Invitrogen). 3. Fetal Bovine Serum (FBS; Hyclone, Thermo Fisher Scientific). 4. Dissection medium: Prepare Iscove’s Modified Dulbecco’s Medium (IMDM) +10% FBS. 5. 12.5 mg/mL heparin (100
) in PBS. 6. BD Falcon 40 μm and 70 μm cell strainers. 7. Cell Dissociation Buffer (Thermo Fisher Scientific). 8. Collagenase (Sigma Aldrich) stock solution; 100 mg/mL collagenase in IMDM supplemented with 20% serum. Aliquots are stored at 20  C.

2.7

Flow Cytometry

1. FACS buffer: heat-inactivated FBS in PBS (see Note 1). 2. DAPI (40 ,6-diamidino-2-phenylindole dihydrochloride). 3. 500 nM propidium iodide (1
). Prepare 1000
stock solution.

2.8 Immunostaining and Microscopy

1. 4% Paraformaldehyde (PFA) in PBS. 2. Washing buffers: PBS with 0.05% Tween-20 (v/v) (PBST); PBST with 0.05% non-fat skim milk powder (Carnation) (PBSMT). 3. Vectashield (Vector Labs) with DAPI or without. 4. Primary and secondary antibodies of choice. 5. Blocking buffer: 2% bovine serum albumin (BSA), 0.1% Triton X-100 in PBS.

2.9 Primitive Erythroid (EryP) Progenitor Assay

1. Primitive erythroid progenitor assay medium: methylcellulose with plasma-derived serum (PDS) (Animal Technologies), protein-free hybridoma medium (PFHM-II) (Gibco), human erythropoietin—Epogen (Amgen). 2. Methylcellulose can be prepared in the lab (see Note 1) or purchased as Methocult (Stem Cell Technologies, without cytokines).

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3. For preparing methylcellulose in house: IMDM containing L-glutamine, penicillin/streptomycin, ascorbic acid, and α-monothioglycerol. 4. Methylcellulose powder. 5. Collagenase or Trypsin/EDTA. 6. 3 and 5 mL syringes. 7. 16-gauge blunt-end needles (Stem Cell Technologies). 8. 3.5 cm petri dishes. 9. 15 cm tissue culture plates. 2.10 Definitive Erythroid and Myeloid Progenitor Assay

1. Methocult™ containing Epo, Scf, IL-3, and IL-6 (Stem Cell Technologies). 2. 16G blunt-end needles. 3. 3.5 cm petri dishes. 4. 15 cm tissue culture plates. 5. Methocult™: per Stem Cell Technologies instructions, thaw the bottle and shake well. Allow bubbles to disperse and aliquot using 16G blunt-end needle. Store at 20  C. Thaw tube once and do not refreeze.

2.11 B Lymphocyte Colony Assay

1. Methocult™ containing IL-7 (Stem Cell Technologies). 2. 16G blunt-end needles. 3. 3.5 cm petri dishes. 4. 15 cm tissue culture plates. 5. Aliquot Methocult™ as above (See Subheading 2.10)

2.12 Megakaryocyte Colony Assay

1. MegaCult™ (Stem Cell Technologies, without cytokines) with recombinant mouse IL-3 (ThermoFisher), mouse IL-6 (ThermoFisher), human IL-11 (ThermoFisher), and rhTPO (Miltyni Biotec); see Subheading 3.6 for concentrations. 2. 16G blunt-end needles. 3. 35 cm petri dishes. 4. 15 cm tissue culture plates. 5. Aliquot Methocult™ as above (see Subheading 2.10).

3

Methods

3.1 DNA Preparation for Microinjection

Transgenes are designed so that the gene to be microinjected can be excised and purified away from plasmid sequences. Prokaryotic sequences from the plasmid do not appear to influence the efficiency of transgene integration but they may impair expression from eukaryotic sequences [15]. Therefore, all plasmid backbone

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should be removed from the transgene construct before microinjection [16, 17]. It is important to include a final purification step to remove particulate material that may clog the injection needle. The following protocol yields clean DNA for microinjection from plasmids smaller than 20 kb. Plasmids larger than 20 kb require a different purification procedure [65], that will not be discussed here. 1. Digest plasmid DNA to completion with the appropriate restriction enzyme(s). 2. Separate the restriction fragments by electrophoresis through an agarose gel (see Note 3). 3. Using an ultraviolet transilluminator, identify and isolate the band containing the transgene to be microinjected. 4. Extract and purify the DNA according to the manufacturer’s protocols for the QIAquick Gel Extraction Kit. 5. Elute the DNA from the QIAquick column using injection buffer (see Subheading 3) and measure the DNA concentration using a NanoDrop spectrophotometer. 6. Pronuclear injection is typically performed in an institutional core transgenic mouse facility. For a detailed protocol, see ref. [66]. At our institution, the purified DNA (1–2 μg, 50–100 ng/μL) is submitted to the Mouse Genetics and Gene Targeting Shared Resource Facility. Filtration and final dilution of the DNA to be used for microinjection is performed by the facility, as is transfer of the injected zygote into pseudopregnant females [17]. Genotyping

Offspring born from injected zygotes are termed “founders” and are usually screened for the presence of the transgene. Genomic screening is most commonly performed using polymerase chain reaction (PCR) analysis of DNA from biopsied tissue (for comments on primer design, see Note 4). In most cases, the microinjected DNA will be stably integrated at the one-cell stage. However, the foreign DNA may integrate at a later (e.g., 4- or 8-cell) stage, resulting in mosaic expression of the transgene and disruption of germline transmission [17]. In addition, silencing of the transgene may occur following integration into or near heterochromatin (see Note 5). It may be possible to identify founders by microscopic analysis of biopsied tissues (see Note 6).

3.2.1 Preparation of Tissue Samples for Genotyping

1. Tail tips 60 years for aged. If you are engaged in a study of immune competence, use donors who are