Liver Carcinogenesis: Methods and Protocols (Methods in Molecular Biology, 2769) [1st ed. 2024] 1071636936, 9781071636930

Table of contents :
Preface
Contents
Contributors
Chapter 1: Orthotopic Model of Hepatocellular Carcinoma in Mice
1 Introduction
2 Materials
2.1 Reagents
2.2 Equipment
2.3 Surgical Supplies
3 Methods
3.1 HCC Cell Preparation for Injection_ Time: 40-60 min
3.2 Implantation of Syngeneic HCC Cells in Liver_ Time: 20-30 min/Mouse
3.2.1 Pre-surgical Setup
3.2.2 Surgical Procedure
3.2.3 Postoperative Care
3.3 Tumor Growth Follow-Up_ Time: 6-8 Weeks
4 Notes
References
Chapter 2: Diethylnitrosamine Induction of Hepatocarcinogenesis in Mice
1 Introduction
1.1 Generalities
1.2 Molecular Mechanisms of DEN-Induced Hepatocarcinogenesis
1.3 Characteristics of DEN-Induced HCC
1.3.1 Histomorphology
1.3.2 Mutational/Transcriptomic Landscape
1.3.3 Immune Infiltration
1.4 General Recommendations
1.5 Advantages and Limitations of the Model
1.6 Materials
2 Methods
3 Notes
References
Chapter 3: Diethylnitrosamine-Induced Liver Tumorigenesis in Mice Under High-Hat High-Sucrose Diet: Stepwise High-Resolution U...
1 Introduction
2 Materials
2.1 DEN Injection
2.2 High-Fat High-Sucrose Diet
2.3 Ultrasonographic Equipment
2.4 Necropsy and Liver Sampling for Histology
3 Methods
3.1 DEN Preparation and Administration to Animals
3.2 High-Fat High-Sucrose Feeding
3.3 Ultrasonographic Assessment of the Liver
3.3.1 Work Station Preparation
3.3.2 Mouse Preparation
3.3.3 Transducer Positioning and Ultrasound Settings
3.3.4 B-Mode Imaging and Ultrasonographic Description
3.3.5 Ultrasound Examination
Cranial Abdomen Imaging and Anatomical Landmarks
Normal Liver Ultrasonography
Pathologic Ultrasonographic Liver Findings
3.3.6 Post-Imaging Mouse Care
3.4 Necropsy and Histological Sample Collection
3.4.1 Sampling
3.4.2 Interpretation of Histopathological Liver Findings
3.5 Tumor Development Modifiers
4 Notes
References
Chapter 4: A Mouse Model of Non-Alcoholic Steatohepatitis and Hepatocellular Carcinoma Induced by Western Diet and Carbon Tetr...
1 Introduction
2 Materials
2.1 Animals
2.2 Diet and Treatment
2.3 CCl4 Injection
2.4 Equipment and Other Materials
3 Methods
3.1 Western Diet (WD) and High-Sugar Water Feeding
3.2 CCl4 Preparation and Administration to Mice
3.3 Ultrasonographic Assessment of HCC
4 Notes
References
Chapter 5: A Mouse Model of Hepatocellular Carcinoma Induced by Streptozotocin and High-Fat Diet
1 Introduction
2 Materials
2.1 Reagents
2.2 Equipment
3 Methods
3.1 Prepare Animals: Time 4-6 h
3.2 Fresh Streptozotocin Buffer Preparation: Time 30 min
3.3 Diabetes Mellitus Induced by Streptozotocin: Time 4 Weeks
3.4 NAS, NASH, Fibrosis, and HCC Accelerated by HFD: Time 20 Weeks
4 Notes
References
Chapter 6: Hydrodynamic Transfection of Hepatocytes for the Study of Hepatocellular Carcinogenesis
1 Introduction
2 Materials
2.1 Equipment
2.2 Disposables
2.3 Mice and Plasmids
3 Methods
3.1 Preparation of Plasmid Mix Solution
3.2 Hydrodynamic Tail Vein Injection
3.3 Monitoring and Tumor Collection
4 Notes
References
Chapter 7: Experimental Model of Biliary Tract Cancers: Subcutaneous Xenograft of Human Cell Lines in Immunodeficient Nude Mice
1 Introduction
2 Materials
2.1 In Vitro Step
2.1.1 Human BTC Cell Lines
2.1.2 Cell Culture Reagents and Equipment
2.2 In Vivo Step
2.2.1 Mouse Strains and Housing
2.2.2 Equipment Required at the Animal Facilities
2.2.3 Equipment Required from the Laboratory
3 Methods
3.1 In Vitro Step
3.1.1 Preparation of BTC Cell Lines
3.1.2 Preparation of Matrigel
3.2 In Vivo Step
3.2.1 Subcutaneous Injection in Mice
3.2.2 Staining and Immunostainings
3.2.3 Representative Results from the BTC Xenograft Model
4 Notes
References
Chapter 8: Oncogene-Driven Induction of Orthotopic Cholangiocarcinoma in Mice
1 Introduction
2 Materials
2.1 Mice
2.2 Preparation of the Plasmid Solution
2.3 Hydrodynamic Injection
2.4 Liver Collection
2.5 MRI Imaging
3 Methods
3.1 In the Animal Facility
3.1.1 Mouse Weighing
3.2 In the Laboratory
3.2.1 Preparation of the Plasmid Solution
3.3 In the Animal Facility
3.3.1 Induction of Cholangiocarcinoma-Hydrodynamic Injection
3.3.2 Liver Collection
3.4 In the Laboratory
3.5 In the MRI Core Facility
3.5.1 MRI Imaging
Setting the Parameters of the MRI Scanner
Liver Immobilization in the MRI Scanner
4 Notes
References
Chapter 9: Isolation of Primary Mouse Hepatocytes and Non-Parenchymal Cells from a Liver with Precancerous Lesions
1 Introduction
2 Materials
2.1 Reagents
2.2 Disposables and Equipment
3 Methods
3.1 Isolation of Primary Mouse Hepatocytes and NPCs by Liver Perfusion
3.1.1 Preparation of Buffers_ Time: 30 min
3.1.2 Isolation of Cell Populations from Liver
Before Perfusion_ Time: 20 min
Perfusion_ Time: 30 min
After Perfusion: Hepatocyte Isolation_ Time: 90 min
After Perfusion: Non-Parenchymal Cells Isolation_ Time: 40 min
3.2 Generation of Leukocyte-Rich Single Cell Suspensions from Non-Perfused Whole Livers
3.2.1 Reagent and Material Preparation_ Time: 15 min
3.2.2 Liver Collection_ Time: 2-5 min per Sample
3.2.3 Liver Processing and Dissociation into a Single Cell Suspension_ Time: 1 h 30 min per Sample (But Simultaneous Sample Is...
4 Notes
References
Chapter 10: Flow Cytometry Assessment of Lymphocyte Populations Infiltrating Liver Tumors
1 Introduction
2 Materials
2.1 Materials and Equipment
2.2 Reagents
3 Methods
3.1 Preparation of Buffers_Time: 10 min
3.2 Preparation of the Single Cell Suspension for Immunostaining_Time: 1 min per Sample (See Notes 1-4)
3.3 Cell Surface Immunostaining_Time: 150 min per Sample (but Simultaneous Sample Staining Is Possible) (See Notes 8-11)
3.4 Intracellular/nuclear Flow Cytometry Staining_Time: 60 min per Sample (but Simultaneous Sample Staining Is Possible)
3.5 Quick Overview of Sample Acquisition Through a Flow Cytometer, Analysis, and Cell Count Normalization
3.5.1 Sample Acquisition Through Flow Cytometer_Time: 120 min for Compensations and 5 min per Sample Acquired (See Note 16)
3.5.2 Analyses and Cell Count Normalization_Time: 120 min per Sample (but Simultaneous Sample Analysis Is Possible) (See Note ...
4 Notes
References
Chapter 11: Immunofluorescent Staining of Human Hepatic Multicellular Spheroids: A Model for Studying Liver Diseases
1 Introduction
2 Materials
2.1 Equipment
2.2 Disposables
2.3 Samples and Reagents
2.3.1 Samples
2.3.2 Reagents Used for Spheroid Cultures and Immunofluorescent Staining
3 Methods
3.1 Spheroid Culture from Primary Cells
3.2 Spheroid Culture from Cell Lines
3.3 Immunofluorescence Staining
3.3.1 Fixation
3.3.2 Permeabilization and Blocking
3.3.3 Immunostaining
3.3.4 Mounting
3.3.5 Data Analysis
3.4 Expected Outcomes
4 Notes
References
Chapter 12: Single-Cell Characterization of the Tumor Ecosystem in Liver Cancer
1 Introduction
2 Materials
2.1 Devices for Tumor Biopsy Collection
2.2 Reagents and Disposables
2.3 Equipment
3 Methods
3.1 Preparation of Tumor Dissociation Kit
3.2 Tumor Biopsy Collection
3.3 Dissociation of Tumor Biopsy into Single Cells
3.4 Single Cell Capture
3.5 Data Analysis
3.5.1 Convert FASTQ Files to Single-Cell Feature Counts
3.5.2 Data Preprocessing
3.5.3 Data Visualization in a Low-Dimensional Space
3.5.4 Separate Malignant and Non-Malignant Cells
3.5.5 Determine Non-Malignant Cell Types
3.5.6 Further Analysis
4 Notes
References
Chapter 13: Chromatin and DNA Dynamics in Mouse Models of Liver Cancers
1 Introduction
2 Materials
2.1 Materials for ATAC-seq
2.1.1 Equipment
2.1.2 Disposables
2.1.3 Reagents
2.2 Materials for ChIP
2.2.1 Equipment
2.2.2 Disposables
2.2.3 Reagents
2.3 Materials for 3C
2.3.1 Equipment
2.3.2 Disposables
2.3.3 Reagents
3 Methods
3.1 ATAC-seq
3.1.1 Transposition Reaction for Primary Hepatocytes (See Note 1)
3.1.2 Transposition Reaction for Non-Tumor and Tumor Samples
3.1.3 Amplification (See Note 2)
3.1.4 Purification
3.1.5 ATAC-Sequencing and Analyses
3.2 ChIP
3.2.1 Crosslinking (See Note 3)
3.2.2 Lysis and Sonication (See Note 4)
3.2.3 Determination of DNA Concentration and Fragment Size
3.2.4 Chromatin Immunoprecipitation (See Note 5)
3.2.5 DNA Isolation
For ChIP Samples
3.2.6 For Inputs
3.2.7 Analysis in qPCR
3.3 3C
3.3.1 Preparation of Nuclei
3.3.2 Cross-Linking
3.3.3 Permeabilization and Restriction Digestion
3.3.4 Ligation
3.3.5 DNA Purification
For Non-Digested and Digested Controls
For 3C Samples
3.3.6 Second Digestion and DNA Purification
3.3.7 Primer Design
3.3.8 Primer Efficiency Control
3.3.9 3C Analysis in qPCR
4 Notes
References
Chapter 14: Targeted Analysis of Glycerophospholipids and Mono-, Di-, or Tri-Acylglycerides in Liver Cancer
1 Introduction
2 Materials
2.1 Disposables
2.2 Equipment
2.3 2.3. Software
2.4 Reagents
3 Methods
3.1 Sample Preparations
3.2 Targeted Analysis of Glycerophospholipids by Flow Injection Analysis (FIA) High-Performance Liquid Chromatography (HPLC) C...
3.3 The Targeted Analysis of Mono-, Di-, and tri-Acylglycerides by Flow Injection Analysis (FIA) High-Performance Liquid Chrom...
3.4 Quality Control Pool
3.5 LC/MS System Rinsing
3.6 Data Processing with LipidView 1.2
4 Notes
References
Chapter 15: Biomarker Identification in Liver Cancers Using Desorption Electrospray Ionization Mass Spectrometry (DESI-MS) Ima...
1 Introduction
2 Materials
2.1 Disposables
2.2 Equipment
2.3 Software
2.4 Reagents
3 Methods
3.1 Sample Preparations (Fig. 1)
3.1.1 Tissue in PFA (Paraformaldehyde) (See Note 1)
3.1.2 HES Staining
3.1.3 Tissue in OCT (Optimal Cutting Temperature) Compound
3.2 MSI Acquisition (Figs. 2 and 3)
3.3 Post-Acquisition Data Treatment
4 Notes
References
Chapter 16: Kinetic Modeling of Hepatic Metabolism and Simulation of Treatment Effects
1 Introduction
2 Methods
2.1 Model Description
2.2 Calibration and Validation
2.3 Application to Proteomic Data
2.4 Scope of Application
2.5 Reference Tissue (Normalization) and Generation of Individual Model Instantiations
2.6 Individual Assessment of Metabolic States under Various Conditions
2.6.1 Maximal Capacities
2.6.2 Assessment of Energetic Capacities and Substrate Utilization Rates under Different Dietary Conditions
2.6.3 Diurnal Metabolic Changes
2.6.4 Simulation of Treatment Effects
2.6.5 Importance of Specific Enzymes for Metabolic Functions
2.6.6 Minimal Hardware and Software Requirements
3 Notes
References
Index

Citation preview

Methods in Molecular Biology 2769

Guido Kroemer Jonathan Pol Isabelle Martins  Editors

Liver Carcinogenesis Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Liver Carcinogenesis Methods and Protocols

Edited by

Guido Kroemer, Jonathan Pol and Isabelle Martins Centre de Recherche des Cordeliers, Paris, France

Editors Guido Kroemer Centre de Recherche des Cordeliers Paris, France

Jonathan Pol Centre de Recherche des Cordeliers Paris, France

Isabelle Martins Centre de Recherche des Cordeliers Paris, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3693-0 ISBN 978-1-0716-3694-7 (eBook) https://doi.org/10.1007/978-1-0716-3694-7 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface Every year, more than 900,000 people are diagnosed with liver cancer, and approximately 800,000 persons succumb to the disease, making it the third leading cause of cancer death worldwide. Primary liver tumors mainly comprise hepatocellular carcinoma (HCC) and cholangiocarcinoma (CCA), encompassing 75–85% and 10–20% of cases, respectively. Other rare hepatic malignancies include angiosarcoma in adults, as well as hepatoblastoma and hemangiosarcoma in children. This tapestry of liver tumors has rather diverse etiologies. On the one hand, HCC finds its origin in a mosaic of risk factors that include chronic hepatitis B and C virus infections, exposure to aflatoxins, excessive alcohol consumption, nonalcoholic fatty liver disease (NAFLD), cirrhosis, diabetes, or the presence of inherited liver disorders like hemochromatosis and alpha-1 antitrypsin deficiency. On the other hand, the emergence of CCA mostly occurs in the context of primary sclerosing cholangitis (PSC), bile duct anomalies (e.g., cysts, stones), parasitic infections (e.g., liver flukes), other chronic liver diseases (e.g., cirrhosis, fatty liver disease, viral hepatitis), smoking, or diabetes. Even more alarming than the current situation is the impending escalation of liver cancer incidence. With an unsettling trajectory, both the incidence of new cases and the harrowing toll of lives claimed by the disease are projected to surge by over 50% within the next two decades. This dire forecast is wrought from a multitude of factors, including the inevitable aging of the world population, the uneven accessibility to vaccinations and antiviral treatments for viral hepatitis, and lifestyles that veer away from healthy diets and physical activity, among others. In response to this looming crisis, it becomes paramount to unveil palliative measures. These encompass the urgent need for public health initiatives, lifestyle adjustments, and improved medical management of liver diseases. Given the current and projected situation, multipronged research efforts must be pursued to unravel the complexities of liver cancer genesis and progression, set up efficient solutions to prevent its occurrence, uncover druggable molecular and cellular targets, design innovative therapeutic interventions, and identify biomarkers allowing timely detection, prognostication, and prediction of treatment responses. Within this overarching framework, this book brings together a collection of modern methodologies employed in the investigation of liver carcinogenesis. The first half of this volume delineates pertinent preclinical models of HCC and CCA, established either through orthotopic induction or ectopic implantation. The second half regroups a diverse array of techniques applied to characterize the biochemical and cellular composition of hepatic malignancies that operate at the singlecell and histological levels. This compilation stands as a valuable resource for scientists and students alike, driving progress in the field of liver cancer research. Paris, France

Guido Kroemer Jonathan Pol Isabelle Martins

v

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

v ix

1 Orthotopic Model of Hepatocellular Carcinoma in Mice . . . . . . . . . . . . . . . . . . . . 1 ˜ o, Le´a Monte´gut, Flavia Lambertucci, Sijing Li, Omar Motin Uxı´a Nogueira-Recalde, Hui Chen, Gerasimos Anagnostopoulos, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins 2 Diethylnitrosamine Induction of Hepatocarcinogenesis in Mice . . . . . . . . . . . . . . 15 Jules Sotty, Pierre Bablon, Paul-Henry Weiss, and Patrick Soussan 3 Diethylnitrosamine-Induced Liver Tumorigenesis in Mice Under High-Hat High-Sucrose Diet: Stepwise High-Resolution Ultrasound Imaging and Histopathological Correlations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 Pierre Cordier, Flora Sangouard, Jing Fang, Christelle Kabore, Chantal Desdouets, and Se´verine Celton-Morizur 4 A Mouse Model of Non-Alcoholic Steatohepatitis and Hepatocellular Carcinoma Induced by Western Diet and Carbon Tetrachloride . . . . . . . . . . . . . . 57 ˜ o, Flavia Lambertucci, Hui Chen, Sijing Li, Omar Motin Gerasimos Anagnostopoulos, Le´a Monte´gut, Uxı´a Nogueira-Recalde, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins 5 A Mouse Model of Hepatocellular Carcinoma Induced by Streptozotocin and High-Fat Diet. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 ˜ o, Sijing Li, Flavia Lambertucci, Omar Motin Gerasimos Anagnostopoulos, Le´a Monte´gut, Uxı´a Nogueira-Recalde, Hui Chen, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins 6 Hydrodynamic Transfection of Hepatocytes for the Study of Hepatocellular Carcinogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Jose´ Ursic-Bedoya and Damien Gregoire 7 Experimental Model of Biliary Tract Cancers: Subcutaneous Xenograft of Human Cell Lines in Immunodeficient Nude Mice. . . . . . . . . . . . . . 87 Bouchra Lekbaby, Javier Vaquero, Allan Pavy, Mirko Minini, Ester Gonzalez-Sanchez, Je´re´my Augustin, and Laura Fouassier 8 Oncogene-Driven Induction of Orthotopic Cholangiocarcinoma in Mice. . . . . . 99 Ce´leste Plantureux, Juliette Paillet, Gwennhael Autret, Maria Pe´rez-Lanz
o n, Guido Kroemer, Maria Chiara Maiuri, and Jonathan Pol 9 Isolation of Primary Mouse Hepatocytes and Non-Parenchymal Cells from a Liver with Precancerous Lesions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 ˜ o, Maria Pe´rez-Lanz
o n, Sijing Li, Flavia Lambertucci, Omar Motin Ce´leste Plantureux, Jonathan Pol, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins

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11

12 13 14

15

16

Contents

Flow Cytometry Assessment of Lymphocyte Populations Infiltrating Liver Tumors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maria Pe´rez-Lanz
on, Ce´leste Plantureux, Juliette Paillet, Jules Sotty, Patrick Soussan, Guido Kroemer, Maria Chiara Maiuri, and Jonathan Pol Immunofluorescent Staining of Human Hepatic Multicellular Spheroids: A Model for Studying Liver Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Me´lanie Couteau and Lynda Aoudjehane Single-Cell Characterization of the Tumor Ecosystem in Liver Cancer. . . . . . . . . Limin Wang, Mahler Revsine, Xin Wei Wang, and Lichun Ma Chromatin and DNA Dynamics in Mouse Models of Liver Cancers . . . . . . . . . . . Julie Sanceau, Thierry Forne´, Sophie Chantalat, and Ange´lique Gougelet Targeted Analysis of Glycerophospholipids and Mono-, Di-, or Tri-Acylglycerides in Liver Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hui Chen, Sylve`re Durand, Me´lanie Bourgin, Flavia Lambertucci, ˜ o, Le´a Monte´gut, Sijing Li, Uxı´a Nogueira-Recalde, Omar Motin Gerasimos Anagnostopoulos, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins Biomarker Identification in Liver Cancers Using Desorption Electrospray Ionization Mass Spectrometry (DESI-MS) Imaging: An Approach for Spatially Resolved Metabolomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hui Chen, Sylve`re Durand, Olivia Bawa, Me´lanie Bourgin, Le´a Monte´gut, ˜ o, Sijing Li, Uxı´a Nogueira-Recalde, Flavia Lambertucci, Omar Motin Gerasimos Anagnostopoulos, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins Kinetic Modeling of Hepatic Metabolism and Simulation of Treatment Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antje Egners, Thorsten Cramer, Iwona Wallach, and Nikolaus Berndt

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

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Contributors GERASIMOS ANAGNOSTOPOULOS • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMICCa, Gustave Roussy, Villejuif, France LYNDA AOUDJEHANE • Sorbonne Universite´, INSERM, Institute of Cardiometabolism and Nutrition (IHU-ICAN), Paris, France; Sorbonne Universite´, INSERM, Centre de Recherche Saint-Antoine (CRSA), Paris, France JE´RE´MY AUGUSTIN • Universite´ Paris Est Cre´teil, INSERM, IMRB, Cre´teil, France; Department of Pathology, AP-HP, Henri Mondor Hospital, Cre´teil, France GWENNHAEL AUTRET • Plateforme Imageries du Vivant, Universite´ de Paris, PARCC, INSERM, Paris, France PIERRE BABLON • Sorbonne Universite´, Institut National de la Sante´ et de la Recherche Me´ dicale (INSERM), Centre de Recherche de Saint Antoine (CRSA), Paris, France OLIVIA BAWA • PETRA, UMS AMICCa, Gustave Roussy, Villejuif, France NIKOLAUS BERNDT • Deutsches Herzzentrum der Charite´ (DHZC), Institute of Computerassisted Cardiovascular Medicine, Berlin, Germany; Charite´ – Universit€ a tsmedizin Berlin, corporate member of Freie Universit€ a t Berlin and Humboldt-Universit€ a t zu Berlin, Berlin, Germany; Department of Molecular Toxicology, German Institute of Human Nutrition Potsdam-Rehbruecke (DIfE), Nuthetal, Germany ME´LANIE BOURGIN • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMICCa, Gustave Roussy, Villejuif, France SE´VERINE CELTON-MORIZUR • Centre de Recherche des Cordeliers, Sorbonne Universite´, INSERM, Universite´ Paris Cite´, Paris, France; Genomic Instability, Metabolism, Immunity and Liver Tumorigenesis Laboratory, Equipe Labellise´e Ligue Contre le Cancer, Paris, France SOPHIE CHANTALAT • Centre National de Ge´notypage, Institut de Ge´nomique, CEA, Evry, France HUI CHEN • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMICCa, Gustave Roussy, Villejuif, France; Faculte´ de Me´decine, Universite´ de Paris Saclay, Kremlin Biceˆtre, France PIERRE CORDIER • Centre de Recherche des Cordeliers, Sorbonne Universite´, INSERM, Universite´ Paris Cite´, Paris, France; Genomic Instability, Metabolism, Immunity and Liver Tumorigenesis Laboratory, Equipe Labellise´e Ligue Contre le Cancer, Paris, France ME´LANIE COUTEAU • Sorbonne Universite´, INSERM, Institute of Cardiometabolism and Nutrition (IHU-ICAN), Paris, France THORSTEN CRAMER • Molecular Tumor Biology, Department of General, Visceral and Transplantation Surgery, RWTH University Hospital, Aachen, Germany; Department of Surgery, Maastricht University Medical Center, Maastricht, The Netherlands; NUTRIM

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Contributors

School of Nutrition and Translational Research in Metabolism, Maastricht University, Maastricht, The Netherlands CHANTAL DESDOUETS • Centre de Recherche des Cordeliers, Sorbonne Universite´, INSERM, Universite´ Paris Cite´, Paris, France; Genomic Instability, Metabolism, Immunity and Liver Tumorigenesis Laboratory, Equipe Labellise´e Ligue Contre le Cancer, Paris, France SYLVE`RE DURAND • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMICCa, Gustave Roussy, Villejuif, France ANTJE EGNERS • Molecular Tumor Biology, Department of General, Visceral and Transplantation Surgery, RWTH University Hospital, Aachen, Germany JING FANG • Centre de Recherche des Cordeliers, Sorbonne Universite´, INSERM, Universite´ Paris Cite´, Paris, France; Genomic Instability, Metabolism, Immunity and Liver Tumorigenesis Laboratory, Equipe Labellise´e Ligue Contre le Cancer, Paris, France THIERRY FORNE´ • IGMM, Univ. Montpellier, CNRS, Montpellier, France LAURA FOUASSIER • Centre de Recherche Saint-Antoine (CRSA), INSERM, Sorbonne Universite´, Paris, France ESTER GONZALEZ-SANCHEZ • TGF-β and Cancer Group, Oncobell Program, Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain; National Biomedical Research Institute on Liver and Gastrointestinal Diseases (CIBERehd), Instituto de Salud Carlos III, Madrid, Spain; Department of Physiological Sciences, Faculty of Medicine and Health Sciences, University of Barcelona, Barcelona, Spain ANGE´LIQUE GOUGELET • Centre de Recherche des Cordeliers, Sorbonne Universite´, Inserm, Universite´ de Paris, Paris, France; Team “Oncogenic functions of beta-catenin signaling in the liver”, E´quipe labellise´e par la Ligue contre le Cancer, Paris, France DAMIEN GREGOIRE • Institut de Ge´ne´tique Mole´culaire de Montpellier, University of Montpellier, CNRS, Montpellier, France CHRISTELLE KABORE • Centre de Recherche des Cordeliers, Sorbonne Universite´, INSERM, Universite´ Paris Cite´, Paris, France; Genomic Instability, Metabolism, Immunity and Liver Tumorigenesis Laboratory, Equipe Labellise´e Ligue Contre le Cancer, Paris, France GUIDO KROEMER • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Department of Biology, Institut du Cancer Paris CARPEM, Hoˆpital Europe´en Georges Pompidou, AP-HP, Paris, France FLAVIA LAMBERTUCCI • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France BOUCHRA LEKBABY • Centre de Recherche Saint-Antoine (CRSA), INSERM, Sorbonne Universite´, Paris, France SIJING LI • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Faculte´ de Me´decine, Universite´ de Paris Saclay, Kremlin Biceˆtre, France LICHUN MA • Liver Cancer Program, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA; Cancer Data Science Laboratory, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA

Contributors

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MARIA CHIARA MAIURI • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Department of Molecular Medicine and Medical Biotechnologies, University of Napoli Federico II, Naples, Italy ISABELLE MARTINS • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France MIRKO MININI • Centre de Recherche Saint-Antoine (CRSA), INSERM, Sorbonne Universite´, Paris, France LE´A MONTE´GUT • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Faculte´ de Me´decine, Universite´ de Paris Saclay, Kremlin Biceˆtre, France OMAR MOTIN˜O • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France UXI´A NOGUEIRA-RECALDE • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Rheumatology Research Group (GIR), Biomedical Research Institute of A Corun˜a (INIBIC), Professor Novoa Santos Foundation, A Corun ˜ a, Spain JULIETTE PAILLET • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Laboratory of Human Lymphohematopoieisis, Imagine Institute, INSERM UMR 1163, Universite´ Paris Cite´, Paris, France; Smart Immune, Paris, France ALLAN PAVY • Centre de Recherche Saint-Antoine (CRSA), INSERM, Sorbonne Universite´, Paris, France MARIA PE´REZ-LANZO´N • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France CE´LESTE PLANTUREUX • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France; Faculte´ de Me´decine, Universite´ Paris-Saclay, Kremlin-Biceˆtre, France JONATHAN POL • Centre de Recherche des Cordeliers, Equipe labellise´e par la Ligue contre le cancer, Inserm U1138, Universite´ Paris Cite´, Sorbonne Universite´, Institut Universitaire de France, Paris, France; Metabolomics and Cell Biology Platforms, UMS AMMICa, Gustave Roussy, Villejuif, France MAHLER REVSINE • Laboratory of Human Carcinogenesis, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA

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Contributors

JULIE SANCEAU • Centre de Recherche des Cordeliers, Sorbonne Universite´, Inserm, Universite´ de Paris, Paris, France; Team “Oncogenic functions of beta-catenin signaling in the liver”, E´quipe labellise´e par la Ligue contre le Cancer, Paris, France FLORA SANGOUARD • Centre de Recherche des Cordeliers, Sorbonne Universite´, INSERM, Universite´ Paris Cite´, Paris, France; Genomic Instability, Metabolism, Immunity and Liver Tumorigenesis Laboratory, Equipe Labellise´e Ligue Contre le Cancer, Paris, France JULES SOTTY • Sorbonne Universite´, Institut National de la Sante´ et de la Recherche Me´dicale (INSERM), Centre de Recherche de Saint Antoine (CRSA), Paris, France PATRICK SOUSSAN • Sorbonne Universite´, Institut National de la Sante´ et de la Recherche Me´ dicale (INSERM), Centre de Recherche de Saint Antoine (CRSA), Paris, France; Assistance Publique – Hoˆpitaux de Paris (AP-HP). Sorbonne Universite´, De´partement de Virologie, GHU Paris-Est, Paris, France JOSE´ URSIC-BEDOYA • Institut de Ge´ne´tique Mole´culaire de Montpellier, University of Montpellier, CNRS, Montpellier, France; Department of Hepatogastroenterology, Hepatology and Liver Transplantation Unit, Saint Eloi Hospital, University of Montpellier, Montpellier, France JAVIER VAQUERO • TGF-β and Cancer Group, Oncobell Program, Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain; National Biomedical Research Institute on Liver and Gastrointestinal Diseases (CIBERehd), Instituto de Salud Carlos III, Madrid, Spain; Centro de Investigaci
on del Ca´ncer and Instituto de Biologı´a Molecular y Celular del Ca´ncer, CSIC-Universidad de Salamanca, Salamanca, Spain IWONA WALLACH • Deutsches Herzzentrum der Charite´ (DHZC), Institute of Computerassisted Cardiovascular Medicine, Berlin, Germany; Charite´ – Universit€ a tsmedizin Berlin, corporate member of Freie Universit€ a t Berlin and Humboldt-Universit€ a t zu Berlin, Berlin, Germany LIMIN WANG • Laboratory of Human Carcinogenesis, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA XIN WEI WANG • Laboratory of Human Carcinogenesis, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA; Liver Cancer Program, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA PAUL-HENRY WEISS • Sorbonne Universite´, Institut National de la Sante´ et de la Recherche Me´dicale (INSERM), Centre de Recherche de Saint Antoine (CRSA), Paris, France

Chapter 1 Orthotopic Model of Hepatocellular Carcinoma in Mice Flavia Lambertucci, Sijing Li, Omar Motin˜o, Le´a Monte´gut, Uxı´a Nogueira-Recalde, Hui Chen, Gerasimos Anagnostopoulos, Maria Chiara Maiuri, Guido Kroemer, and Isabelle Martins Abstract Orthotopic models of hepatocellular carcinoma (HCC) consist in the implantation of tumor cells into the liver by direct intrahepatic injection. In this model, tumorigenesis is triggered within the hepatic microenvironment, thus mimicking the metastatic behavior of HCC. Herein, we detail a surgically mediated methodology that allows the reproducible and effective induction of liver-sessile tumors in mice. We enumerate the steps to be followed before and after the surgical procedure, including HCC cell preparation, the quantity of cancer cells to be injected, presurgical preparation of the mice, and finally, postoperative care. The surgical procedure involves laparotomy to expose the liver, injection of cells into the left-lateral hepatic lobe, and closure of the incision with sutures followed by wound clips. We also provide information concerning the subsequent tumor growth follow-up, as well as the application of bioluminescence imaging to monitor tumor development. Key words Bioluminescent imaging, Hepatocellular carcinoma, Intrahepatic injection, Orthotopic cancer model, Surgical procedure

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Introduction Most cases of liver cancer occur in the setting of chronic liver disease, making the design of adequate mouse models challenging [1]. For a detailed and effective characterization of hepatocellular carcinoma (HCC) treatment, different models should be employed. Animal models in which development of initially non-malignant liver diseases is followed by hepatic oncogenesis and tumor progression are best suited to study the biology of hepatocarcinogenesis [2]. Nevertheless, mimicking the physiological changes of HCC development requires long-lasting treatments. Injection of HCC tumor cells into mice is suitable for establishing a

Guido Kroemer et al. (eds.), Liver Carcinogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2769, https://doi.org/10.1007/978-1-0716-3694-7_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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biologically relevant model of the disease in order to evaluate novel treatment strategies. Thanks to their simplistic setup and increased reproducibility, implantation models are often used to test new drugs [2]. Injection of murine or human cancer cell lines as well as implantation of tumor tissue subcutaneously or into the liver of recipient mice, represent the most common methods to establish HCC tumors [3]. Injection of a tumor into its tissue of origin is known as an orthotopic model, whereas tumors inoculated into a foreign tissue, for example, subcutaneously, constitute heterotopic models. The dual hepatic vascularization with its connection to the gut, the tumor–stroma interactions, and the metastatic behavior of HCC require orthotopic models more than in any other cancer field [4]. When compared to subcutaneous implantations, orthotopic models are technically more challenging to perform. However, they may reflect more accurately the tumor microenvironment (TME). Orthotopic tumors derived from mouse or human cells can be introduced via direct intrahepatic, intrasplenic, or intraportal injection [5]. In syngeneic models, murine cancer cells or murine tumors are implanted in immunocompetent mice. In xenograft models, human cancer cell lines or human tumors are injected in immunocompromised murine recipients to avoid rejection of the foreign tissue. For this reason, xenograft models are not expected to mimic the TME and the immune reactions against the tumor [6, 7]. Nowadays, it is of major importance to consider the role of the TME on HCC development, progression, and response to therapy [8, 9]. The present protocol describes in detail all the steps required for a syngeneic mouse liver orthotopic implantation model. This HCC model constitutes a powerful tool to uncover the mechanisms regulating tumor initiation and progression [10]. As an example, we established a unifocal HCC model developing a single tumor nodule by implantation of Hep55.1C-Luc murine tumor cells in the liver of syngeneic 7-week-old female C57BL/6J mice. Tumor evolution can be monitored by bioluminescent imaging (BLI) as well as histologically. The implantation of Hep55.1C cells in the syngeneic mice liver enables the fast growth of a highly differentiated tumor, expressing fibrotic and cancer marker genes [5]. This model represents a reproducible HCC model in the C57BL/6J mouse strain, which has a low propensity to develop spontaneous cancers. Established luciferase-expressing HCC cell lines have been successfully used in an orthotopic immunodeficient xenograft mouse model, which enables easy repeated monitoring of tumor growth by in vivo bioluminescence that correlates well with MRI imaging [11]. BLI consists in the detection of luminescence emitted following the expression of luciferase genes in tumor cells and the administration of the substrate luciferin [12]. Although BLI has been

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found to correlate with other imaging modalities, when tumors reach the end stage of disease, BLI may underestimate tumor growth as the tumor outgrows its blood supply [13]. Nevertheless, due to the local injection of tumor cells, mice develop a solid single tumor nodule that it is easily detected with BLI. Additionally, BLI is a useful tool to monitor changes in tumor growth induced by different treatment modalities [14, 15]. Prior to injection, HCC cells are trypsinized, resuspended in culture medium, and maintained in ice until the moment of injection (Subheading 3.1). After the recipient mouse is anesthetized, a parallel incision along the linea alba is made in the abdominal wall to expose the liver. Then, the injection is performed orthotopically into the left-lateral hepatic lobe. The peritoneum is closed with suture, and wound clips are used for closure of the skin. Intrahepatic implantation of mouse HCC cells requires 20–30 min per mouse (Subheading 3.2). Tumor growth is followed by BLI for 2 to 8 weeks after implantation. Mice are sacrificed 6–8 weeks after implantation, and histological analysis of the liver by HES staining is performed (Subheading 3.3). Tumor growth varies according to the tumor cell line and mouse strain used. Additionally, in later stages of the disease, the mice may develop local metastases in adjacent lobes of the liver, as well as lung metastases. Development of one big tumor nodule, which can be easily distinguished from the rest of the liver tissue, is useful for a precise measurement of tumor size and for immunotherapy studies, as mononuclear cells from the tumor and surrounding liver tissue can be analyzed separately. This distinguishes this model from others (e.g., genetically engineered mice), which may generate several tumors throughout the liver [16]. Since the protocol consists in a surgical procedure on living mice, it requires the acquisition of basic surgery skills to perform the procedure in an ethically accepted, effective, safe, and reproducible manner. Finally, variations of this protocol can be performed in the context of non-alcoholic fatty liver disease (NAFLD), injecting HCC cells orthotopically in preconditioned livers with high-fat-diet feeding or underlying liver cirrhosis [10, 15]. Indeed, simultaneous induction of tumor growth and development of liver disease can help to understand the crosstalk between hepatic tumors and the local environment.

2 2.1

Materials Reagents

1. Culture medium: DMEM high-glucose medium (pH 7.0 to 7.4, with 4.5 g/L D-glucose, L-glutamine, phenol red, sodium pyruvate), 10% fetal bovine serum (FBS), 1% penicillin/streptomycin antibiotics.

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2. Trypsin suitable for cell culture. 3. 70% ethanol. 4. 10% povidone iodine solution or Betadine. 5. Luciferase-expressing HCC line: Hep 55.1c-Luc. 6. Serological pipettes and pipette tips. 7. T75 Flask (suitable for cell culture). 8. Eppendorf and centrifuge tubes. 9. C57Bl/6J female mice, 7 weeks-old. 10. Isoflurane (gas anesthesia system; according to the specific animal protocol at your institution and animal facility). 11. Analgesia (according to the specific animal protocol at your institution). 12. 1X PBS, pH 7.4. 13. 4% Formaldehyde solution, buffered, pH 6.9 (approx. 10% formalin solution) for histology. 14. IVISbrite D-luciferin potassium salt bioluminescent substrate, XenoLight. 2.2

Equipment

1. Centrifuge. 2. Bright-field microscope. 3. Incubator. 4. Laminar flow hood. 5. Heating pad. 6. Heat lamp. 7. Isoflurane vaporizer. 8. Electric trimmer. 9. Scale (capable of measuring mouse weight). 10. Bioluminescence in vivo imaging system (IVIS Spectrum). 11. Caliper.

2.3

Surgical Supplies

1. Eye protective gel. 2. Tape. 3. 1 mL syringe. 4. 25-Gauge needle. 5. Autoclip applier plus 9-mm autoclips. 6. Sterile cotton tips. 7. Sterile gauze. 8. 5-0 PDS (Polydioxanone Suture), vicryl, or silk suture for peritoneum closure.

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9. Sterile absorbable gelatin sponge. 10. Straight forceps. 11. Curved fine forceps. 12. Straight scissor. 13. Hartman hemostats. 14. Wound clip remover.

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Methods

3.1 HCC Cell Preparation for Injection_ Time: 40– 60 min

1. Prewarm culture medium (previously prepared) in a 37 °C water bath before use and prewarm trypsin at room temperature. 2. Remove the medium from the flask containing Hep55.1C-Luc cells at 80% confluence (see Notes 1, 2, and 3). Rinse cells twice with 1X PBS. 3. Add 2 mL of trypsin to the cells. Gently shake the flask to ensure full coverage of the surface area by the trypsin solution. Put the flask at 37 °C in the incubator (see Note 4). 4. After 3–5 min, check the cells detaching with a bright-field microscope. Gently tap the flask to facilitate cell detachment (see Note 5). 5. Once cells are detached, add 6–8 mL of serum containing culture medium to neutralize the trypsin in the flask. Gently resuspend and mix the cell suspension to finish detaching cells. Collect the cells in a 15-mL centrifuge tube. 6. Centrifuge the cells for 5 min at 200–250 g at room temperature and resuspend the cells in culture medium. 7. Count cells to determine the number of cells in the suspension. Adjust the cell concentration to 30 × 104 cells per 50 μL in DMEM-10% FBS (corresponding to the final cell concentration and volume to be injected). Keep the cell suspension in ice (see Note 6).

3.2 Implantation of Syngeneic HCC Cells in Liver_ Time: 20– 30 min/Mouse 3.2.1

Pre-surgical Setup

(see Notes 7, 8, and 9) (Fig. 1) 1. Properly arrange all instrumentations and solutions that are used during the experimentation prior to surgery. 2. For anesthesia of mice, an isoflurane chamber with nose cone attachment is needed. Anesthetize the mouse by inhalation of 1.5–3 vol% isoflurane in 100% oxygen at a flow rate of 1 L/min. The depth of anesthetization is sufficient when the following vital criteria are reached: regular spontaneous breathing, no reflex after setting of pain stimuli between toes, and no response to pain. Apply pre-operative analgesia subcutaneously,

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a

b

Peritoneal incision and liver exposure

d

c

Injection of cell suspension

Intrahepatic HCC cell injection

Surgical closure of incision

Fig. 1 Graphical abstract of the surgical procedure

either meloxicam (5 mg/kg, subcutaneous injection) or buprenorphine (0.1 mg/kg, subcutaneous injection). 3. During the complete experimentation, keep the animal on a heating pad at a temperature of 37 °C and fix the legs of the animal with stripes of tape. 4. Shave the abdominal fur of the mouse using an electric trimmer and disinfect the skin with 70% ethanol followed by sterile gauzes with a 10% povidone-iodine solution. Protect the eyes from drying out by usage of eye protective gel (see Note 10). 3.2.2

Surgical Procedure

1. Grab the skin with a pair of straight blunt forceps and perform a midline abdominal incision of the skin (below the thoracic cavity). Incision should be 2.0–2.5 cm long. Do not penetrate into the peritoneal cavity. 2. Gently free the skin from the peritoneum. Grab the peritoneum lifting straight upward and open the peritoneal cavity with a midline laparotomy of a length of approximately 1.5–2 cm, ensuring to see the tip of the scissors as to not injure underlying organs. Incision in the peritoneum should be performed along the linea alba (midlines white fascia). If the incision is correctly made, the liver should be the first organ to be visualized (see Note 11) (Fig. 2a). 3. Gently pull out the left-lateral lobe with a saline-moistened cotton tip and expose the lobe out of the peritoneal cavity (see Note 12) (Fig. 2b). 4. Immediately before orthotopic implantation, cell suspension should be mixed. Fill a 1 mL syringe attached to a 25-gauge needle with 50 μl of cell suspension (all 50 μl should be injected). This will ensure a final cell number of 30 × 104 cells implanted (see Note 13).

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Fig. 2 Surgical procedure for intrahepatic HCC cell injection. (a) Skin and peritoneal incisions and liver exposure. (b) Exteriorization of the left-lateral hepatic lobe. (c) Insertion of the needle under the liver capsule. (d) Injection of 50 μl cell suspension and visualization of the change in the liver color. (e) Covering of the leftlateral lobe injection site with a piece of sterile sponge. (f) Relocation of the left-lateral lobe in the peritoneal cavity with the piece of sterile sponge. (g) Closure of muscle layer by a continuous suture. (h) Skin layer closed with 9-mm autoclips

5. Keep immobilizing the liver lobe with the help of a cotton tip and visualize the injection site. 6. Insert the needle into the liver parenchyma traversing several millimeters deep (under the liver capsule) and gently inject the cell suspension (Fig. 2c). The color of the liver surface injected should slowly change to a brown-to-white shade during cell implantation (see Notes 14 and 15) (Fig. 2d). 7. After injecting the cell suspension, keep the needle for a few seconds in the same position to then slowly retract it. 8. Immediately after removing the needle of the liver parenchyma, place a small piece of sterile sponge (0.5 × 0.5 cm) on the injected site applying gentle pressure with a cotton tip for a few minutes to prevent possible bleeding or leakage (see Note 16) (Fig. 2e). 9. Replace the liver to its physiological position, leaving the sterile sponge onto the injection site which will be slowly resorbed (Fig. 2f). Using a 5-0 suture, close the peritoneal layer by continuous or interrupted sutures (Fig. 2g). Cut the ends of the sutures and sterilize the operation area with a gauze swab moistened with 10% povidone-iodine solution.

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10. Close the skin layer of the abdominal wall by placing two or three 9-mm autoclips using the autoclip applier. Simple interrupted sutures can also be used (see Note 17) (Fig. 2h). 3.2.3

Postoperative Care

1. Allow the mouse to recover in a cage warmed up by a heat lamp until the mouse is fully awake and active. A heating pad may also be used instead of a heat lamp (see Note 18). 2. Apply postoperative analgesia according to the specific animal protocol at your institution. 3. Place animals back in their cages with unlimited access to food and water.

3.3 Tumor Growth Follow-Up_ Time: 6– 8 Weeks

1. Wound clips must be removed in 7–10 days once the incision has healed. 2. Closely monitor the mice for tumor progression and continue to assess their health until the experiment is finished (see Note 19). 3. Anesthetize the mice using 2% isoflurane in an anesthesia chamber. 4. Administer an intraperitoneal injection of 150 mg/kg of D-luciferin in 1X PBS. Wait 5 min. 5. Perform imaging. We utilized the IVIS Spectrum in vivo imaging system (Fig. 3). Images were acquired with the same settings and exposure times for all animals (see Note 20). 6. After imaging, let the mice recover from anesthesia in a cage with a heat lamp. 7. After 6–8 weeks sacrifice animals and retrieved liver for histological and biochemical analysis (see Note 21) (Fig. 4). 8. Excised tumors and normal liver can be fixed in 10% formalin solution for later histological analyses like hematoxylin-eosinsafranin (HES) staining (Fig. 5).

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Notes 1. Avoid using antibiotics to cultured cells at least 1 week before this step. Cells should be in culture for approximately four passages prior to injection and cell passaging should be avoided at least 2 days before this step. 2. Here we use Hep55.1C-Luc cells as HCC cells since they are syngeneic and can be injected and grown in C57BL6J mice; other HCC cells can also be used. Also, the use of a luciferase expressing cell line allows a follow-up on tumor growth by BLI after implantation of cells.

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Fig. 3 Representative images obtained with IVIS Spectrum in vivo imaging system. Bioluminescent signal measured in three anesthetized female mice, two positive controls with tumor, and one negative control without intrahepatic tumor injection

Fig. 4 Representative livers after euthanasia of control and tumor implanted mice after 6 weeks

3. Hep55.1C-Luc cells were maintained in Dulbecco’s modified Eagle’s medium supplemented with 4.5 g/L glucose (see Subheading 2) and 10% FBS in a 5% CO2 humidified chamber. Cell lines used should be regularly checked to ensure they are not infected with mycoplasma. 4. Some commercial trypsin reagents do not require a previous wash with PBS before trypsinization. 5. Do not leave the cells in trypsin for more than 5 min.

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Fig. 5 Representative HES staining images of a mice tumor-bearing liver. (a) Image of an HES stained liver slide. Digital magnification from 5a, of the periphery of tumors in the liver (b) and the tumor morphology (c)

6. Prepare 20% more volume of the cell suspension than the exact volume needed for injection. Use Eppendorf tubes, to allow easier collection of cells with the syringe in the moment of injection. 7. Use latex gloves, face mask, surgical gown, and hair cap as protective equipment both to protect the human operator and to keep the surgical field reasonably aseptic. 8. Experiments must comply with institutional and ethics regulations concerning the use of animals for research purposes. 9. Carry out all procedures under clean but non-sterile conditions. All the surgical instruments must be autoclaved prior to use and opened on a clean bench. Use small surgical instruments, which should be appropriate for rodent surgery. Clean surgical instruments should be used for each mouse. 10. Surgical area should be completely shaved and disinfected, to avoid contamination of the wound through contact with fur and to keep a sterile surgical field. 11. A proper abdominal incision makes the procedure easier with a good surgical view of the injection site. Improper placement may lead to a struggle delivering the liver out of the wound for injection. A smaller incision allows a better and faster mice recovery from surgery. A Colibri retractor can be used for a better visualization of the surgical field if necessary.

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12. We inject the left-lateral lobe of the murine liver as it is generally easily accessible and larger than the right or middle lobes. It is useful to use a moistened cotton tip to pull out the hepatic lobe, if necessary, pulling out the lobe with the fingers can be done, applying no pressure on the tissue. Do not use forceps to manipulate the hepatic lobes, since the liver can be lacerated and this may provoke some undesirable bleeding. 13. We have tested different gauge size needles from 31G to 25G. The larger needle would be preferred to decrease the impact of shear stress on the cells. The 25G needle did not cause any significant hemostatic or leakage problems. 14. Inject the cell suspension continuously and as slowly as possible to avoid leakage and bleeding. A precise location of the needle is crucial in this procedure to prevent backflow of the cell suspension and allow consistent and reproducible injection of the same number of cells. If the cell suspension is not properly injected under the liver capsule, a bubble will be visible. Do not insert the needle more than 2–3 mm to not cross the hepatic lobe. Do not move the needle from its initial position while injecting to prevent laceration of the liver. 15. This procedure requires surgical skill and practice to obtain reproducible results and perform the implantation in an acceptable amount of time. It is critical to inject viable cells, the tumor cells should not to be in ice for an extended period of time. 16. Possible issues for orthotopic tumor implantation are bleeding, leakage, or undesirable peritoneal metastasis. To avoid these problems, we propose (1) to reduce the volume of cell suspension injected (50 μl per injection or less), (2) to inject cell suspension as slowly as possible, (3) to extend the time applying gentle pressure with the sterile sponge after injection. 17. Clips may be used if BLI imaging is not planned on being performed during the following week, since clips may cause interference with the imaging. We normally use wound clips to close the skin, remove them after 7 days (necessary time for skin healing), and start to perform tumor growth follow-up by BLI imaging weekly. 18. An animal should not be left unattended until it has regained sufficient consciousness. An animal which has undergone a surgery should not be returned to the company of other animals until fully recovered. 19. Additionally, after removing wound clips, BLI can be utilized. Tumor growth is monitored on a weekly basis with BLI. Expression of the luciferase gene synthesizes the enzyme luciferase which allows monitoring tumor progression during the course of treatment.

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20. Before starting a large experiment, it is important to determine the proper exposure time required as to not saturate the image but have a good signal. When obtaining images with BLI, several factors should be kept in mind including mouse position, the depth of tumor in the tissue, and imaging settings [17]. These are important considerations as imaging settings may be different depending on the tumor cell line, and the location of the tumor. 21. The tumors take 2 to 8 weeks to engraft and grow enough to be analyzed [18, 19]. In our experience, 90–100% of animals have tumors after a few weeks of implantation. The mice sacrifice is performed by cervical dislocation following the institutional and ethics regulations concerning the use of animals for research purposes.

Acknowledgments GK is supported by the Ligue contre le Cancer (e´quipe labellise´e); Agence National de la Recherche (ANR) – Projets blancs; AMMICa US23/CNRS UMS3655; Association pour la recherche sur le cancer (ARC); Cance´ropoˆle Ile-de-France; Fondation pour la Recherche Me´dicale (FRM); a donation by Elior; Equipex Onco-Pheno-Screen; European Joint Programme on Rare Diseases (EJPRD); Gustave Roussy Odyssea, the European Union Horizon 2020 Projects Oncobiome and CRIMSON (grant agreement No. 101016923); Fondation Carrefour; Institut National du Cancer (INCa); Institut Universitaire de France; LabEx Immuno-Oncology (ANR-18IDEX-0001); a Cancer Research ASPIRE Award from the Mark Foundation; the RHU Immunolife; Seerave Foundation; SIRIC Stratified Oncology Cell DNA Repair and Tumor Immune Elimination (SOCRATE); and SIRIC Cancer Research and Personalized Medicine (CARPEM). This study contributes to the IdEx Universite´ de Paris ANR-18-IDEX-0001. SL and HC are supported by the China Scholarship Council (CSC, file n°. 201907060011 and file n° 201908070134, respectively). UN-R is supported by Axudas de apoio a´ etapa de formacio´n posdoutoral da Xunta de Galicia – GAIN. N°Expediente: IN606B-2021/015. Conflicts of Interest GK has been holding research contracts with Daiichi Sankyo, Eleor, Kaleido, Lytix Pharma, PharmaMar, Osasuna Therapeutics, Samsara Therapeutics, Sanofi, Sotio, Tollys, Vascage, and Vasculox/Tioma. GK has been consulting for Reithera. GK is on the Board of Directors of the Bristol Myers Squibb Foundation France. GK is a scientific co-founder of everImmune, Osasuna Therapeutics, Samsara Therapeutics, and Therafast Bio. GK is the inventor of patents covering therapeutic targeting of aging, cancer, cystic fibrosis, and metabolic disorders.

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References 1. Brown ZJ, Heinrich B, Greten TF (2018) Mouse models of hepatocellular carcinoma: an overview and highlights for immunotherapy research. Nat Rev Gastroenterol Hepatol 15: 536–554 2. Bakiri L, Wagner EF (2013) Mouse models for liver cancer. Mol Oncol 7(2):206–223 3. He L, Tian D-A, Li P-Y et al (2015) Mouse models of liver cancer: progress and recommendations. Oncotarget 6:23306–23322 4. Hernandez-Gea V, Toffanin S, Friedman SL et al (2013) Role of the microenvironment in the pathogenesis and treatment of hepatocellular carcinoma. Gastroenterology 144:512–527 5. Bour G, Martel F, Goffin L et al (2014) Design and development of a robotized system coupled to mCT imaging for intratumoral drug evaluation in a HCC mouse model. PLoS One 9:e106675 6. Richmond A, Yingjun S (2008) Mouse xenograft models vs GEM models for human cancer therapeutics. Dis Model Mech 1:78–82 7. Jung J (2014) Human tumor xenograft models for preclinical assessment of anticancer drug development. Toxicol Res 30:1–5 8. Ma C, Kesarwala AH, Eggert T et al (2016) NAFLD causes selective CD4(+) T lymphocyte loss and promotes hepatocarcinogenesis. Nature 531:253–257 9. Makarova-Rusher OV, Medina-Echeverz J, Duffy AG et al (2015) The yin and yang of evasion and immune activation in HCC. J Hepatol 62:1420–1429 10. Kasashima H, Duran A, Cid-Diaz T et al (2020) An orthotopic implantation mouse model of hepatocellular carcinoma with underlying liver steatosis. STAR Protoc 1:100185 11. Wu T, Heuillard E, Lindner V et al (2016) Multimodal imaging of a humanized

orthotopic model of hepatocellular carcinoma in immunodeficient mice. Sci Rep 6(1):35230 12. Lee TK, Na KS, Kim J et al (2014) Establishment of animal models with orthotopic hepatocellular carcinoma. Nucl Med Mol Imaging 48(2010):173 13. Fleten KG, Bakke KM, Mælandsmo GM et al (2017) Use of non-invasive imaging to monitor response to aflibercept treatment in murine models of colorectal cancer liver metastases. Clin Exp Metastasis 34:51–62 14. Brown ZJ, Heinrich B, Greten TF (2018) Establishment of orthotopic liver tumors by surgical intrahepatic tumor injection in mice with underlying non-alcoholic fatty liver disease. Methods Protoc 1:1–9 15. Reiberger T, Chen Y, Ramjiawan RR et al (2015) An orthotopic mouse model of hepatocellular carcinoma with underlying liver cirrhosis. Nat Protoc 10:1264 16. Febbraio MA, Reibe S, Shalapour S et al (2019) Preclinical models for studying NASH-driven HCC: how useful are they? https://doi.org/10.1016/j.cmet.2018. 10.012 17. Zinn KR, Chaudhuri TR, Szafran AA et al (2008) Noninvasive bioluminescence imaging in small animals. ILAR J 49:103–115 18. Kauntz H, Bousserouel S, Gosse´ F et al (2011) Silibinin triggers apoptotic signaling pathways and autophagic survival response in human colon adenocarcinoma cells and their derived metastatic cells. Apoptosis 16:1042–1053 19. Kudo Y, Sugimoto M, Arias E et al (2020) PKCλ/ι loss induces autophagy, oxidative phosphorylation, and NRF2 to promote liver cancer progression. Cancer Cell 38:247–262. e11

Chapter 2 Diethylnitrosamine Induction of Hepatocarcinogenesis in Mice Jules Sotty, Pierre Bablon, Paul-Henry Weiss, and Patrick Soussan Abstract Diethylnitrosamine (DEN) is a chemical hepatocarcinogenic agent that triggers a large array of oncogenic mutations after a single injection. Initiated hepatocytes subsequently undergo clonal expansion within a proliferative environment, rendering the DEN model a comprehensive carcinogen. In rodent studies, DEN finds extensive utility in experimental liver cancer research, mimicking several aspects of human hepatocellular carcinoma (HCC), including angiogenesis, metabolic reprogramming, immune exhaustion, and the ability to metastasize. Beyond the wealth of scientific insights gleaned from this model, the objective of this chapter is to review morphological, genomic, and immunological characteristics associated to DEN-induced HCC. Furthermore, this chapter provides a detailed procedural guide to effectively induce hepatocarcinogenesis in mice through a single intraperitoneal injection of DEN. Key words Diethylnitrosamine, Hepatocellular carcinogenesis, Mouse, Chemical carcinogen, Mutagenesis, Genotoxicity

1 1.1

Introduction Generalities

Primary liver cancer is the sixth most commonly diagnosed type of cancer and the third leading cause of cancer-related mortality worldwide [1]. Given its rising global incidence, it is likely to remain a global health concern in the coming decades [2]. Hepatocellular carcinoma (HCC) is the major subtype accounting for 75–85% of primary liver cancer [1]. Most cases arise from underlying liver disease related to chronic viral infection (hepatitis B or C), excessive alcohol consumption, or metabolic associated steatohepatitis (MASH). A cirrhotic background is found in 90% of cases [3, 4] and malignancies are often diagnosed at an advanced stage, contributing to the poor prognostic of HCC. In the past few years, next generation sequencing and singlecell omics analyses highlighted a broad heterogeneity in terms of genetic and epigenetic alterations, immune microenvironment, and

Guido Kroemer et al. (eds.), Liver Carcinogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2769, https://doi.org/10.1007/978-1-0716-3694-7_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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metabolic reprogramming that could be associated with patient outcomes [5–8]. Despite promising advancements in combining immune checkpoint inhibitors with antiangiogenic factor and other targeted therapies, the clinical management of HCC lacks mechanistic insights that could pave the way for early predictive biomarkers and efficacious personalized treatments. In the realm of cancer research, animal models stand as indispensable tools affording a lens through which to discern the intricate interplay among biological compartments and the complexity intrinsic to tumor ecosystems. Among these models, diethylnitrosamine (DEN)-induced carcinogenesis is a well-established and widely employed cornerstone in experimental liver cancer investigations. While recapitulating the multistep process of carcinogenesis, DEN can be administered in combination with other chemicals or genetic modifications to emulate a range of conditions (e.g., steatosis, fibrosis, viral infection). The objective of this chapter is twofold: (i) to summarize major phenotypic features characterizing DEN-induced HCC in mice, contemplating the model’s advantages and limitations, and (ii) to provide a reliable methodology for inducing hepatocarcinogenesis via a single DEN injection. 1.2 Molecular Mechanisms of DENInduced Hepatocarcinogenesis

Diethylnitrosamine (N-Nitrosodiethylamine, C4H10N2O), akin to other N-nitrosamines, necessitates metabolic conversion to manifest genotoxic properties. The metabolic pathway of DEN in mice mirrors that observed in human liver microsomes [9]. Its biotransformation is initiated through a cytochrome P450-dependent α-hydroxylation predominantly occurring in centrilobular hepatocytes. The hydroxylated compound is spontaneously dealkylated and ultimately decomposed into an electrophilic ethyl diazonium ion [10–12]. This ethyl diazonium species can react with various nucleophilic biomolecules, including nucleic acids, to form covalent ethyl DNA adducts. Moreover, reactive oxygen species (ROS) generated via P450 cytochrome activity contribute to additional DNA damages [13]. In the end, a single dose of DEN administered to rodents incites a large array of mutagenic lesions, affecting DNA phosphodiester bonds and inducing strand breaks, or modifying DNA bases (e.g., O4-EtdT, O6-EtdG), which can result in heritable base mispairing [12, 14, 15]. The formation of preneoplastic cell clones requires a high rate of proliferation in the liver to facilitate mutation transmission. Therefore, DEN is conventionally administered to young mice at 2 weeks of age to exploit their heightened hepatocyte proliferation rates, rendering DEN a complete carcinogen during this phase. Alternatively, it can be administered chronically (once a week), or paired with a tumor promoter (e.g., phenobarbital, carbon tetrachloride, N-nitrosomorpholine) in adult mice, where hepatocytes are quiescent [16]. The introduction of a tumor promoter can amplify the rate of carcinogenesis, albeit potentially impacting

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model reproducibility of the model and modifying tumor genotype [17, 18]. Although HCC does not arise on an inflamed liver, inflammatory mechanisms appear pivotal in promoting DEN-induced hepatocarcinogenesis. At 24 hours post-treatment (15 mg/kg), Li et al. observed an acute accumulation of inducible nitric oxide synthase (iNOS) in intrahepatic macrophages along with increased expression of interleukin (IL)-6 and IL-1β [13]. Cytokines secreted by activated Kupffer cells have been shown to incite compensatory proliferation in response to hepatocyte death, even at low doses [19–22]. Neutrophils have also been implicated in promoting DEN-mediated HCC since their depletion led to decreased ROS-mediated telomere damage and a 3.5-fold reduction in tumor burden [23]. Conversely, tumor number and size were elevated in recombination-activating gene 1 (RAG1)-/- mice devoid of T and B cells, indicating that the adaptative immune response governs early tumor formation and the growth of established tumors [24]. Additionally, natural killer T (NKT) and CD4+ T cells have been shown to inhibit DEN-induced carcinogenesis by clearing pre-malignant hepatocytes [25]. 1.3 Characteristics of DEN-Induced HCC 1.3.1

Histomorphology

Murine liver tumors exhibit histological similarities to their corresponding human counterparts [26]. At 5–6 months following DEN administration, focal lesions are easily discernable on liver sections, some of which stain positive for alpha fetoprotein (AFP) [27]. Hepatocytes in dysplastic nodules often possess a distinct cytoplasmic tinctorial affinity, differing from adjacent parenchyma cells [28]. Depending on timing and mouse strain, cytoplasmic vacuoles, pseudo-nucleoli, mitotic figures, and hyalin inclusion bodies can be observed [28–31]. While hepatocytes within initial dysplastic nodules (10–20 weeks) exhibit relative uniformity, they can display significant zonal heterogeneity at later stages. Deeper regions might appear eosinophilic or hydropic, contrasting with basophilic cells at the periphery, and potentially contain neutral lipids and necrotic areas [28] (Fig. 1). HCC ultimately arises from regenerative dysplastic nodules, where an elevated nuclear/cytoplasm ratio is indicative of malignancy [13]. HCC in mice typically assumes a trabecular pattern [32] and can be characterized through staining positive for AFP or Glypican 3 (Gpc3) [19, 27, 28]. While a single carcinogenic dose appears not to alter parenchyma architecture, chronic DEN administration (35 mg/kg once a week) can incite acute lesions, including diffuse hepatocellular hydropic degeneration, apoptosis, and necrosis [29, 31]. Prolonged exposure may lead to HCC development against a backdrop of liver inflammation and fibrosis after 25 weeks [31, 33].

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Fig. 1 Intrahepatic lesions resulting from a single injection of DEN at 5 mg/kg in 15-day-old male mice. (a) Basophilic dysplastic nodule at 6 months post injection. (b) Higher magnification of panel A. Note the increased nuclear/cytoplasmic ratio and the presence of hyaline inclusion bodies. (c) Macrotrabecular hepatocellular carcinoma at 10 months post-injection. (d) Higher magnification of panel C highlighting zonal heterogeneity within nodules, the presence of lipids, vacuolated cells, and inclusion bodies. DEN, diethylnitrosamine 1.3.2 Mutational/ Transcriptomic Landscape

Tumors originating from DEN-injected mice exhibit a high burden of somatic mutations, surpassing the average number of mutated genes in human HCC [34]. Frequent genetic mutations affect proto-oncogenes encoding constituents of the mitogen-activated protein kinase (MAPK) signaling pathways, such as epithelial growth factor receptor (EGFR), B-Raf, and H-Ras, thereby constitutively activating downstream effectors like the Raf/MEK/ERK cascade [26, 34, 35]. While Ctnnb1 mutations are not frequently encountered in the classic DEN model, the administration of DEN + phenobarbital in adult mice results in a selective clonal outgrowth of hepatocytes harboring Ctnnb1 mutations [18]. Accordingly, the transcriptomic profile of DEN-mediated tumors has been characterized by the overexpression of genes related to cell growth and proliferation, a pattern distinct from other tested mouse models. While this gene expression profile is not commonplace in human HCC, it is linked to a subclass of human HCC associated with poor survival prognosis [36].

Diethylnitrosamine Induction of Hepatocarcinogenesis in Mice 1.3.3

Immune Infiltration

1.4 General Recommendations

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Numerous studies demonstrated that a single DEN injection can trigger both innate and adaptative immune responses alongside carcinogenesis. Elevated levels of certain chemokines (i.e., C-C motif chemokine ligand [CCL]2, CCL3, CCL4, CCL5, C-X-C motif chemokine ligand [CXCL]9) have been observed in liver prior to the detection of visible nodules, underscoring the early induction of a chronic inflammatory response during hepatocarcinogenesis [24]. Ten months after exposure, Deust et al. identified intrahepatic infiltration of neutrophils, macrophages, and T cells, including programmed cell death protein 1 (PD-1)+ subtypes, along with an accumulation of T cells in the spleen. Kupffer cell, NK, and NKT numbers remained unchanged [37]. Additionally, a classical immunosuppressive phenotype was described in young mice following a single dose, marked by intrahepatic Treg infiltration [37, 38] and the accumulation of myeloid-derived suppressor cells (MDSCs) in the bone marrow, liver, and spleen [24, 39]. Chronic DEN administration typically yields more pronounced and early immune alterations. Using this model, Heindryckx et al. have observed a fourfold and eightfold increase in inflammatory foci after 16 and 20 weeks, respectively, with macrophage preferentially accumulating around HCC tumors [33]. Fifteen weeks of chronic DEN exposure in rats resulted in liver infiltration by IL-10-producing Breg and Treg, decreased expression of dendritic cell co-stimulatory molecules (i.e., major histocompatibility complex (MHC)-II, CD80, CD86), along with reduced T cell proliferative capacity [40]. Additionally, T cell exhaustion was noted at 15 weeks post-injection in a DEN + CCl4 model, characterized by increased expressions of the markers T cell immunoglobulin and mucin containing protein 3 (Tim-3), lymphocyte-activation gene 3 (LAG-3), and PD-1 [41]. Recent findings revealed that inhibiting the PD-1-PD-L1 axis could impede HCC development in mice injected with DEN + CCl4 [42, 43], suggesting that this model may be adapted for preclinical study of immunotherapies. The susceptibility to chemical hepatocarcinogenesis exhibits a distinct specificity based on mouse strains. Some of them such as C57BL/6 and BALB/c are relatively resistant compared to others, notably C3H and CBA [44]. This variation in susceptibility has been linked to differing mutation sites across mouse strains [35]. Furthermore, males have a higher incidence of HCC development than females due to the protective role of estrogen [21, 45]. To mitigate gender-related biases and reduce the number of animals in experimental procedures, it is recommended to exclusively utilize males. Among the different administration routes for DEN, intraperitoneal injection is often favored to ensure accurate dosing and enhance reproducibility. Numerous factors influence tumor incidence and progression, including manipulator, diet, genetic background, hormones, and environment. To minimize

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Table 1 List of studies employing a single injection of DEN and their corresponding incidences of HCC at various timepoints. Incidence of liver tumors is computed based on the macroscopic detection of surface nodules >1 mm. DEN, diethylnitrosamine Mouse strain

Dose

Tumor incidence

Metastasis

References

129svj

5 mg/kg

0% at 24 weeks 82% at 36 weeks

45% lung metastasis at 36 weeks

Teoh et al. [46]

C57BL/6

15 mg/kg

90 mg/kg

20% at 20 weeks 77% at 32 weeks 0% at 16 weeks 83% at 40 weeks 100% at 48 weeks 100% at 32 weeks 0% at 22 weeks 71% at 44 weeks 81% at 40 weeks 33% at 77 weeks

20 mg/kg

100% at 25 weeks

20 mg/kg

25 mg/kg

C3H

Li et al. [13] 33% lung metastasis at 72 weeks

Kapanadze et al. [39]

Naugler et al. [21] Mossanen et al. [25] No metastasis at 40 weeks 14% lung metastasis at 77 weeks

Shalini et al. [47] Diwan and Meyer [48] Connor et al. [26]

result variability, it is crucial to designate a single manipulator for injections, maintain consistent cage environments, and randomize mouse groups to the greatest extent possible. The classical protocol entails a single intraperitoneal injection in 2-week-old pups. A panel of studies adhering to this protocol and displaying corresponding incidence rates is outlined in Table 1. Liver harvesting typically occurs 6 to 12 months after a single DEN injection at doses ranging from 5 to 50 mg/kg. The emergence of initial mice harboring macroscopic tumors is typically observed around 5–6 months, though the incidence varies across mouse strains [13, 27]. However, some studies do not report surface tumors at this time point [25, 46], thereby warranting a pilot experiment with a small number of mice to fine-tune timings and group sizes. 1.5 Advantages and Limitations of the Model

The injection of DEN stands as the most commonly employed model for inducing hepatocarcinogenesis in rodents. Consequently, the scientific literature abounds with a wealth of related information, comparison points, and specific protocols. Despite being time-consuming, this model offers ease of implementation and reproducibility, effectively simulating the multistep process of carcinogenesis, spanning sporadic mutagenesis, clonal expansion, and advanced tumor progression. Sporadic mutagenesis mirrors the circumstances in HCC patients. Yet, it inherently introduces significant variability in tumor incidence and clonal diversity, thereby

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necessitating larger sample sizes to glean meaningful insights. This particularity underscores a contrast with genetically engineered mouse models where mutation sites are targetable. Furthermore, DEN-induced carcinogenesis has been demonstrated to capture several facets of human HCC, encompassing neoangiogenesis, metabolic reprogramming, immune evasion, and metastatic potential. While tumors originate from accentuated DNA damage, they do not arise in a context of underlying chronic hepatitis (typically characterized by liver inflammation and fibrosis), as often seen in human HCC cases. Nevertheless, several approaches exist to establish a fibrotic environment intertwined with chronic inflammation, such as incorporating CCl4 treatment and employing repeated DEN exposure. Overall, despite possessing certain distinct attributes (e.g., mutational landscape, absence of chronic inflammation), DEN-induced hepatocarcinogenesis remains among the most pertinent and versatile models, affording researchers the opportunity to investigate various aspects of HCC across its spectrum from early stages to advanced malignancy. 1.6

Materials

1. Disposable sterile syringes and needles (25–27 g). 2. Balance. 3. Gauzes or cotton swabs. 4. Chemical hood. 5. Chemical absorbent paper. 6. Diethylnitrosamine C4H10N2O).

(DEN,

N-Nitrosodiethylamine,

7. Phosphate-buffered saline (PBS). 8. Sharp biohazard waste container. 9. 14–15-day-old mice.

2

Methods 1. Prepare your workplan within a chemical hood, using absorbent paper. 2. Prepare a sterile DEN solution in PBS at a concentration of 0.5–2 mg/ml, in a microtube. The recommended volume for IP injection in mice is approximately 1% of the body weight. For example, for a 5 mg/kg injection in a 20 g mouse, prepare a solution at 0.5 mg/ml to inject 200 μl (see Notes 1, 2, 3, 4, 5, 6, 7). 3. Disinfect the tube containing DEN solution with 70% ethanol, and place it on a tube rack to allow it to reach room temperature. Dispose cotton swabs or gauzes, syringes, and biohazard waste container on absorbent paper.

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4. Weigh the mice, calculate and note the appropriate injection volumes. 5. If possible, position the cage under the chemical hood. Otherwise, ensure that the mice are placed under the hood during the injection (see Note 6). 6. Prewarm the syringe by moving the plunger up and down to avoid friction during injection. Draw the appropriate volume of the DEN solution into the syringe and place it horizontally on absorbent paper (see Note 8). 7. Gently pick up a mouse from the cage and immobilize it by grasping the loose skin at the back of the neck between your thumb and forefinger. 8. Insert the needle to a depth of about ½, with the bevel facing up, in the right lower quadrant of the abdomen (left when facing the animal). Slowly inject the DEN solution without moving the needle within the abdomen. Keep the needle in place for 1 second after injection before gently removing it. If a drop of liquid leaks from the injection site after removing the needle, use a cotton swab or gauze to absorb it (see Note 9). 9. Return the animal to the cage and inspect for any complications (i.e., bleeding at the injection site, peritonitis) at the end of procedure, and the next day (see Note 10). 10. Mice must be carefully monitored from the injection procedure until the time of sacrifice. Monitoring includes regular weighting and observing behavior for signs of pain and distress. It is crucial to adhere to the list of humane endpoints that has been established in conjunction with the ethical committee.

3

Notes DEN is a mutagenic and teratogenic chemical classified as a human carcinogen (group 2A) by the international agency for research on cancer (IARC). Acute exposure by inhalation or ingestion may cause irritation of the respiratory tract, nausea, vomiting, and fever. It may also cause skin and eye irritation by dermal contact. DEN should always be handled with the following prudent practices. 1. Always handle the product while wearing a white coat, airtight nitrile gloves, and safety glasses. If the product comes into direct contact with gloves, replace them. 2. DEN is highly volatile and must be manipulated under a chemical hood to prevent exposure to hazardous vapors. 3. A chemical absorbent paper should be disposed on the work plan to prevent any spillage.

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4. Label tubes and waste containers with the appropriate carcinogenic, mutagenic or reprotoxic (CMR) designation. 5. DO NOT recap syringes after use. 6. Dispose of waste items such as gloves, absorbent paper, cones, and tubes that have come into contact with DEN in sealed and labeled containers (denoting diethylnitrosamine and CMR classification). 7. Inform the personnel in the animal facility that you have administered DEN to the animals. Label the cages accordingly, wait 24 hours before changing the bedding, and place the used bedding in labeled autoclavable bags. 8. DEN is sensitive to light and should be stored protected from light at 4 °C. 9. As it is flammable, it must be kept away from sources of open flames. 10. The storage area should be designated for storing CMR chemicals and located as close as possible to the handling area. 11. Only aliquot and transport the quantity required to minimize risks associated with handling and transportation and to reduce the generation of hazardous waste. References 1. Sung H, Ferlay J, Siegel RL et al (2021) Global cancer statistics 2020: GLOBOCAN estimates of incidence and mortality worldwide for 36 cancers in 185 countries. CA A Cancer J Clin 71:209–249 2. Dasgupta P, Henshaw C, Youlden DR et al (2020) Global trends in incidence rates of primary adult liver cancers: a systematic review and meta-analysis. Front Oncol 10:171 3. Fattovich G, Stroffolini T, Zagni I et al (2004) Hepatocellular carcinoma in cirrhosis: Incidence and risk factors. Gastroenterology 127: S35–S50 4. Mittal S, El-Serag HB, Sada YH et al (2016) Hepatocellular carcinoma in the absence of cirrhosis in US veterans is associated with non-alcoholic fatty liver disease. Clin Gastroenterol Hepatol 14:124–131.e1 5. Losic B, Craig AJ, Villacorta-Martin C et al (2020) Intratumoral heterogeneity and clonal evolution in liver cancer. Nat Commun 11:291 6. Sia D, Jiao Y, Martinez-Quetglas I et al (2017) Identification of an immune-specific class of hepatocellular carcinoma, based on molecular features. Gastroenterology 153:812–826 7. Sun Y, Wu L, Zhong Y et al (2021) Single-cell landscape of the ecosystem in early-relapse

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Diethylnitrosamine Induction of Hepatocarcinogenesis in Mice different murine models of hepatocellular carcinoma. J Hepatol 59:1007–1013 40. Song S, Yuan P, Li P et al (2014) Dynamic analysis of tumor-associated immune cells in DEN-induced rat hepatocellular carcinoma. Int Immunopharmacol 22:392–399 41. Yin C, Han Q, Xu D et al (2019) SALL4mediated upregulation of exosomal miR-146a-5p drives T-cell exhaustion by M2 tumor-associated macrophages in HCC. OncoImmunology 8:e1601479 42. Chung AS, Mettlen M, Ganguly D et al (2020) Immune checkpoint inhibition is safe and effective for liver cancer prevention in a mouse model of hepatocellular carcinoma. Cancer Prev Res (Phila) 13:911–922 43. Liu C, Yang Y, Chen C et al (2021) Environmental eustress modulates β-ARs/CCL2 axis to induce anti-tumor immunity and sensitize immunotherapy against liver cancer in mice. Nat Commun 12:5725 44. Kemp CJ, Drinkwater NR (1989) Genetic variation in liver tumor susceptibility, plasma

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testosterone levels, and androgen receptor binding in six inbred strains of mice. Cancer Res 49:5044–5047 45. O’Brien MH, Pitot HC, Chung S-H et al (2021) Estrogen receptor-α suppresses liver carcinogenesis and establishes sex-specific gene expression. Cancers 13:2355 46. Teoh NC, Dan YY, Swisshelm K et al (2008) Defective DNA strand break repair causes chromosomal instability and accelerates liver carcinogenesis in mice. Hepatology 47:2078– 2088 47. Shalini S, Nikolic A, Wilson CH et al (2016) Caspase-2 deficiency accelerates chemically induced liver cancer in mice. Cell Death Differ 23:1727–1736 48. Diwan BA, Meier H (1975) Carcinogenic effects of a single dose of diethylnitrosamine in three unrelated strains of mice: Genetic dependence of the induced tumor types and incidence. Cancer Letters 1:249–253

Chapter 3 Diethylnitrosamine-Induced Liver Tumorigenesis in Mice Under High-Hat High-Sucrose Diet: Stepwise High-Resolution Ultrasound Imaging and Histopathological Correlations Pierre Cordier, Flora Sangouard, Jing Fang, Christelle Kabore, Chantal Desdouets, and Se´verine Celton-Morizur Abstract The hepatotoxic N-nitroso compound diethylnitrosamine (DEN) administered intraperitoneally (i.p.) induces liver neoplasms in rodents that reproducibly recapitulate some aspects of human hepatocarcinogenesis. In particular, DEN drives the stepwise formation of pre-neoplastic and neoplastic (benign or malignant) hepatocellular lesions reminiscent of the initiation-promotion-progression sequence typical of chemical carcinogenesis. In humans, the development of hepatocellular carcinoma (HCC) is also a multistep process triggered by continuous hepatocellular injury, chronic inflammation, and compensatory hyperplasia that fuel the emergence of dysplastic liver lesions followed by the formation of early HCC. The DEN-induced liver tumorigenesis model represents a versatile preclinical tool that enables the study of many tumor development modifiers (genetic background, gene knockout or overexpression, diets, pollutants, or drugs) with a thorough follow-up of the multistage process on live animals by means of highresolution imaging. Here, we provide a comprehensive protocol for the induction of hepatocellular neoplasms in wild-type C57BL/6J male mice following i.p. DEN injection (25 mg/kg) at 14 days of age and 36 weeks feeding of a high-fat high-sucrose (HFHS) diet. We emphasize the use of ultrasound liver imaging to follow tumor development and provide histopathological correlations. We also discuss the extrinsic and intrinsic factors known to modify the course of liver tumorigenesis in this model. Key words Diethylnitrosamine (DEN), Non-alcoholic fatty liver disease (NAFLD), Hepatocellular carcinoma (HCC), Hypercaloric diet, Mouse model, Ultrasound, Histopathology

1

Introduction Hepatocellular carcinoma (HCC) is one of the most common malignancies in humans and a major threat to public health due to a rising global incidence with 905,677 new cases in 2020 [1].

Pierre Cordier, Flora Sangouard and Jing Fang contributed equally. Guido Kroemer et al. (eds.), Liver Carcinogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2769, https://doi.org/10.1007/978-1-0716-3694-7_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Well-characterized infectious and environmental risk factors for HCC development include chronic hepatitis B virus (HBV) or hepatitis C virus (HCV) infection, long-term excessive alcohol intake, chronic aflatoxin B1 exposure, and chronic overfeeding and/or sedentary lifestyle-associated obesity, which promotes non-alcoholic fatty liver disease (NAFLD) [2]. Epidemiological studies clearly indicate that chronic liver diseases (CLD) and cirrhosis represent a fertile soil for HCC emergence and underlie more than 80% of the cases [3]. Starting from a cirrhotic liver, human hepatocarcinogenesis is widely recognized as a protracted, multistep process defined by a precise sequence of lesions. Indeed, accumulating somatic DNA alterations accompany the evolution from low-grade dysplastic nodule (L-DN) to high-grade dysplastic nodule (H-DN), early HCC, and finally progressed and advanced HCCs [4]. Interestingly, among other dysplastic features, H-DNs frequently show evidence of small cell change (smaller hepatocytes with increased nuclear-tocytoplasmic ratio and mild nuclear hyperchromasia) and large cell change (larger hepatocytes with enlarged pleomorphic nuclei) [5]. These premalignant hepatocellular changes are the earliest morphologic alterations recognized during hepatocyte transformation toward full-blown HCC. Extensive research in the field has made it possible to gain insights into the biological mechanisms driving HCC development and progression. Interestingly, the molecular alterations during hepatocarcinogenesis depend to some extent on the initial genotoxic insult and underlying etiology (reviewed by [6]). Pioneer work using whole-exome sequencing (WES) identified recurrent driver mutations in TERT (60%, promoter mutations), CTNNB1 (37%, mostly activating point mutations), and TP53 (24%, mostly inactivating point mutations) [7]. More recently, HCCs have been separated into two major molecular classes (proliferative versus non-proliferative) based on genomic, epigenomic, and signaling pathway analyses. Importantly, each class correlates with specific histopathological hallmarks, immunological features, etiologies, and finally clinical outcomes [2]. NAFLD and its more severe form, non-alcoholic steatohepatitis (NASH), have emerged as leading causes of chronic liver injury and HCC in the West [8]. The initial step is characterized by excessive hepatocellular lipid accumulation (NAFL, steatosis) that progressively sets the stage for lipotoxic liver damage in the context of liver insulin resistance [9]. According to the multiple hit hypothesis, the synergistic effects of lipotoxicity, oxidative stress, endoplasmic reticulum (ER) stress, and mitochondrial dysfunction induce hepatocyte cell death, chronic inflammation, and progressive fibrosis of the liver parenchyma [10]. In particular, hepatocyte ER stress cooperates with TNF signaling to promote de novo lipogenesis, toxic lipid production, and liver necroinflammation, illustrating

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one of the many self-amplifying loops perpetuating and aggravating NASH disease [11, 12]. In this noxious environment, attempt at proliferation of surviving hepatocytes to compensate for cell loss exacerbates DNA damage and is thought to be a prerequisite for HCC development [13]. Given the partial comprehension of the pathogenesis and the lack of effective treatment for NASH, relevant NAFLD-driven HCC mouse models are still urgently needed. Murine models of HCC development under NAFLD settings can be broadly categorized into diet-induced with or without the use of chemicals, genetically engineered, or mixed [14]. Among toxin and diet-based models, N-Nitrosodiethylamine, better known as diethylnitrosamine (DEN), coupled to a hypercaloric, obesogenic diet is one of the most popular and best-studied combinations [15]. DEN is a classic intrinsic, indirect-acting hepatotoxin widely used as a procarcinogen in rodents [16]. After intraperitoneal injection, it is transported to the liver by the portal vein and undergoes an important first pass effect. Bioactivation occurs predominantly in centrilobular hepatocytes in a cytochrome P450 2E1 (CYP2E1)-dependent manner and leads to the emergence of a highly reactive ethyl diazonium ion intermediate [17]. This transient electrophilic metabolite acts as an alkylating agent and exerts its toxic effect by DNA-adduct formation, resulting in ethylation of nucleobases like N7- and O6-ethylguanine and O4-ethylthymine. Ensuing base mispairing or abasic site formation leads to a high burden of stochastic point mutations due to base substitution, chiefly T > A transversion and T > C and C > T transitions [18, 19]. Reactive oxygen species (ROS) generated by CYP2E1 activity are also thought to significantly contribute to oxidative DNA damage upon DEN administration [20]. Combined with high-fat diet (HFD) feeding as a promoter, a single intra-peritoneal injection of DEN (25 mg/kg) to 14-day-old mice consistently induces hepatocellular tumors with a prevalence rate approaching 100% in males after 34 weeks [21]. Reminiscent of the multistep hepatic carcinogenesis in humans, multiple welldefined preneoplastic liver lesions as well as benign and malignant neoplasms can be observed sequentially in mice by conventional histology [22]. Female mice appear to be at least partially refractory to DEN-induced tumorigenesis due to the protective effect of estrogen-mediated inhibition of IL-6 signaling [23]. Indeed, circulating estrogen inhibits the early secretion of IL-6 by Kupffer cells exposed to damaged hepatocytes, thus reducing the initial proliferation of surviving hepatocytes bearing DNA alterations. These findings underline the major involvement of innate immunity in liver tumorigenesis. Here, we describe a detailed protocol for the induction of hepatocellular neoplasms in wild-type C57BL/6J mice following i.p. DEN injection (25 mg/kg) at 14 days of age and 36 weeks of feeding a high-fat high-sucrose (HFHS) diet (Fig. 1). We

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Fig. 1 Overview of the experimental procedure. C57BL/6J mice are given a single intra-peritoneal injection of DEN (25 mg/kg) at 14 days of age. Standard chow is replaced by HFHS diet one week after weaning (week 5) and mice are fed ad libitum for 36 weeks. Ultrasonographic follow-up can be started at 5 months after the initial carcinogenic challenge and performed monthly until euthanasia at 9 months post-DEN exposure (week 41). We recommend that the frequency of ultrasound examinations be increased in the last two months (e.g., every two weeks) because of the unpredictable behavior of DEN-induced neoplasms. Mice should be weighed regularly during the experimental procedure to monitor obesity and tumor development. DEN: Diethylnitrosamine, HFHS: High-Fat High-Sucrose

monitored the stepwise development of hepatocellular tumors in this mouse model by ultrasonographic and histopathological liver examinations. This ultrasonographic approach is a versatile, non-invasive and highly sensitive technique that can be easily integrated to the routine evaluation of mice with a little practice [24]. Furthermore, it contributes to refine the use of animals in cancer research by avoiding unnecessarily high tumor burden or multiplicity and consequently animal distress, which are common disadvantages of chemical carcinogenesis models and are easily adaptable to most genetic backgrounds, including genetically engineered mice.

2 Materials 2.1

DEN Injection

1. 14-day-old (see Note 1) C57BL/6J male mice (see Note 2). 2. N-Nitrosodiethylamine (density 0.95 g/mL) (see Note 3). 3. Personal protective equipment (nitrile gloves, goggles, lab coat) (see Note 4) and a laboratory fume hood (see Note 5). 4. Physiological saline serum. 5. 1.5 mL microcentrifuge tubes. 6. Insulin syringes (U-100, 0.5 mL, 29G).

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7. Disposable plastic cages, water bottles, and feeders. 8. Container dedicated to hazardous chemical waste (see Note 6). 2.2 High-Fat HighSucrose Diet

2.3 Ultrasonographic Equipment

High-fat high-sucrose diet (SAFE, #U8954 v.0141) (see Note 7), composed of 23.2% crude fat and 35.8% sugars (49.8% nitrogen free extract). According to the Atwater formula, diet energy is 4760 kcal/kg, 43.8% of which derives from lipids and 41.8% from nitrogen free extract. 1. Ultrasound imaging system equipped with a high-frequency linear array probe (synonym: transducer) adapted to laboratory animals. Here, we illustrate the use of the VEVO® 3100 LT Imaging System (FUJIFILM, VisualSonics) with the highfrequency linear array MX550D transducer (central frequency: 40 MHz, axial resolution: 40 μm) (Fig. 2a) (see Note 8).

Fig. 2 Workstation, mouse preparation and transducer positioning. (a) The workstation consists of a gas anesthesia system (1), an imaging station setup (2), and an ultrasound imaging system (3). The imaging station presented here includes a monitoring tablet display (4) connected to a heated platform unit with ECG electrode pads (5) and a rectal temperature probe (6). The linear array ultrasound probe (7) is linked up to an integrated rail base (8) via a transducer mounting system (9). (b) Once properly anesthetized, the mouse is transferred to a nose cone in dorsal recumbency and supine position (1). Depilatory cream is spread on the skin over the cranial abdomen (2) and the area is thoroughly wiped clean to remove the hair (3). (c) The probe is positioned just below the ribcage in transverse orientation on top of a layer of transmission gel. Note that the transducer orientation indicator (red arrow) is directed toward the right side of the animal. ECG: Electrocardiogram

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2. Imaging station setup including a heated platform unit with an animal physiological monitoring system to control the body temperature and basic physiologic parameters (e.g., heart rate respiratory rate, and possibly electrocardiogram). We use the VEVO® Animal Monitoring System fully equipped with the heated SM200 platform with built-in ECG electrode pads, a rectal temperature probe, and a tablet display unit (Fig. 2a). An integrated rail base and a transducer mounting system may be added to the imaging station setup (see Note 9). 3. Laboratory animal gas anesthesia system equipped with an induction chamber, a nose cone, and a waste-gas scavenging canister (TEMSEGA, Minihub® V2.1) (Fig. 2a) (see Note 10). 4. Isoflurane (PIRAMAL Critical Care, ISO-VET®, 100%, 1000 mg/g) (see Note 11). 5. Commercial depilatory cream, widely available as an over-thecounter product. 6. Lubricating gel (PARKER, Aquagel®, #57-05) (see Note 12). 7. Ophthalmic liquid gel (TVM, Ocry-gel® with carbopol 980 NF) (see Note 12). 8. Electrode gel for electrocardiogram (PARKER, Signa Gel®, #15-60) (see Note 12). 9. Ultrasound transmission gel 100, #01-02) (see Note 12).

(PARKER,

Aquasonic®

10. Surface disinfectant without alcohol and with neutral pH. 11. Common disposables: cotton buds, medical adhesive tape, gauzes. 2.4 Necropsy and Liver Sampling for Histology

1. Personal protective equipment (nitrile gloves, goggles, lab coat) and a laboratory fume hood (see Note 5). 2. Laboratory animal weighing scale. 3. Insulin syringes for blood collection (U-100, 0.5 mL, 29G). 4. Ethanol 70% v/v pure (see Note 13). 5. Basic necropsy instruments including a dissection board, scissors, forceps, scalpel, cotton buds, and a ruler. 6. Common plastic materials: cryogenic tubes, 30 mL sample containers, 1.5 mL microcentrifuge tubes, histology embedding cassettes, histological glass slides. 7. Liquid nitrogen (see Note 14). 8. Formaldehyde solution 4% w/v buffered at pH 6.9 (see Note 15). 9. Container dedicated to hazardous chemical waste (see Note 6).

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10. Plastic bags for mouse cadaver disposal. 11. Physiological saline serum. 12. Digital photographic equipment to take photographs of livers.

3

Methods

3.1 DEN Preparation and Administration to Animals

1. Dilute the DEN stock solution (density 0.95 g/mL) in physiological saline serum to obtain a working solution of 1.25 mg/ mL (see Note 16). Due to DEN toxicity, this step should be performed carefully with adequate personal protective equipment under a dedicated laboratory fume hood (see Note 17). 2. Weigh the 14-day-old C57BL/6J male mice before injection (week 2). The corresponding volume of working solution is subsequently calculated for each pup to reach a dose of 25 mg/ kg body weight (see Note 18). 3. Gently restrain mouse pups by the scruff and perform an intraperitoneal (i.p.) injection with the adequate volume of working solution (see Note 19). It is recommended that the injection be performed in the animal’s caudal right abdominal quadrant, laterally to the midline, using an insulin syringe with the needle bevel up (see Note 20). 4. Discard the tubes and insulin syringes in an appropriate container for hazardous chemical waste at the end of the experiment (see Note 6). 5. Put the DEN-injected mouse pups in a disposable plastic cage with the lactating mother. They should be inspected one hour after injection and at regular intervals (see Notes 21 and 22). 6. In the following days, urine and feces from injected animals and plastic cages need to be collected and disposed as hazardous waste (see Note 23). 7. After 5 days, the pups and their mother can be put back in conventional cages. Since DEN-induced acute liver damage can have a negative impact on growth rate, late weaning at 4 weeks of age is recommended (see Note 24).

3.2 High-Fat HighSucrose Feeding

1. Replace the standard chow by HFHS diet one week after weaning (week 5). 2. Feed mice ad libitum for 36 weeks until euthanasia at 9 months post-DEN exposure (week 41) (see Note 25). 3. Monitor the body weight at regular intervals to follow the development of dietary obesity and liver tumors (Fig. 1) (see Note 26).

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Note that the duration of HFHS promotion may be adjusted according to your personal workflow and the length of the tumor latency period in your conditions (see Subheading 3.5). 3.3 Ultrasonographic Assessment of the Liver 3.3.1 Work Station Preparation

1. Set up the connection between the tablet display unit and the heated platform unit for real-time animal monitoring. The heated platform should be adjusted at 38 °C (Fig. 2a) (See Note 27). 2. Prepare the gas anesthesia station by switching on the air delivery system and the waste-gas scavenger canister. 3. Verify that the vaporizing chamber is properly filled with isoflurane and set the air flow at 1.5–2 L/min and isoflurane vaporization at 3–4% for anesthesia induction. 4. Once the system is saturated with fresh gases, put 2 to 3 mice in the induction box at the same time to achieve general anesthesia and transfer one to the nose cone for imaging (see Note 28). 5. Set the isoflurane vaporization at 1.5–2% for anesthesia maintenance during mouse preparation and ultrasound examination (approximately 15–20 minutes per animal, see Note 29). 6. Switch on and initialize the ultrasound imaging system.

3.3.2

Mouse Preparation

1. The mouse should be placed in dorsal recumbency with the head toward the top of the platform (Fig. 2b). 2. Connect the electrocardiogram monitoring equipment to the animal. If using a heated platform with built-in ECG electrode pads, apply first ECG gel between each paw and its respective electrode. 3. Immobilize the mouse in supine position on the heated platform by applying medical adhesive tape on the four limbs. Insert with great care the temperature probe in the rectum using lubricating gel (see Note 30) and tape the base of the probe to the platform. 4. Apply ophthalmic gel to lubricate the eyes during the anesthetic procedure. 5. For optimal liver ultrasonographic evaluation, hair from the cranial abdominal region needs to be removed beforehand. Use a cotton bud to spread the depilatory cream over the cranial abdomen and wait for 1–2 minutes before removing the hair with a gauze (Fig. 2b). Wipe thoroughly the area clean with warm water and a wet gauze after hair has been removed (see Note 31). 6. Finally, apply a sufficient amount of prewarmed ultrasound transmission gel (1 cm thick band of gel is enough) on the skin over the cranial abdomen (see Note 32).

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The mouse should be closely monitored during the whole imaging procedure (body temperature, heart rate, respiratory rate) (see Note 33). 3.3.3 Transducer Positioning and Ultrasound Settings

1. Position first the transducer in transverse orientation, on the ultrasound gel, just below the ribcage and start imaging in B-mode (Fig. 2c). 2. Apply only a slight pressure on the abdomen at the beginning to visualize the most ventral liver parenchyma (just beneath the skin). A higher pressure can then be applied to better scan the deepest dorsal regions of the liver. Sagittal views can be obtained by rotating the transducer 90°. The transducer orientation indicator has to be orientated correctly, both on the probe and on the image display (Fig. 2c). As a convention, this probe orientation marker needs to be directed toward the animal’s right in transverse views (animal’s right on the left side of the screen) and toward the head in sagittal views (animal’s head on the left side of the screen). The imaging station setup may include an integrated rail base and a transducer mounting system to secure the transducer in stationary position, allow fine movements of the probe with the micro-manipulators, and achieve increased stability during image acquisition and measurements (Fig. 2a). The screen display can be optimized by adjusting the field of view to fit the region of interest (e.g., liver parenchyma, tumor), the focus depth (depth of the focal zone), and the gain (intensity of echo amplification). We recommend to keep constant the ultrasound settings between mice and imaging sessions in longitudinal studies and save the set of acquisition parameters in a custom preset (see Note 34).

3.3.4 B-Mode Imaging and Ultrasonographic Description

The liver parenchyma should be scanned in B-mode (brightness mode) to assess morphology (see Note 35). This mode provides a two-dimensional (2-D) real-time ultrasound image display composed of bright dots on a gray scale, representing the ultrasound echo signal amplitude [25]. B-mode is the most effective imaging representation for locating anatomical structures as well as for describing lesions and taking measurements. The description of anatomical structures and lesions should include: shape, size, location in the abdomen or organ, echostructure (homogeneous or heterogeneous), echotexture (finely to coarsely granular), echogenicity, and presence of ultrasound artefacts [26]. Structures that appear brighter than surrounding tissues are said to be hyperechoic (e.g., liver capsule) whereas darker structures vary from hypoechoic (e.g., lumen of blood vessels) to anechoic (e.g., gallbladder content).

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3.3.5 Ultrasound Examination Cranial Abdomen Imaging and Anatomical Landmarks

The ultrasonographic assessment of the liver should be systematic and follow a consistent approach. 1. Locate the liver in the cranial abdominal cavity, immediately behind the diaphragm and examine its relationships with adjacent organs in transverse imaging. 2. In the median plane, close to the hilus, find the major abdominal blood vessels: abdominal aorta (most dorsal vessel with hyperechoic, pulsating walls), caudal vena cava (largest vessel, running laterally to the aorta on its right side, with thin walls and renal vein branching), and portal vein (most ventral vessel with hyperechoic walls, in close contact with the liver parenchyma and a caliber close to that of the aorta) (Fig. 3a). 3. Move the transducer to the right side of the cranial abdomen and locate the right kidney which contacts the caudate process of the caudate lobe of the liver at the level of the renal fossa (Fig. 3b) (see Note 36). 4. In the left cranial abdomen, find the stomach which contacts the visceral surface of the left lateral lobe of the liver (a large sac-like structure frequently filled with hyperechoic gas and feed content creating acoustic shadowing artefacts) and continues with the duodenum, running transversally from the pylorus to the right side of the abdominal cavity (Fig. 3a, c). 5. Finally, visualize the left kidney situated caudally to the stomach and medially to the spleen (Swiss-cheese appearance). The latter contacts the liver dorsally and the pancreas medially (hyperechoic, striated appearance).

Normal Liver Ultrasonography

The normal liver parenchyma appears homogeneously hypoechoic to slightly hyperechoic in nonfasted mice (glycogen accumulation) with an intermediate granular echotexture (Fig. 3a, b). Individual lobes are generally not easily identifiable in B-mode (see Note 37) but may sometimes be seen when differently involved by a disease process (e.g., steatosis, severe inflammation, or fibrosis). Examine successively all the compartments of the liver: 1. The margins of the liver should be smooth, regular, and delineated by a curved hyperechoic line (liver capsule) (Fig. 3c). The tip of liver lobes should appear sharp and pointy. 2. Intra-hepatic blood vessels are seen as round (cross-section) or branching tubular (longitudinal section) hypoechoic to anechoic structures that can be followed by moving the transducer along their course. The walls of portal vein branches are hyperechoic due to the presence of more abundant surrounding connective tissue (Fig. 3c).

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Fig. 3 Normal liver ultrasonography and anatomic landmarks. (a) Transverse view of the cranial abdomen in the median plane. Major abdominal blood vessels are visible dorsally as well as the stomach in the left abdomen with a hyperechoic content. The duodenum is seen running from the left to the right side and the surrounding liver appears hypoechoic. (b) Transverse view of the right cranial abdomen. The liver is visible ventrally (right lobe) and contacts the right kidney (blue arrowheads) at the level of the renal fossa. Note that the kidney cortex is isoechoic to the caudate process of the caudate hepatic lobe. Major abdominal vessels are seen on the right side of the image. (c) Transverse view of the left cranial abdomen. The majority of the field is occupied by the stomach which has a heterogeneous echostructure due to gas and feed content. Note intrahepatic blood vessels with hyperechoic walls (portal vein branches) in the left lateral liver lobe. (d) Sagittal view of the cranial abdomen in the median plane. The liver is visible cranially with a hypoechoic parenchyma and sharp extremities. The gallbladder can be seen on the left side of the image as a hollow organ with an anechoic content. Large abdominal veins are observable dorsally. AA: Abdominal Aorta, CP: Caudate Process, CVC: Caudal Vena Cava, D: Duodenum, GB: Gallbladder, L: Liver, PV: Portal Vein, RK: Right Kidney, S: Stomach. Blue dot at the top left corner indicates right lateral side and cranial in transverse and sagittal orientations, respectively

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3. The gallbladder is visible cranially as an anechoic oval to pearshaped hollow organ with a thin, slightly hyperechoic wall and produces a posterior acoustic enhancement artefact (Fig. 3d). 4. Intra-hepatic bile ducts are not identifiable. A similar ultrasonographic evaluation can be performed in sagittal orientation (Fig. 3d). For further analyses (annotations and measurements), save B-mode frames and video clips of transverse and sagittal views. Pathologic Ultrasonographic Liver Findings

For a comprehensive monitoring of liver tumor development, ultrasound follow-up may be started 5 months after the initial carcinogenic challenge. At the beginning, the slow growth of liver tumors authorizes a monthly ultrasonographic evaluation. Starting from 7 months to 9 months post-DEN exposure (time of euthanasia), we recommend that the frequency of ultrasound examinations be increased (e.g., every two weeks) as DEN-induced liver tumors may grow very rapidly and sometimes unpredictably (Fig. 1). In every case, cumulative tumor volume should not exceed 2 cm3 in any single mouse and signs of distress call for a decision of euthanasia (humane endpoint). HFHS-induced obesity is accompanied by a massive liver triglyceride overload responsible for hepatomegaly (hepatic steatosis) (see Note 38). On ultrasound images, diffuse lipid accumulation increases the overall echogenicity of the liver to the point that the hepatic parenchyma appears brighter than the renal cortex (liver parenchyma and kidney cortex are normally isoechoic) [24]. The site of anatomical apposition of the caudate process of the caudate liver lobe to the right kidney at the level of the renal fossa offers the opportunity for size and echogenicity comparison between the two organs (Fig. 4a). Quantitative and accurate assessments of hepatic steatosis can be obtained at that site by measuring the ultrasound hepatic/renal echo-intensity ratio (average hepatic gray scale divided by average renal gray scale) in sagittal or transverse views [27]. However, in our experience, we frequently see variations in echogenicity and thus steatosis between individual liver lobes. Increased acoustic attenuation with poor visualization of deep dorsal regions and loss of echoes from the walls of portal vein branches are additional ultrasonographic features of diffuse hepatic steatosis. Fibrosis progressively disrupts liver architecture and creates a heterogeneous echostructure with multifocal hyperechoic spots corresponding to fibrotic areas. When severe diffuse hepatic fibrosis occurs, the capsule is thickened with an irregular outline, liver lobes are retracted, and the organ appears smaller than normal (microhepatica) with portal vein widening [28]. Occasionally, anechoic unilocular or multilocular thin-walled biliary cysts can be seen.

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Fig. 4 Pathologic ultrasonographic liver findings in the DEN-HFHS mouse model. (a) Diffuse hepatic steatosis. The caudate process of the liver is markedly enlarged and distinctly hyperechoic compared to the right kidney cortex. Transverse view. (b) Pre-neoplastic focus. The liver shows a small, hyperechoic focal lesion with irregular outlines and no compression of the surrounding parenchyma. Note the close association with a small blood vessel on its right side. Transverse view. (c) Pre-neoplastic focus. A round, hypoechoic, and non-compressive focal lesion in the right part of the liver. Transverse view. (d) Hepatocellular adenoma. A large, round, well-demarcated, and hypoechoic benign tumor causing distinct compression of the adjacent liver parenchyma. The expansive tumor growth impinges on surrounding blood vessels. Transverse view. (e) Hepatocellular adenoma. A large, round, and expansive tumor isoechoic to adjacent liver and with ill-defined margins. Note the bending of the dorsal blood vessel with a compressed lumen due to tumor expansion. Transverse view. (f) Hepatocellular adenoma. A small liver neoplasm with a hyperechoic center surrounded by a narrower hypoechoic periphery, giving it the typical target appearance. Transverse view. (g) Hepatocellular carcinoma. A large malignant tumor with ill-defined boundaries and a heterogeneous echostructure. Note the variegated appearance with hyperechoic areas and multiple anechoic foci corresponding to multifocal necrosis (*). Sagittal view. (h) Hepatocellular carcinoma. A second example with a similar ultrasonographic aspect as in G. The lesion shows a prominent heterogeneous echogenicity and effaces an entire lobe. Transverse view. (i) Hepatocellular carcinoma. A smaller, round, and well-demarcated malignant hepatic neoplasm arising in the papillary process of the caudate lobe. Several anechoic cavities can be seen (*). Transverse view. AA: Abdominal Aorta, CP: Caudate Process, CVC: Caudal Vena Cava, L: Liver, PV: Portal Vein, RK: Right Kidney, S: Stomach. Blue dot at the top left corner indicates right lateral side and cranial in transverse and sagittal orientations, respectively

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DEN-induced liver tumorigenesis occurs sequentially with a stepwise tumor development over time (see Note 39): (a) Pre-neoplastic lesions manifest as small, focal, generally 1 cm in diameter but is not a reliable criterion per se to distinguish pre-neoplastic, benign, and malignant lesions. (c) Hepatocellular carcinomas (HCCs) present as large, frequently multinodular, compressive, and/or multifocally invasive malignant growths (Fig. 4g, h, and i). The echogenicity is most often variegated and highly heterogeneous with hypo/ anechoic cavities corresponding to angiectasis, necrosis, or hemorrhages intermingled with hyperechoic areas due to patchy lipid accumulation or abundant connective tissue (Fig. 4i) (see Note 41) [26]. HCCs are generally more than 1 cm in diameter, sometimes effacing entire lobes, and the surrounding non-tumoral liver parenchyma may show decreased echogenicity due to lipid and glycogen depletion. 3.3.6 Post-Imaging Mouse Care

1. Put back the transducer on its holder with great care (see Note 42) and stop the ultrasound scanning. 2. Carefully remove the rectal temperature probe and wipe off the ultrasound gel with a wet gauze. 3. Withdraw the adhesive tape on the mouse limbs and let the animal recover from anesthesia on a heating source. 4. Monitor the mouse for full recovery before placing it back into its original cage (see Note 21). 5. At the end of the session, turn off the gas anesthesia system and the ultrasound imaging system.

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6. Use an alcohol-free disinfectant to clean the ECG electrodes, the temperature probe, and the ultrasound transducer (see Note 43). Ultrasound images can be saved on an external storage disk (DICOM file format) and visualized with an adequate medical imaging software. 3.4 Necropsy and Histological Sample Collection 3.4.1

Sampling

1. Prepare the work station and annotate cryogenic tubes, Eppendorf tubes, and sample containers with the mouse identification number, sample date, and nature of the sample (e.g., liver tumor, non-tumoral liver, blood). 2. Weigh the mouse and collect venous blood according to your usual sampling route if required (see Note 44). Animals should be humanely euthanized in agreement with your institution’s approved procedures and the necropsy workflow should be conducted in a systematic manner (see Note 45) [29]. 3. Dampen the hair coat with 70% ethanol, open the abdominal cavity by incising the wall on the midline, and expose the cranial abdomen (see Note 46). 4. Excise entirely the liver by cutting its retro-diaphragmatic ligamentous attachments and the caudal vena cava before its entry into the thoracic cavity (see Note 47). 5. Blot the liver against an absorbent material such as a wet gauze to remove excess blood (see Note 48). 6. Tumor load should be evaluated by weighing the liver (see Note 49) and counting the number of all macroscopic tumors visible from the surface of the organ (see Note 50). Given the large number of neoplastic lesions usually present, in our practice, we also specifically count the number of tumors larger than 5 mm in diameter and briefly describe their gross morphology (lobar distribution, size, shape, color, consistency). 7. Recording macroscopic features may prove useful for interpreting histopathological findings (e.g., tumor necrosis and hemorrhage as indicators of malignancy). It is recommended that photographs be taken of the diaphragmatic and visceral surfaces of the liver (see Note 51). 8. Other organs may be inspected for additional lesions or metastases (e.g., lung, abdominal cavity), especially in susceptible strains or genetically engineered mice. 9. Separate individual liver lobes at their base. Cut lobe slices encompassing both tumors and surrounding non-tumoral liver parenchyma with a scalpel blade and immerse them immediately in 10% formalin for fixation (see Note 52). Each lobe should be sampled for histology to take account of interlobar heterogeneity.

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10. Sample multiple sites of non-tumoral liver for molecular analyses (e.g., RNA and protein extractions) in a cryogenic tube snap-frozen in liquid nitrogen. The largest tumors (e.g., >5 mm in diameter) may be bisected along their longest axis and half of it fixed in 10% formalin with associated surrounding non-tumoral liver parenchyma and the other half cut in smaller pieces for cryopreservation in liquid nitrogen (molecular analyses). 11. Alternatively, tumors and non-tumoral liver may be employed for a variety of experimental assessments according to your custom workflow (e.g., flow cytometry, frozen tissue preparation for cryosectioning). 12. At the end of the experiment, cryogenic tubes should be transferred from liquid nitrogen to a - 80 °C freezer for long-term storage or processed immediately according to your usual protocol. 13. After adequate fixation (see Note 52), transfer liver samples from formalin-filled containers to annotated histology embedding cassettes and immerse the latter in 70% ethanol. Due to formalin toxicity, this step should be performed with adequate personal protective equipment under a dedicated laboratory fume hood. Discard formalin in an appropriate container for hazardous chemical waste or store it for a second use (see Note 6). 14. After paraffin embedding, cut 3-μm-thick liver tissue sections with a microtome for conventional hematoxylin and eosin (H&E) staining and histopathology (see Note 53). 3.4.2 Interpretation of Histopathological Liver Findings

The histopathological assessment of neoplastic and non-neoplastic liver lesions is best performed with an upright brightfield microscope equipped with 4X, 10X, and 40X objectives (see Note 54). On H&E-stained slides, microscopic structures with an affinity for hematoxylin are said to be basophilic (violet-blue) whereas those structures that stain with eosin are referred to as acidophilic or eosinophilic (pink-red). To guarantee the reproducibility of histopathological findings [30], the description and diagnosis of mouse liver lesions should strictly follow the International Harmonization of Nomenclature and Diagnostic Criteria (INHAND) guidelines for Lesions of the Rat and Mouse Hepatobiliary System, as comprehensively defined by Thoolen and colleagues [31], and the accepted consensual terminology used in mouse pathology, as published by international committees and experts [32]. HFHS-induced hepatic steatosis is usually severe, involving all liver lobes with a prominent centrilobular (zone 3) to panlobular (panacinar) zonal pattern. Hepatocytes contain either a single large intracytoplasmic vacuole displacing the nucleus to the periphery

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(macrovesicular steatosis) or numerous smaller vacuoles approximately the size of the nucleus with no peripheralization of the latter (mediovesicular steatosis). True microvesicular steatosis, in which hepatocytes are filled with innumerable minute vacuoles giving the cytoplasm a foamy appearance, is seldom observed. Multifocal aggregates of lymphocytes, macrophages, and neutrophils are seen within liver lobules (lobular inflammation) along with mild to moderate portal inflammation. Isolated or small clusters of pigment-laden macrophages are sometimes visible. Hepatocellular injury is generally mild to moderate and includes cytoplasmic alteration (increased granularity, swelling, and eosinophilia), a higher occurrence of hepatocellular apoptosis (rounded, condensed hyperacidophilic cytoplasm surrounded by a clear halo and with pyknotic hyperbasophilic nucleus), and more infrequently, hepatocyte karyomegaly with intra-nuclear cytoplasmic invagination (pseudoinclusion). Bona fide hepatocellular ballooning and Mallory-Denk bodies, major features of human NASH, are not typical responses of mouse hepatocytes to chronic metabolic injury and are thus rarely seen, except when severe chronic steatohepatitis occurs (see Note 55). Mild to moderate centrilobular (zone 3) perisinusoidal fibrosis and periportal (zone 1) oval cell hyperplasia (rows of oval cells streaming from portal tracts into liver lobules) may be part of the histopathological findings. NAFLD hallmark lesions may be semi-quantitatively assessed using the histological grading (activity) and staging (chronicity) systems defined by Kleiner and collaborators for the human disease [33]. Several DEN-induced hepatocellular proliferative lesions can be recognized: (a) Foci of cellular alteration (FCA) represent early DEN-induced pre-neoplastic liver lesions that have been well characterized histologically [31]. Those dysplastic foci are focal proliferations of phenotypically abnormal hepatocytes further sub-classified based on cytoplasmic tinctorial features, cell size, and morphology. Foci range from infra-lobular to supra-lobular in size and although sharply demarcated from surrounding parenchyma by their specific tinctorial affinity, they cause no or only minimal compression of adjacent hepatocytes (Fig. 5a). The most frequent type is the basophilic FCA which consists of cells smaller than normal hepatocytes with cytoplasmic basophilia, frequent intra-cytoplasmic glycogen accumulation, nuclear pleomorphism, nuclear hyperchromasia, and prominent nucleoli (Fig. 5a, b, and c) (see Note 56) [22]. Foci may show increased mitoses and are commonly found in close association with blood vessels (vascular pseudo-invasion) (Fig. 5c).

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Fig. 5 Histopathologic liver findings in the DEN-HFHS mouse model. (a) Focus of cellular alteration, basophilic type. The lesion consists of closely packed basophilic hepatocytes forming a cluster approximately the size of a liver lobule. (b) Focus of cellular alteration, basophilic type. Despite being sharply demarcated from adjacent liver tissue, this infra-lobular lesion causes no peripheral compression and merges imperceptibly with the surrounding parenchyma. Note diffuse mixed macro-mediovesicular steatosis in background liver. (c) Higher magnification of B. Altered hepatocytes demonstrate a decreased cell size and prominent cytoplasmic basophilia. Some mitotic figures and intra-cytoplasmic inclusion bodies can be seen (yellow arrowhead). The focus partially surrounds a central vein. (d) Hepatocellular adenoma. A round, nodular, well-demarcated, non-encapsulated, and expansive benign tumor showing prominent central fatty change and a densely cellular basophilic periphery (target appearance on ultrasound images). (e) Hepatocellular adenoma. Same morphology as in D. This benign neoplasm arises in a background of diffuse hepatic steatosis and shows less prominent central fatty change. Peripheral compression of adjacent liver is evident. (f) Hepatocellular adenoma at tumor margin. A benign tumor is partially visible on the left and shows loss of lobular architecture with an expansive growth, manifesting as a peripheral rim of pressure atrophy of non-neoplastic hepatocytes. Note the portal tract being distorted and progressively surrounded by tumor expansion. (g) Hepatocellular carcinoma. This large malignant tumor has almost replaced an entire hepatic lobe and shows diffuse loss of lobular architecture. A clear demarcation with non-tumoral liver is visible on the left. Both solid and trabecular (top right) growth patterns can be seen as well as a necrotic focus on the right (N). (h) Same HCC as in G, region with trabecular growth pattern. 5- to 10-cell-thick trabeculae of well-differentiated neoplastic hepatocytes alternates with sinusoid-like, blood-filled vascular spaces. A large necrotic focus is visible on

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(b) Hepatocellular adenomas (HCAs) are classically multiple, nodular, 1 mm to >1 cm in diameter, well demarcated, non-encapsulated, and expansive benign lesions that often bulge from the liver surface (Fig. 5d, e, and f). Necrosis is usually absent and the stroma is scant. Expansive growth manifests as a distinct compression of the adjacent liver parenchyma with a peri-tumoral rim of pressure atrophy of non-neoplastic hepatocytes and a parallel alignment of surrounding blood vessels (Fig. 5f). Loss of lobular architecture is a key diagnostic criterion and no portal tract should be visible within the tumor, however, adjacent portal triads may be entrapped at the margin by tumor growth (Fig. 5f) [34]. By definition, HCAs consist of well-differentiated neoplastic hepatocytes with minimal atypia (nuclear hyperchromasia, occasional karyomegaly) and varying cell size and tinctorial affinity, as described for foci of cellular alteration. In our experience, DEN-induced HFHS-promoted HCAs almost always show central fatty change (intracytoplasmic lipid accumulation) with a narrower peripheral zone of smaller, basophilic neoplastic hepatocytes similar in appearance to basophilic FCA (Fig. 5d) [35]. This specific histologic arrangement is responsible for the target appearance of those benign tumors on ultrasound images (Fig. 4f). Additionally, hepatocytes within DEN-induced adenomas and sometimes FCA invariably accumulate intra-cytoplasmic inclusion bodies, in the form of either several to multiple globular brightly acidophilic inclusions of varying size or a single pale pink inclusion that fills the cytoplasm and displaces the nucleus to the periphery [36] (Fig. 5c). (c) Hepatocellular carcinomas (HCCs) present generally as unique, large, nodular with sometimes irregular outlines, non-encapsulated and moderately to densely cellular malignant lesions which often replace entire lobes (Fig. 5g, h, and i). Necrosis and hemorrhages are frequently seen and may be so extensive as to involve up to 25–50% of the neoplasm (Fig. 5g, h). Tumor growth may appear expansive with multifocal infiltration or overtly invasive. Lobular architecture is lost and replaced by 3–4 to >10-cell-thick trabeculae of welldifferentiated hepatocytes (trabecular growth pattern) or ä Fig. 5 (continued) the left (N). (i) Same HCC as in G, region with solid growth pattern. A diffuse sheet of moderately differentiated neoplastic hepatocytes with cellular atypia (anisokaryosis, karyomegaly, nuclear pleomorphism, and nuclear hyperchromasia) and increased mitotic activity can be seen. Note glandular formation with the presence of a central clear space lined by neoplastic hepatocytes, suggesting pseudoglandular growth pattern in the tumor (yellow arrowhead). CV: Central Vein, N: Necrosis, PT: Portal Tract. Hematoxylin and Eosin staining

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diffuse sheets of moderately to poorly differentiated, pleomorphic neoplastic hepatocytes (solid growth pattern) (Fig. 5h, i) [34, 37]. HCCs frequently contain different histological patterns within different areas (Fig. 5g). The stroma is usually inconspicuous and leukocyte infiltration is variable. Cytological atypia includes anisocytosis, anisokaryosis, karyomegaly, nuclear pleomorphism, nuclear hyperchromasia, prominent nucleoli, and multinucleation (Fig. 5i). Intra-cytoplasmic inclusion bodies as described for HCAs and FCA may be found with an apparently lower frequency. Increased mitotic index, aberrant mitotic figures, fatty or clear cell change, and foci of extramedullary hematopoiesis are common findings. HCCs may sometimes be seen arising in pre-existing hepatocellular adenomas [31]. 3.5 Tumor Development Modifiers

Many intrinsic and extrinsic factors have been shown to modify the course of liver tumorigenesis after DEN injection, either affecting tumor multiplicity, the latency period, or progression to malignancy [20]. Intrinsic factors include sex (females are more resistant), age at DEN administration (younger mice are more prone to tumorigenesis), and mouse strain (C57BL/6J mice are more resistant) [23, 38]. Accordingly, a dose of DEN as high as 80 mg/kg is necessary to achieve a 90% prevalence rate of liver neoplasms after 34 weeks in mice injected at 16 weeks of age and put under HFD at 6 weeks of age [21]. Levels of CYP2E1 expression or activity are also critical for the initial DEN bioactivation and mice defective for this cytochrome isoform are greatly protected from tumorigenesis [39]. All those modifiers mainly influence tumor initiation or early promotion and finally tumor multiplicity. Xenobiotics or genetic manipulations that modify the susceptibility of hepatocytes to succumb to DEN bioactivation, the response of the innate immune system to hepatocyte injury and death, or the ensuing compensatory hepatocellular hyperplasia are also known to modulate the extent of early liver damage and tumorigenesis [23, 40, 41]. Extrinsic factors encompass the route of administration (parenteral route or per os) and quantity of DEN administered to mice, DEN-induced tumorigenesis being a dose-dependent process [42]. Promoters favor tumor development by various mechanisms such as enzymatic induction (e.g., phenobarbital) or release of tumor-promoting cytokines and increased proliferation (e.g., CCl4, hypercaloric diets) [20]. Various hypercaloric diets have been described in conjunction with DEN exposure, such as highfat diet (HFD) and choline-deficient high-fat diet (CD-HFD), and may modulate background liver lesions (e.g., fibrosis, inflammation) and/or tumor latency [15]. It is interesting to consider that promoters may influence at least to some extent the genetic landscape of DEN-induced hepatocellular neoplasms, probably by supporting the emergence of tumors with specific mutations. Indeed,

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in C57BL/6J male mice, a single dose of DEN with or without hypercaloric feeding results in tumors which predominantly carry an activating mutation in Braf (the main oncogenic driver in C3H/HeOuJ males is Hras) whereas phenobarbital-promoted neoplasms show more frequent Ctnnb1 (β-catenin) activating mutations [17, 18]. Intestinal and biliary microbiota have emerged as leading factors accelerating or halting tumor development depending on their composition. Notably, various Helicobacter species (e.g., Helicobacter hepaticus, H. bilis) are well known to induce chronic liver injury in mice and create a microenvironment permissive to malignant transformation or progression [43]. Extrinsic modifiers may thus influence each step of tumor development from initiation to promotion and progression.

4

Notes 1. DEN susceptibility varies according to age, sex, and mouse strains. Older mice can be used as well but the dose of DEN needs to be increased accordingly (doses between 50–90 mg/ kg body weight for animals between 10–16 weeks of age). 2. To obtain a sufficient number of young mice for the experiment, we recommend, when possible, that synchronized pregnant C57BL/6J female mice be ordered from your local mouse supplier company. In our experience, we generally obtain an average of 7 mouse pups per litter. The pregnant females should be shipped at day 15 post-coitum and housed in a calm environment with enough materials for nest building. Several days after parturition, pups can be sexed, and homogeneous groups of the adequate size can be obtained by fostering. 3. DEN is a volatile, clear yellow, oily liquid solution at room temperature that should be stored protected from light in a well-ventilated place at 4 °C. It is advisable to keep it locked up with an access restricted to authorized persons. 4. DEN is classified as toxic if swallowed (H301), may cause cancer (H350), and is harmful to aquatic life with long-lasting effects (H412). It should always be handled with appropriate personal protective equipment (nitrile gloves, goggles, lab coat). 5. The fume hood should expulse the filtrated air outside the laboratory. 6. Please refer to your institution’s procedure for the proper disposal of carcinogenic, mutagenic, and reprotoxic (CMR) substances.

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7. To insure sterilityirradiation of feed pellets is recommended (25 kGy per 2 kg bag of diet). HFHS pellets should be stored in a clean and dry place, at 4 °C, protected from light. 8. Transducers with a lower central frequency (180 mg/dL) from group 2 and the animals are fed with high-fat diet (HFD). As a control, group 1 is fed with regular chow diet (RCD). All mice are fed for 20 weeks. From weeks 21 to 24, tumor growth is monitored by weekly echography. All animals are sacrificed 24 weeks after the beginning of the experiment

euthanized after 20 weeks of HFD, and livers are collected and photographed to confirm the presence of macroscopic tumors (Fig. 2b). To validate HCC diagnosis, histological analysis from the livers can also be performed by hematoxylin eosin saffron (HES) staining. Additionally, the HCC can be characterized by immunohistochemistry, for instance to measure the levels of proliferation marker Ki67 and the expression of the HCC marker alpha fetoprotein (AFP) [13, 14]. Hepatic damage can also be indirectly assessed by measuring plasma levels of alanine and aspartate transaminases activity (ALT and AST, respectively) [15].

2 Materials 2.1

Reagents

1. 5% Glucose. 2. Glucose test strips. 3. High-fat diet (U8978 Version 19, Safe diets). 4. 1 M Hydrochloric acid (HCl). 5. Isoflurane (anesthetic gas). 6. Neonatal C57Bl/6J male mice. 7. Ophthalmic liquid gel. 8. Oxygen tank.

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Fig. 2 Representative pictures of the liver with HCC derived by diabetes/NAFLD. (a) The development of the hepatic tumor is analyzed by echography in mice (treated with vehicle plus RCD) and in mice from the experimental group (treated with STZ plus HFD) before sacrifice. (b) Representative photographs of the livers from the control and experimental groups are shown. The tumor area is labeled by a red circle

9. Regular chow diet (A04, Safe diets). 10. 50 mM sodium citrate buffer, pH 4.5. 11. Electric trimmer and depilatory cream. 12. Streptozotocin (S0130, Sigma Aldrich). 13. 10% Sucrose. 2.2

Equipment

1. 1 ml syringes. 2. 2 ml microcentrifuge tubes. 3. 100 μm Filter. 4. 27-G needles. 5. Blood glucometer. 6. Aluminum foil. 7. Echography in vivo imagen system (Vevo® 3100 from FUJIFILM VisualSonics). 8. Cotton buds.

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9. Heparin tube (16443, Sarstedt). 10. Isoflurane vaporizer system. 11. Scale (for measuring mouse weight). 12. Medical tapes. 13. Gauzes.

3

Methods

3.1 Prepare Animals: Time 4–6 h

At least 4–6 h prior to initiating the experiment, house 2–5 neonatal (2 days old) male mice with mother female mice per cage at 24 °C ± 1 °C and 55% ± 5% (see Note 1).

3.2 Fresh Streptozotocin Buffer Preparation: Time 30 min

1. Prepare 50 mL (final volume) of a 50 mM sodium citrate buffer, pH 4.5. (a) Dissolve 535 mg in 40 mL of deionized water. (b) Set the pH to 4.5 by slowly adding 1 M HCl (see Note 2). (c) Once pH = 4.5, complete the volume to 50 mL. (d) Filter the solution. 2. Weigh 4 mg of streptozotocin into a 2 ml microcentrifuge tube and cover the tubes with aluminum foil (see Note 3).

3.3 Diabetes Mellitus Induced by Streptozotocin: Time 4 Weeks

1. Immediately prior to injection, dissolve the streptozotocin in 50 mM sodium citrate buffer (pH 4.5) to a final concentration of 2 mg/ml (see Notes 3 and 4). 2. Perform intraperitoneal injection (i.p.; lower side of the abdomen with a 27-gauge needle) of 200 μg contained in 100 μL of streptozotocin buffer into C57BL/6J mice at 2 days of age. Inject an equal volume of citrate buffer (pH 4.5) i.p. into the control group mice (see Note 5). 3. Return the mice to their home cages. Provide free access to normal food and 10% sucrose water. 4. Inspect animals every 2 h after injection for 8 h and every day for at least 1 week (see Note 6). 5. After one week, place the animals in a new cage without food to start fasting (fresh bedding is crucial to avoid the risk that food leftovers or feces being eaten during the fasting period). 6. After 4–6 h fasting, measure glycemia from one drop of blood from the tail vein by a blood glucometer. after this, allow food and water ad libitum (see Note 7). 7. Check mice weight and measure the glycemia (see Subheading 3.3, Step 6) every week for 3 weeks. 8. Euthanize mice with levels of glycemia less than 180–200 mg/ dL after 3 weeks.

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3.4 NAS, NASH, Fibrosis, and HCC Accelerated by HFD: Time 20 Weeks

1. Remove the control diet (regular chow diet) and feed mice with high fat diet (HFD). Allow the food and water ad libitum (see Note 8). 2. Follow the weight weekly and glycemia monthly (as described in Subheading 3.3, Step 6). 3. After 16 weeks, anesthetize mice with 2% isoflurane and follow the developing of the tumors by echography (see Note 9): (a) Turn on the Vevo® 3100 imaging system and set up the heated platform to 38 °C. (b) Anesthetize the animal with 2% isoflurane and 1.5–2 L/ min oxygen flux in the anesthesia chamber. (c) Transfer the mouse to the anesthesia table and fix it with adhesive plaster, maintaining the vaporization of isoflurane/oxygen mix. (d) Eliminate the abdomen hair with electric trimmer and then with depilatory cream and use wet gauzes for cleaning the observational area. (e) Put transmission gel in the abdomen region. (f) Use the B-mode of MX550D transducer from the imaging system in transversal position and take adequate images of the liver (see Note 10). (g) Return the animal to the cage and re-inspect animals after they wake up. 4. Repeat Subheading 3.4, Step 3 weekly to follow up the tumor growth up to 20 weeks. 5. One day before week 20 with HFD, remove the food and keep in fasting for 16 h (see Note 11). 6. Measure the glycemia (see Subheading 3.3, Step 6), collect blood from the submandibular vain in a heparin tube, and keep it at 4 °C. 7. Euthanize all animals by cervical dislocation. 8. Collect and take images of the liver.

4

Notes 1. Most of streptozotocin-induced diabetic mouse studies are carried on male animals because female mice are less sensitive to islet-cell destruction [16]. Moreover, we recommend creating group sizes of 12 to 20 due to the morbidity associated with the streptozotocin treatment, although the protocol detailed is designed to minimize variability. The final number of mice should be equal for each group.

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2. Be careful to let the solution homogenize and the pH stabilize between each drop of HCl. 3. IMPORTANT: Use one tube for three mice because streptozotocin degrades within 15 to 20 min after dissolving in the citrate buffer. 4. Create one control group of mice with regular chow diet and injection of control citrate buffer (vehicle), and one control group with HFD without the initial streptozotocin injection (vehicle). 5. The streptozotocin solution should be prepared immediately before use and injected within 5 min of dissolution. 6. Although the streptozotocin dose is low, some mice will die soon after receiving streptozotocin because of the release of insulin from the pancreas (rapid and massive β-cell necrosis), causing fatal hypoglycemia during the first 24 h [17]. Check the levels of glucose in plasma to prevent fatal hypoglycemia, if glucose levels are