Genome Editing in Biomedical Sciences (Advances in Experimental Medicine and Biology, 1429) [1st ed. 2023] 3031333241, 9783031333248

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Advances in Experimental Medicine and Biology  1429

Geraldo A. Passos   Editor

Genome Editing in Biomedical Sciences

Advances in Experimental Medicine and Biology Series Editors Wim E. Crusio, Institut de Neurosciences Cognitives et Intégratives d’Aquitaine, CNRS and University of Bordeaux, Pessac Cedex, France Haidong Dong, Departments of Urology and Immunology, Mayo Clinic, Rochester, MN, USA Heinfried H. Radeke, Institute of Pharmacology and Toxicology, Clinic of the Goethe University Frankfurt Main, Frankfurt am Main, Hessen, Germany Nima Rezaei , Research Center for Immunodeficiencies, Children’s Medical Center, Tehran University of Medical Sciences, Tehran, Iran Ortrud Steinlein, Institute of Human Genetics, LMU University Hospital, Munich, Germany Junjie Xiao, Cardiac Regeneration and Ageing Lab, Institute of Cardiovascular Sciences, School of Life Science, Shanghai University, Shanghai, China

Advances in Experimental Medicine and Biology provides a platform for scientific contributions in the main disciplines of the biomedicine and the life sciences. This series publishes thematic volumes on contemporary research in the areas of microbiology, immunology, neurosciences, biochemistry, biomedical engineering, genetics, physiology, and cancer research. Covering emerging topics and techniques in basic and clinical science, it brings together clinicians and researchers from various fields. Advances in Experimental Medicine and Biology has been publishing exceptional works in the field for over 40 years, and is indexed in SCOPUS, Medline (PubMed), EMBASE, BIOSIS, Reaxys, EMBiology, the Chemical Abstracts Service (CAS), and Pathway Studio. 2021 Impact Factor: 3.650 (no longer indexed in SCIE as of 2022)

Geraldo A. Passos Editor

Genome Editing in Biomedical Sciences

Editor Geraldo A. Passos Laboratory of Genetics and Molecular Biology, Department of Basic and Oral Biology, School of Dentistry of Ribeirão Preto University of São Paulo Ribeirão Preto Campus, SP, Brazil Molecular Immunogenetics Group Department of Genetics Ribeirão Preto Medical School University of São Paulo Ribeirão Preto Campus, SP, Brazil

ISSN 0065-2598 ISSN 2214-8019 (electronic) Advances in Experimental Medicine and Biology ISBN 978-3-031-33324-8 ISBN 978-3-031-33325-5 (eBook) https://doi.org/10.1007/978-3-031-33325-5 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 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 Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

The science of genetics is only possible because organisms exhibit both phenotypic and DNA sequence variability. Variations in the nucleotide sequence involving a single nucleotide, both in the protein-coding regions of a gene (exons) and in the non-coding regions (introns), occur spontaneously and refer to single-nucleotide polymorphisms (SNPs), which may or may not influence the phenotype. Mutations that significantly affect the phenotype, whether due to variations of a single nucleotide or more, are rarer in the population. When they occur, they are usually deleterious or cause genetic diseases in the human species. However, we need mutations to be able to do experimental genetics. For this reason, researchers have been looking for methods to induce mutations since the beginning of the twentieth century, when Herman Müller’s seminal experiments demonstrated that when the fly Drosophila melanogaster was exposed to high doses of X-rays, there was an increase in genetic mutations (www.nobelprize.org/prizes/ medicine/1946/muller/facts). This was the first demonstration of the possibility of inducing mutations with a physical agent (X-rays), and other experimental examples followed, by other researchers, demonstrating that irradiating bacteria with ultraviolet light caused an increase in mutations. Mustard gas was another example, this time of a chemical agent, of inducing experimental mutations. These discoveries advanced genetics enormously, as an immense collection of mutant experimental organisms (such as Drosophila, for example) could be identified and isolated. However, the use of physical or chemical mutagenic agents has the disadvantage of not being able to choose “a priori” the target gene in which we want to induce mutations, as this is a non-site-specific process generating random mutations. With these agents, we need to analyze hundreds or even thousands of treated individuals until we identify the one that harbors a particular mutation in a gene of interest. Moreover, this becomes complicated when the organism studied is an arthropod like Drosophila or a mammal like the Mus musculus mouse.

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For this reason, geneticists began to develop other strategies to produce sitedirected mutagenesis, such as using the nucleotide analog N4-hydroxycytidine to provoke the transition from GC to AT in the bacteriophage Qbeta (Flavell et al. 1975). Geneticists have achieved a significant improvement, such as the “scanning mutagenesis of oligo-directed targets (SMOOT)” technique (Cerchione et al. 2020), with which it is possible to generate a plasmid library with inserts harboring single-nucleotide mutations across an entire gene. However, this method proved limited and needed to be more specific. Several examples of attempts to induce sitedirected mutations followed, but all make use of bacteriophages, plasmids, bacteria, or yeast. Thus, it is possible to transfect cells with these plasmids and evaluate the effect of each point mutation. This protocol is very accurate but does not represent actual genome editing. Genetics needed a protocol to edit genes directly in living organisms accurately, that is, choose a target gene, select an exon in this gene, and cause mutation in this exact location of the genome in a living cell or intact individual. The emergence of knockout mice (KO) by homologous recombination, which significantly boosted biomedical research, could serve as an example to meet this need. However, not all experimental animals other than the mouse have been genetically knocked out by this technique. In addition, KO animals usually present interruption of the gene sequence from exon 2 or 3, generating a truncated protein. The generation of KO mice was close, but this still did not represent gene editing. It was when a little over 10 years ago, in 2012, that the first scientific papers on the use of the Crispr technique began to be published, the acronym for “clustered regularly interspaced short palindromic repeats” by Jennifer A. Doudna and Emmanuelle Charpentier and collaborators (Jinek et al. 2012). The authors showed that the crRNA, together with the tracRNA, form a structure that “guides” the Cas9 enzyme to a “target” region of the genome and there causes double-strand breaks. Therefore, they managed to direct a molecular complex capable of producing mutations in a specific genome region. These researchers did not discover the Crispr system. Still, it was previously described independently by the groups of Y. Ishino and M. J. Mojica when they analyzed the genome of the bacterium E. coli (Ishino et al. 2018). The Crispr system is ancient in evolution and is part of the defense system of bacteria against bacteriophages (Makarova et al. 2020). The brilliance was to use this system to cause mutations in other organisms. Applying the Crispr system earned Doudna and Charpentier the Nobel Prize in 2020 “for the development of a method for genome editing” (www.nobelprize.org/prizes/ chemistry/2020/summary). Currently, the Crispr technique, or “Crispr toolbox,” is widely used both in basic research for the identification of the function of eukaryotic or prokaryotic genes, in medicine as a means of gene therapy, in the pharmaceutical industry for the production of new therapeutic drugs, and in agriculture and veterinary in the control of pests or the generation of plants and animals resistant to diseases (Passos et al. 2016).

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If I had to select a few examples of Crispr use that have most impacted biomedical research and medicine, the focus of this book would first say the elimination of the HIV-1 virus genome from infected T cells (Kaminski et al. 2016) or the correction of the genetic disease hypertrophic cardiomyopathy (Ma et al. 2017) or the reprogramming of human T lymphocytes ex vivo for their use in the treatment of cancer in vivo (Zhou et al. 2022; Tomasik et al. 2022). This book has been organized to show examples of using the Crispr system in biomedicine. It includes 11 chapters and covers the fundamental concepts of Crispr to the ethical controversy generated by the possibility of gene-edited babies. This book will be helpful to specialized researchers and to students who aim to introduce themselves to modern genome editing research. I thank all the authors who contributed to their chapters and Springer Nature for this opportunity and continued support. Ribeirão Preto, São Paulo, Brazil

Geraldo A. Passos

References Cerchione D, Loveluck K, Tillotson EL, Harbinski F, DaSilva J, Kelley CP, Keston-Smith E, Fernandez CA, Myer VE, Jayaram H, Steinberg BE (2020) SMOOT libraries and phageinduced directed evolution of Cas9 to engineer reduced off-target activity. PLoS One 15(4): e0231716. https://doi.org/10.1371/journal.pone.0231716 Flavell RA, Sabo DL, Bandle EF, Weissmann C (1975) Site-directed mutagenesis: effect of an extracistronic mutation on the in vitro propagation of bacteriophage Qbeta RNA. Proc Natl Acad Sci U S A 72(1):367–371. https://doi.org/10.1073/pnas.72.1.367 Ishino Y, Krupovic M, Forterre P (2018) History of CRISPR-Cas from encounter with a mysterious repeated sequence to genome editing technology. J Bacteriol 200(7):e00580-17. https://doi.org/ 10.1128/JB.00580-17 Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337(6096):816– 821. https://doi.org/10.1126/science.1225829 Kaminski R, Chen Y, Fischer T, Tedaldi E, Napoli A, Zhang Y, Karn J, Hu W, Khalili K (2016) Elimination of HIV-1 genomes from human T-lymphoid cells by CRISPR/Cas9 gene editing. Sci Rep 6:22555. https://doi.org/10.1038/srep22555. Erratum in: Sci Rep 2016 July 07;6:28213 Ma H, Marti-Gutierrez N, Park SW, Wu J, Lee Y, Suzuki K, Koski A, Ji D, Hayama T, Ahmed R, Darby H, Van Dyken C, Li Y, Kang E, Park AR, Kim D, Kim ST, Gong J, Gu Y, Xu X, Battaglia D, Krieg SA, Lee DM, Wu DH, Wolf DP, Heitner SB, Belmonte JCI, Amato P, Kim JS, Kaul S, Mitalipov S (2017) Correction of a pathogenic gene mutation in human embryos. Nature 548(7668):413–419. https://doi.org/10.1038/nature23305 Makarova KS, Wolf YI, Iranzo J, Shmakov SA, Alkhnbashi OS, Brouns SJJ, Charpentier E, Cheng D, Haft DH, Horvath P, Moineau S, Mojica FJM, Scott D, Shah SA, Siksnys V, Terns MP, Venclovas Č, White MF, Yakunin AF, Yan W, Zhang F, Garrett RA, Backofen R, van der Oost J, Barrangou R, Koonin EV (2020) Evolutionary classification of CRISPR-Cas systems: a burst of class 2 and derived variants. Nat Rev Microbiol 18(2):67–83. https://doi.org/10.1038/ s41579-019-0299-x

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Passos GA, Speck-Hernandez CA, Sousa LC, Felicio RF, Souza TA (2016) Aplicações da técnica. In: Introdução à técnica de Crispr (T. C. Pereira, Editor). Sociedade Brasileira de Genética. ISBN 978-85-89265-24-9 Tomasik J, Jasiński M, Basak GW (2022) Next generations of CAR-T cells – new therapeutic opportunities in hematology? Front Immunol 13:1034707. https://doi.org/10.3389/fimmu.2022. 1034707 Zhou Z, Tao C, Li J, Tang JC, Chan AS, Zhou Y (2022) Chimeric antigen receptor T cells applied to solid tumors. Front Immunol 13:984864. https://doi.org/10.3389/fimmu.2022.984864

Contents

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What Is the CRISPR System and How It Is Used? . . . . . . . . . . . . . Danyel F. Contiliani, Vitor N. Moraes, Geraldo A. Passos, and Tiago Campos Pereira

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Modern Approaches to Mouse Genome Editing Using the CRISPR-Cas Toolbox and Their Applications in Functional Genomics and Translational Research . . . . . . . . . . . . Cintia J. Monteiro, David M. Heery, and Jonathan B. Whitchurch

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Long Non-coding RNAs and CRISPR-Cas Edition in Tumorigenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cristiana Libardi Miranda Furtado, Renan da Silva Santos, Sarah Leyenne Alves Sales, Louhana Pinheiro Rodrigues Teixeira, and Claudia do Ó. Pessoa

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The CRISPR/Cas System in Human Cancer . . . . . . . . . . . . . . . . . . Martín Hérnan Bonamino and Eduardo Mannarino Correia

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Using CRISPR/Cas9 to Edit a Thyroid Cancer Cell Line . . . . . . . . Cesar Seigi Fuziwara and Edna Teruko Kimura

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Genome Editing for Engineering the Next Generation of Advanced Immune Cell Therapies . . . . . . . . . . . . . . . . . . . . . . . Sarah Caroline Gomes de Lima, Daianne Maciely Carvalho Fantacini, Izadora Peter Furtado, Rafaela Rossetti, Roberta Maraninchi Silveira, Dimas Tadeu Covas, and Lucas Eduardo Botelho de Souza

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CRISPR Genome Editing and the Study of Chagas Disease . . . . . . 111 Gabriela de Assis Burle-Caldas, Viviane Grazielle-Silva, Lídia Paula Faustino, and Santuza Maria Ribeiro Teixeira

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Genome Editing Tools for Lysosomal Storage Disorders . . . . . . . . . 127 Esteban Alberto Gonzalez, Helena Nader, Marina Siebert, Diego A. Suarez, Carlos J. Alméciga-Díaz, and Guilherme Baldo

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CRISPR Libraries and Whole-Genome Screening to Identify Essential Factors for Viral Infections . . . . . . . . . . . . . . . . . . . . . . . 157 Isadora Marques Paiva, Samara Damasceno, and Thiago Mattar Cunha

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Gene Editing Technologies Targeting TFAM and Its Relation to Mitochondrial Diseases . . . . . . . . . . . . . . . . . . . 173 Vanessa Cristina de Oliveira, Kelly Cristine Santos Roballo, Clesio Gomes Mariano Junior, and Carlos Eduardo Ambrósio

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The Gene-Edited Babies Controversy: Reactions in the Scientific Community, Social Media, and the Press . . . . . . . . 191 Morgan Meyer and Frédéric Vergnaud

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205

About the Editor

Geraldo A. Passos received his Ph.D. degree in Biochemistry in 1988 from Ribeirão Preto Medical School, University of São Paulo (USP), Brazil. His postdoctoral studies were conducted at the Molecular Genetics Institute of Montpellier (Centre National de la Recherche Scientifique, CNRS), France (1992–1994), where he worked with sequencing and physical mapping of the human immunoglobulin lambda locus on chromosome 22q11.2. For several years, he worked in close collaboration with the transcriptome and microarray team of the TAGC Lab (Technologies Avancées pour le Génome et la Clinique) at the Centre d’Immunologie de Marseille-Luminy in Marseille, France (1999–2001) and then at the Institut National de la Santé et de la Recherche Médicale (INSERM) Unité 1090 at the Luminy Scientific Park in Marseille, France (2002–2017) to study the large scale gene expression profiling of the murine thymus gland. He is currently an Associate Professor of Genetics and Molecular Biology at the School of Dentistry of Ribeirão Preto, USP, and a collaborator at the Ribeirão Preto Medical School, USP, where he is the Head of the Molecular Immunogenetics Group.

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What Is the CRISPR System and How It Is Used? Danyel F. Contiliani, Vitor N. Moraes, Geraldo A. Passos, and Tiago Campos Pereira

1 History The clustered regularly interspaced short palindromic repeats (CRISPR) system is currently a cutting-edge technology used in many facets of the life sciences. Before becoming a powerful biotechnology tool, the CRISPR system was initially a conundrum found in prokaryotic genomes. Its first report came in 1987 when Yoshimuzi Ishino and colleagues were analyzing the bacterial genome of Escherichia coli and encountered an enigmatic locus composed of short repeated palindromic nucleotide sequences interspersed with spacer sequences (Forterre et al. 2018) (Fig. 1.1). At the dawn of the genomic era, these mysterious repeat sequences were found in archaea (Ishino et al. 1987; Mojica et al. 1993) and other bacterial species (Mojica et al. 2000). Thus, the conservation of those unknown genetic elements, especially in

D. F. Contiliani Centro de Cana, Instituto Agronômico de Campinas, Ribeirão Preto, SP, Brazil Graduate Program in Genetics, Ribeirão Preto Medical School, University of São Paulo (USP), Ribeirão Preto, SP, Brazil V. N. Moraes · T. C. Pereira (✉) Graduate Program in Genetics, Ribeirão Preto Medical School, University of São Paulo (USP), Ribeirão Preto, SP, Brazil Department of Biology, Faculty of Philosophy, Sciences and Letters of Ribeirão Preto, USP, Ribeirão Preto, SP, Brazil e-mail: [email protected] G. A. Passos Laboratory of Genetics and Molecular Biology, Department of Basic and Oral Biology, School of Dentistry of Ribeirão Preto, University of São Paulo, Ribeirão Preto Campus, SP, Brazil Molecular Immunogenetics Group, Department of Genetics, Ribeirão Preto Medical School, University of São Paulo, Ribeirão Preto Campus, SP, Brazil © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_1

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Fig. 1.1 Process of CRISPR-Cas defense system. Fragments of the viral genetic material are incorporated into the bacterial CRISPR locus as spacers (adaptation). These stored foreign sequences are subsequently transcribed and processed into mature crRNAs (CRISPR expression), which form ribonucleoprotein complexes with the Cas9 enzyme and guide it towards cleavage of the invading DNA (interference)

prokaryotes, started to draw the attention of the scientific community and was distinctively named short regularly spaced repeats (SRSRs), spacers interspersed direct repeats (SPIDRs), and large cluster of tandem repeats (LCTRs). Ultimately, the acronym CRISPR was coined (Jansen et al. 2002) and became widely accepted by the community. Leveraging the plethora of CRISPR genomic region sequences retrieved from numerous microorganisms, Jansen and his colleagues were able to identify conserved genes physically close to CRISPR loci, which were referred to as CRISPRassociated genes (Cas genes) (Jansen et al. 2002). These genes are responsible for encoding DNA-binding enzymes, such as nucleases, polymerases, and helicases, whose structures were later resolved. However, the molecular mechanisms underlying the CRISPR-Cas system involving these proteins remained a mystery. Three years later, two independent research groups highlighted the strong similarities between the spacer sequences and bacteriophage and plasmid sequences (Pourcel et al. 2005; Mojica et al. 2005). This evidence prompted the hypothesis that the CRISPR-Cas system might be a prokaryotic adaptive defense mechanism against viral infections. Further investigations confirmed that these spacer sequences are derived from foreign genetic elements, and only the bacteria that possessed them

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were resistant to subsequent phage infections (Bolotin et al. 2005; Barrangou et al. 2007). Based on these findings, the initial understanding was that the spacer-decoded RNA molecules would be complementary to foreign mobile genetic elements, resembling the eukaryotic sequence-specific mechanism of RNA interference (RNAi). Finally, in 2007, Barrangou and colleagues demonstrated how CRISPR spacers could drive the specificity of this heritable, nucleic acid-based, prokaryotic immune system (Barrangou et al. 2007).

2 Conceptual Aspects Within the vast diversity of microorganisms, CRISPR and its molecular components vary remarkably (e.g., spacer size and composition, number of spacer-repeat units, cas genes composition). Still, the entire defense machinery acts in a similar way against foreign mobile genetic elements, such as transposons, plasmids, and bacteriophages (Weiss and Sampson 2014). Thus, the CRISPR-Cas defense system (Fig. 1.1) can be divided into three steps: (i) adaptation, (ii) CRISPR expression, and (iii) interference. The first step – adaptation – is defined by the acquisition of bacterial immunity to the infectious pathogen. Upon the entry of viral genetic material into the prokaryotic cell, the most conserved Cas genes, Cas1 and Cas2, are transcriptionally induced, thereby prompting the formation of the Cas1-Cas2 enzymatic complex. These macromolecules are capable of cleaving the viral DNA into small fragments (28–48 nucleotides) known as spacers, which are subsequently integrated into the bacterial genome, thus extending the repetitive elements in the CRISPR array (Makarova et al. 2011). Afterwards – in the CRISPR expression phase –, the CRISPR locus is relentlessly transcribed from an AT-rich leader sequence, yielding the CRISPR-derived RNA (crRNA). In general, a long primary precursor crRNA (pre-crRNA) is first generated and processed into a set of short crRNAs, each one of which corresponds to a distinct spacer (Makarova et al. 2011). This crRNA processing (or RNA maturation step) may be performed by different ways (e.g., via specific Cas endonucleases or RNase III) (Jiang and Marraffini 2015), thus resulting in diverse “types” of CRISPR system (types I, II, III, etc). Particularly, the type-II system requires the annealing between the crRNA and a small non-coding RNA, called trans-activating crRNA (tracrRNA) (Jinek et al. 2012). Finally, in the interference step, the mature crRNA docks to Cas proteins to assemble ribonucleoprotein (RNP) complexes. Once the bacteriophage returns to infect the host bacterium, the crRNA serves as a molecule that guides the Cas endonuclease to recognize the viral nucleotide sequence (Brouns et al. 2008), known as protospacer (complementary to the bacterial spacer in the CRISPR array). Ultimately, the Cas enzyme sharply performs the double-strand break (DSB) of the viral DNA.

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In summary, the CRISPR-Cas system naturally consists of a prokaryotic acquired-immunity mechanism, in which Cas proteins act as effectors, crRNAs serve as carrier molecules, and the CRISPR locus stands as genetic memory. Analogously to the RNA interference (RNAi) system from eukaryotes, CRISPRCas acts to cleave target sequences by its protein complex anchored to a small non-coding RNA (Bhaya et al. 2011). Based on these similarities and leveraging key molecular aspects (i.e., ability to generate DSBs) of the CRISPR-Cas system, the development of a novel genome editing technology has become feasible.

3 Molecular Mechanisms of the CRISPR-Cas System Genome editing tools such as meganucleases, zinc finger nucleases (ZFNs), and transcription activator-like effector nucleases (TALENs) were reconstructed from observable natural phenomena to suit biotechnology and our understanding of the relationships between genotypes and phenotypes (e.g., genetic diseases). In particular, ZFNs and TALENs have promoted a more flexible manipulation of DNA with their programmable DNA-binding proteins. Nevertheless, these strategies are laborious and pricey. In 2012, the groundbreaking article on Science (Jinek et al. 2012) revealed how the prokaryotic CRISPR-Cas immunity system could be converted into a simple, effective, and versatile genome engineering tool. Thus, the traditional CRISPR-Cas genome editing technique comprises two main components: an endonuclease (usually a Cas9) and a single-guide RNA (sgRNA). Based on the simplest type of CRISPR-Cas system – the type II – in which the Cas9 endonuclease is responsible for cleaving the foreign genetic element, the most commonly used Cas9 enzyme in genome editing was isolated from the bacterium Streptococcus pyogenes (SpCas9). Its tridimensional structure features two catalytic domains, HNH and RuvC, which independently cut the complementary and non-complementary DNA strands, respectively, causing a DSB (Chen et al. 2014). Moreover, the Cas9 structure also has a positively charged groove (PAM-interacting domain) that nests the protospacer adjacent motif (PAM) of the target (Nishimasu et al. 2014). This is particularly important since PAM recognition is a critical aspect of DNA strand separation and the formation of the sgRNA:DNA heteroduplex (Sternberg et al. 2014). Thus, the SpCas9 enzyme especially recognizes the 5’-NGG-3’ PAM motif (N means any nucleotide) and performs the DSB approximately 3 base pairs upstream of this site, resulting in blunt ends (Jinek et al. 2012). Interestingly, a diversity of PAM sequences can be functional depending on the system type, or on engineered variants of Cas9 (Anders et al. 2016; Leenay et al. 2016; Gleditzsch et al. 2019), thus not restricting the CRISPR-Cas machinery to a single motif. However, Cas9-mediated cleavage is reliant on activation and guidance by an RNA molecule. As previously illustrated, the type-II CRISPR system entails the pairing of two non-coding RNAs, crRNA and tracrRNA, which form an active and stable structure capable of guiding Cas9 to the target genetic material (Deltcheva

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Fig. 1.2 CRISPR-Cas9 mechanism of action. The illustrated ribonucleoprotein (RNP) complex is bound to the target DNA and opens both of its strands. The RNP is composed of the Cas9 endonuclease (orange) and the single-guide RNA (gRNA), which is formed by two parts: (i) the guide sequence (brown; including the seed region, in blue) and (ii) the scaffold (black). The complementarity between the guide sequence and the target DNA sequence (red) upstream of the PAM site enables the activation of Cas9, whose catalytic domains, RuvC and HNH (scissors), act cooperatively to perform the double-strand break (DSB)

et al. 2011). To simplify this process for experimental procedures, Jinek and colleagues engineered a single chimeric RNA molecule (Fig. 1.2) – the singleguide RNA (sgRNA) – whose structural aspects mimic the crRNA:tracrRNA complex (Jinek et al. 2012). The sgRNA is composed of a secondary hairpin structure formed by the guide sequence and the 3′ end-scaffold (~80 nt). The guide sequence is highlighted because of its 20 nucleotides complementary to the target sequence (upstream to the PAM site) of a particular gene of interest (Anders et al. 2016). More importantly, the first 10 nucleotides (3′ end) of the guide sequence correspond to the seed region, where the pairing between sgRNA and the target gene is initiated (Jiang and Doudna 2017). Therefore, the seed sequence must be fully complementary to the target; otherwise, mismatches in this region cause instabilities in the interaction of the RNP complex and the target DNA (Semenova et al. 2011; Jinek et al. 2012; Cong et al. 2013). After cleavage of the target sequence, the cell can trigger two main DNA repair pathways: the non-homologous end joining (NHEJ) and the homology-directed repair (HDR) (Fig. 1.3). Briefly, the most prevalent DSB repair mechanism in mammalian cells is mediated by the NHEJ pathway, which can occur promptly throughout the cell cycle (Mao et al. 2008). This process ligates the cleaved ends of the target DNA with minor end processing. However, as an error-prone mechanism, some nucleotides can be deleted or inserted (indels) in the cleavage site, thereby

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Fig. 1.3 Repair mechanisms of CRISPR-induced double-strand breaks. Upon cleavage of the target DNA mediated by the Cas9 endonuclease upstream to the PAM site (blue), the cell may trigger two potential DNA repair mechanisms: non-homologous end joining (NHEJ) and homology-directed repair (HDR). The error-proneness of the NHEJ mechanism can result in small insertions or deletions (indels) at the cut site, leading to a frameshift in the coding region of the target gene. Besides, the HDR mechanism relies on a template donor DNA featuring the desired mutation (purple) and the modified PAM site (green) flanked by homology arms (highlighted in gray) to the target DNA. After recombination, the target DNA is accurately edited

resulting in a frameshift, which often impairs the functionality of the translated protein (Hsu et al. 2014). Therefore, NHEJ is suitable for gene knockout strategies. Conversely, the error-free HDR mechanism conducts DSB repair by relying on an exogenous DNA donor molecule (Gallagher and Haber 2018). The long-range resection at the chromosome break results in 3′ single-strand DNA (ssDNA) tails that allow the invasion of a strand of the cleaved DNA into the homologous region of the donor DNA. Once the target DNA is resynthesized based on the donor DNA, the region flanked by homology arms (which carries the desired modification) is inserted into the cut site (Nambiar et al. 2022). Thus, this mechanism provides a precise repair that allows not merely allelic substitution (knock-in), but large insertions comprising endogenous/exogenous genes (Shan et al. 2013; Wu et al. 2013; Gratz et al. 2014).

4 The Versatile CRISPR Toolbox The programmability of the CRISPR-Cas system enables its application beyond genome editing. One such approach is the regulation of gene expression employing the catalytically inactive endonuclease, dead Cas9 (dCas9), which is unable to

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cleave the target DNA double-strand. Thus, a complex result from the fusion of dCas9 to transcriptional regulatory proteins can be addressed by sgRNAs to bind on cis-regulatory regions of a target DNA (Balderston et al. 2021). For instance, the fusion of dCas9 and KRAB (a heterochromatin inducer) can be used to repress gene expression – CRISPR interference (CRISPRi) – when directed to the promoter region of a gene, thereby achieving up to 15-fold gene down-regulation in human cells (Gilbert et al. 2013). Conversely, endogenous gene overexpression – CRISPR activation (CRISPRa) – can be performed by dCas9 fused to transcription activator modules, such as VP64 and SunTag (Balderston et al. 2021), which can elicit an 820-fold gene up-regulation (Perez-Pinera et al. 2013; Tanenbaum et al. 2014). To date, these CRISPR-based gene modulation approaches have already been strategically implemented in therapeutic contexts (Bialek et al. 2016; Matharu et al. 2019, Matharu et al. 2019). Nevertheless, some obstacles still impinge on CRISPRa/ibased therapies, such as nucleosome occupancy (Horlbeck et al. 2016) and dCas9 off-targets (Matharu et al. 2019). Another CRISPR system adaptation that has shown its importance to the field is CRISPR-based diagnostics (Fig. 1.4). A first example is the DNA endonuclease targeted CRISPR trans-reporter (DETECTR) method (Fig. 1.4 – left), which employs the Cas12a endonuclease as a biosensing molecule. In this approach, Cas12a binds to the target viral DNA (clinical sample) previously amplified by

Fig. 1.4 CRISPR-based nucleic acid detection mechanisms. At first, nucleic acid molecules are extracted from biological samples and amplified either by recombinase polymerase amplification (RPA) – for DETECTR (left) – or by reverse-transcription recombinase polymerase amplification (RT-RPA) followed by in vitro transcription (IVT) – for SHERLOCK (right). In both technologies, the Cas endonucleases, Cas12a and Cas13, remain inactive in the absence of the respective target nucleic acid molecules. The activation of these enzymes mediated by the detection of target molecules (red) leads to indiscriminate cleavage of off-targets, including reporter probes. Thus, the cleavage of these reporter molecules results in the release of their fluorophores (green), thereby emitting a stable fluorescence signal

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recombinase polymerase amplification (RPA), which triggers its non-specific collateral activity and degradation of single-stranded DNAs (ssDNAs). Then, as the ssDNA reporter (probe) is cleaved by activated Cas12a, a fluorescent signal is emitted and detected by a fluorimeter, indicating that the clinical sample was virus-positive (Chen et al. 2018). Alternatively, other methods for detecting RNA molecules have also been described (Gootenberg et al. 2017, 2018; Myhrvold et al. 2018; Ackerman et al. 2020; Broughton et al. 2020). The sensitivity enzymatic reporter unlocking (SHERLOCK) method uses previously RNA-amplified (reverse-transcription recombinase polymerase amplification – RT-RPA) and transcribed (in vitro transcription) samples to react with the Cas13 enzyme (Fig. 1.4 – right). Similar to DETECTR, the interaction of Cas13 with the target RNA molecule leads to enzymatic activation and consequent collateral cleavage of fluorescent reporters, emitting a signal readout (Gootenberg et al. 2017). Interestingly, these CRISPR-based strategies for detecting nucleic acids are already effectively suitable for monitoring highly incident viral infections, such as Zika, dengue, and SARSCoV-2. As a prospect, further optimizations might detect a wider range of humanassociated viruses and other diseases (Qiu et al. 2018; Ackerman et al. 2020). The dramatic rise of the CRISPR-Cas system since its discovery is astounding. Other features of the CRISPR toolbox (Table 1.1) have also shown their feasibility, such as CRISPR-based imaging (Qin et al. 2017), DNA labeling (Anton et al. 2014), RNA cleavage (O’Connell et al. 2014), gene mapping (Sadhu et al. 2016), and RNA Table 1.1 The comprehensive CRISPR toolbox Variation of the technique Gene editing Nick – reverse ranscription Stochastic marks Mutagenic chain reaction

Name CRISPR Prime editing GESTALT/MEMOIR MCR

Digital encoding Recording of cellular events High-fidelity SNP genotyping CRISPR-responsive smart materials Programmed chromosome fission/ fusion Gene mapping Nucleic acid detection RNA cleavage Regulation of gene expression Identification of chromatin interactions 3D positioning of genomic loci

– CAMERA/SCRIBE/record-Seq Cas14-DETECTR – –

Base editing Labeling/racking

REPAIR LiveFISH/CASFISH

Objective Modification Editing Lineage tracing Allelic dissemination Data storage Recording Genotyping Material plasticity Synthetic biology

– SHERLOCK – CRISPRi/CRISPRa CAPTURE

Localization Detection Expression

CRISPR-GO

Genomic positioning Modification Localization

Interactions

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tracking (Nelles et al. 2016), among others. Collectively, CRISPR-derived strategies are expected to contribute substantially to fundamental research, agriculture, industry, and medicine. Therefore, biomedical sciences have gained several innovative CRISPR-based applications, which are discussed in the following chapters.

References Ackerman CM, Myhrvold C, Thakku SG et al (2020) Massively multiplexed nucleic acid detection with Cas13. Nature 582:277–282. https://doi.org/10.1038/s41586-020-2279-8 Anders C, Bargsten K, Jinek M (2016) Structural plasticity of PAM recognition by engineered variants of the RNA-guided endonuclease Cas9. Mol Cell 61:895–902. https://doi.org/10.1016/ j.molcel.2016.02.020 Anton T, Bultmann S, Leonhardt H, Markaki Y (2014) Visualization of specific DNA sequences in living mouse embryonic stem cells with a programmable fluorescent CRISPR/Cas system. Nucleus 5:163–172. https://doi.org/10.4161/nucl.28488 Balderston S, Clouse G, Ripoll J-J et al (2021) Diversification of the CRISPR toolbox: applications of CRISPR-Cas systems beyond genome editing. CRISPR J 4:400–415. https://doi.org/10. 1089/crispr.2020.0137 Barrangou R, Fremaux C, Deveau H et al (2007) CRISPR provides acquired resistance against viruses in prokaryotes. Science 315:1709–1712. https://doi.org/10.1126/science.1138140 Bhaya D, Davison M, Barrangou R (2011) CRISPR-Cas systems in bacteria and archaea: versatile small RNAs for adaptive defense and regulation. Annu Rev Genet 45:273–297. https://doi.org/ 10.1146/annurev-genet-110410-132430 Bialek JK, Dunay GA, Voges M, Schäfer C, Spohn M, Stucka R, Hauber J, Lange UC (2016) Targeted HIV-1 latency reversal using CRISPR/Cas9-derived transcriptional activator systems. PLoS One 11(6):e0158294. https://doi.org/10.1371/journal.pone.0158294. PMID: 27341108; PMCID: PMC4920395. Bolotin A, Quinquis B, Sorokin A, Ehrlich SD (2005) Clustered regularly interspaced short palindrome repeats (CRISPRs) have spacers of extrachromosomal origin. Microbiology 151: 2551–2561. https://doi.org/10.1099/mic.0.28048-0 Broughton JP, Deng X, Yu G et al (2020) CRISPR-Cas12-based detection of SARS-CoV-2. Nat Biotechnol 38:870–874. https://doi.org/10.1038/s41587-020-0513-4 Brouns SJJ, Jore MM, Lundgren M et al (2008) Small CRISPR RNAs guide antiviral defense in prokaryotes. Science 321:960–964. https://doi.org/10.1126/science.1159689 Chen H, Choi J, Bailey S (2014) Cut site selection by the two nuclease domains of the Cas9 RNA-guided endonuclease. J Biol Chem 289:13284–13294. https://doi.org/10.1074/jbc.M113. 539726 Chen JS, Ma E, Harrington LB et al (2018) CRISPR-Cas12a target binding unleashes indiscriminate single-stranded DNase activity. Science 360:436–439. https://doi.org/10.1126/science. aar6245 Cong L, Ran FA, Cox D et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339:819–823. https://doi.org/10.1126/science.1231143 Deltcheva E, Chylinski K, Sharma CM et al (2011) CRISPR RNA maturation by trans-encoded small RNA and host factor RNase III. Nature 471:602–607. https://doi.org/10.1038/ nature09886 Gallagher DN, Haber JE (2018) Repair of a site-specific DNA cleavage: old-school lessons for Cas9-mediated gene editing. ACS Chem Biol 13:397–405. https://doi.org/10.1021/acschembio. 7b00760 Gilbert LA, Larson MH, Morsut L et al (2013) CRISPR-mediated modular RNA-guided regulation of transcription in eukaryotes. Cell 154:442–451. https://doi.org/10.1016/j.cell.2013.06.044

10

D. F. Contiliani et al.

Gleditzsch D, Pausch P, Müller-Esparza H et al (2019) PAM identification by CRISPR-Cas effector complexes: diversified mechanisms and structures. RNA Biol 16:504–517. https://doi.org/10. 1080/15476286.2018.1504546 Gootenberg JS, Abudayyeh OO, Lee JW et al (2017) Nucleic acid detection with CRISPR-Cas13a/ C2c2. Science 356:438. https://doi.org/10.1126/science.aam9321 Gootenberg JS, Abudayyeh OO, Kellner MJ et al (2018) Multiplexed and portable nucleic acid detection platform with Cas13, Cas12a, and Csm6. Science 360:439–444. https://doi.org/10. 1126/science.aaq0179 Gratz SJ, Ukken FP, Rubinstein CD et al (2014) Highly specific and efficient CRISPR/Cas9catalyzed homology-directed repair in drosophila. Genetics 196:961–971. https://doi.org/10. 1534/genetics.113.160713 Horlbeck MA, Gilbert LA, Villalta JE et al (2016) Compact and highly active next-generation libraries for CRISPR-mediated gene repression and activation. eLife 5:e19760. https://doi.org/ 10.7554/eLife.19760 Hsu PD, Lander ES, Zhang F (2014) Development and applications of CRISPR-Cas9 for genome engineering. Cell 157:1262–1278. https://doi.org/10.1016/j.cell.2014.05.010 Ishino Y, Shinagawa H, Makino K et al (1987) Nucleotide sequence of the iap gene, responsible for alkaline phosphatase isozyme conversion in Escherichia coli, and identification of the gene product. J Bacteriol 169:5429–5433. https://doi.org/10.1128/jb.169.12.5429-5433.1987 Ishino Y, Krupovic M, Forterre P (2018) History of CRISPR-cas from encounter with a mysterious repeated sequence to genome editing technology. https://journals.asm.org/doi/epub/10.1128/ JB.00580-17. Accessed 28 Mar 2022 Jansen R, Van Embden JDA, Gaastra W, Schouls LM (2002) Identification of genes that are associated with DNA repeats in prokaryotes. Mol Microbiol 43:1565–1575. https://doi.org/10. 1046/j.1365-2958.2002.02839.x Jiang F, Doudna JA (2017) CRISPR–Cas9 structures and mechanisms. Annu Rev Biophys 46:505– 529. https://doi.org/10.1146/annurev-biophys-062215-010822 Jiang W, Marraffini LA (2015) CRISPR-Cas: new tools for genetic manipulations from bacterial immunity systems. Annu Rev Microbiol 69:209–228. https://doi.org/10.1146/annurev-micro091014-104441 Jinek M, Chylinski K, Fonfara I et al (2012) A programmable dual-RNA–guided DNA endonuclease in adaptive bacterial immunity. Science 337:816. https://doi.org/10.1126/science. 1225829 Leenay RT, Maksimchuk KR, Slotkowski RA et al (2016) Identifying and visualizing functional PAM diversity across CRISPR-Cas systems. Mol Cell 62:137–147. https://doi.org/10.1016/j. molcel.2016.02.031 Makarova KS, Haft DH, Barrangou R et al (2011) Evolution and classification of the CRISPR–Cas systems. Nat Rev Microbiol 9:467–477. https://doi.org/10.1038/nrmicro2577 Mao Z, Bozzella M, Seluanov A, Gorbunova V (2008) DNA repair by nonhomologous end joining and homologous recombination during cell cycle in human cells. Cell Cycle 7:2902–2906. https://doi.org/10.4161/cc.7.18.6679 Matharu N, Rattanasopha S, Tamura S et al (2019) CRISPR-mediated activation of a promoter or enhancer rescues obesity caused by haploinsufficiency. Science 363:eaau0629. https://doi.org/ 10.1126/science.aau0629 Mojica FJM, Juez G, Rodriguez-Valera F (1993) Transcription at different salinities of Haloferax mediterranei sequences adjacent to partially modified PstI sites. Mol Microbiol 9:613–621. https://doi.org/10.1111/j.1365-2958.1993.tb01721.x Mojica FJM, Díez-Villaseñor C, Soria E, Juez G (2000) Biological significance of a family of regularly spaced repeats in the genomes of archaea, bacteria and mitochondria. Mol Microbiol 36:244–246. https://doi.org/10.1046/j.1365-2958.2000.01838.x Mojica FJM, Díez-Villaseñor C, García-Martínez J, Soria E (2005) Intervening sequences of regularly spaced prokaryotic repeats derive from foreign genetic elements. J Mol Evol 60: 174–182. https://doi.org/10.1007/s00239-004-0046-3

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Myhrvold C, Freije CA, Gootenberg JS et al (2018) Field-deployable viral diagnostics using CRISPR-Cas13. Science 360:444–448. https://doi.org/10.1126/science.aas8836 Nambiar TS, Baudrier L, Billon P, Ciccia A (2022) CRISPR-based genome editing through the lens of DNA repair. Mol Cell 82:348–388. https://doi.org/10.1016/j.molcel.2021.12.026 Nelles DA, Fang MY, O’Connell MR et al (2016) Programmable RNA tracking in live cells with CRISPR/Cas9. Cell 165:488–496. https://doi.org/10.1016/j.cell.2016.02.054 Nishimasu H, Ran FA, Hsu PD et al (2014) Crystal structure of Cas9 in complex with guide RNA and target DNA. Cell 156:935–949. https://doi.org/10.1016/j.cell.2014.02.001 O’Connell MR, Oakes BL, Sternberg SH et al (2014) Programmable RNA recognition and cleavage by CRISPR/Cas9. Nature 516:263–266. https://doi.org/10.1038/nature13769 Perez-Pinera P, Kocak DD, Vockley CM et al (2013) RNA-guided gene activation by CRISPRCas9–based transcription factors. Nat Methods 10:973–976. https://doi.org/10.1038/nmeth. 2600 Pourcel C, Salvignol G, Vergnaud GY (2005) CRISPR elements in Yersinia pestis acquire new repeats by preferential uptake of bacteriophage DNA, and provide additional tools for evolutionary studies. Microbiology 151:653–663. https://doi.org/10.1099/mic.0.27437-0 Qin P, Parlak M, Kuscu C et al (2017) Live cell imaging of low- and non-repetitive chromosome loci using CRISPR-Cas9. Nat Commun 8:14725. https://doi.org/10.1038/ncomms14725 Qiu X-Y, Zhu L-Y, Zhu C-S et al (2018) Highly effective and low-cost microRNA detection with CRISPR-Cas9. ACS Synth Biol 7:807–813. https://doi.org/10.1021/acssynbio.7b00446 Sadhu MJ, Bloom JS, Day L, Kruglyak L (2016) CRISPR-directed mitotic recombination enables genetic mapping without crosses. Science 352:1113–1116. https://doi.org/10.1126/science. aaf5124 Semenova E, Jore MM, Datsenko KA et al (2011) Interference by clustered regularly interspaced short palindromic repeat (CRISPR) RNA is governed by a seed sequence. Proc Natl Acad Sci 108:10098–10103. https://doi.org/10.1073/pnas.1104144108 Shan Q, Wang Y, Li J et al (2013) Targeted genome modification of crop plants using a CRISPRCas system. Nat Biotechnol 31:686–688. https://doi.org/10.1038/nbt.2650 Sternberg SH, Redding S, Jinek M et al (2014) DNA interrogation by the CRISPR RNA-guided endonuclease Cas9. Nature 507:62–67. https://doi.org/10.1038/nature13011 Tanenbaum ME, Gilbert LA, Qi LS et al (2014) A protein-tagging system for signal amplification in gene expression and fluorescence imaging. Cell 159:635–646. https://doi.org/10.1016/j.cell. 2014.09.039 Weiss D, Sampson T (2014) CRISPR-Cas systems: new players in gene regulation and bacterial physiology. Front Cell Infect Microbiol 4 Wu Y, Liang D, Wang Y et al (2013) Correction of a genetic disease in mouse via use of CRISPRCas9. Cell Stem Cell 13:659–662. https://doi.org/10.1016/j.stem.2013.10.016

Chapter 2

Modern Approaches to Mouse Genome Editing Using the CRISPR-Cas Toolbox and Their Applications in Functional Genomics and Translational Research Cintia J. Monteiro, David M. Heery, and Jonathan B. Whitchurch

1 Introduction The development of gene targeting approaches has enabled the generation of gene knockout, knock-in, mutant or conditionally expressed alleles and transgenes in animals used in research. Application of gene-targeting methods to zygotes or embryonic stem cells can facilitate the germline transmission of these genomic alterations. Many recent advances in the study of functional genomics in animal models have resulted from the development and application of bacterial CRISPRCas technologies. The discovery of CRISPR-Cas9 and its development into a range of simple, cost-effective and well-defined protocols has produced an invaluable toolbox of molecular techniques for genetic editing and regulation of animal genomes. These have allowed scientists to create animal models of human diseases which accurately mimic the genetic alterations seen in affected individuals, much easier and precisely than could be achieved before the CRISPR-Cas9 revolution. These animal disease models are highly clinically relevant due to their disease aetiology and often accurate presentation of the human disease phenotype. A wealth of knowledge has been discovered about the molecular mechanisms contributing to disease development in cancer, metabolic disorders and neurological conditions like Alzheimer’s, as well as congenital conditions such as heterotaxia, sickle cell anaemia and cystic fibrosis, to name just a few areas of biomedical research where

C. J. Monteiro Department of Genetics, Molecular Immunogenetics Group, Ribeirão Preto Medical School, University of São Paulo, Ribeirão Preto, SP, Brazil D. M. Heery School of Pharmacy, University of Nottingham, Nottingham, UK J. B. Whitchurch (✉) Mammalian Genetics Unit, MRC Harwell Institute, Oxfordshire, UK © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_2

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CRISPR-Cas has been utilised to great benefit in the production of animal models of human disease. While elucidating the genetic contribution to a disease can be regarded as fundamental in truly understanding the aetiology, having clinically relevant and well-characterised animal models of disease also allows researchers to test promising therapeutics identified from cell-based assays, in vivo. Many CRISPR-Cas-generated animal models of human diseases have been utilised in these investigations, with some demonstrating partial alleviation of symptoms with novel treatments or even the apparent absence of disease after experimental gene therapy. Such exciting reports of therapeutic breakthroughs offer hope for people who have incurable genetic diseases. It wouldn’t be an overstatement to say that CRISPR-Cas technology has and will no doubt continue to have a very profound effect upon biomedical research, with the use of CRISPR-Cas to study mammalian genomes likely to benefit human health and enhance the understanding of our own genome on a monumental scale. The animal genome in which CRISPR-Cas has been most widely used, is undoubtedly the mouse, one of a geneticist’s best friends when studying mammalian genetics and whose contribution to research has been immense. Editing and regulation of this organism’s genome via CRISPR-Cas will be the main focus of this chapter; however the humble mouse is not alone when it comes to encounters with the CRISPR-Cas toolkit. An ever-increasing list of organisms are having their genomes modified using these techniques. Here, we will review some of the latest applications of CRISPR-Cas technologies in mice, as well as delivery methods and the use of alternative Cas enzymes for base editing, prime editing and transcriptome targeting. We will also discuss examples where CRISPR-Cas technologies have been applied to target clinically relevant genes in mice, either to generate humanised models of disease or for experimental gene therapies.

2 CRISPR-Cas Systems: An Expanding Toolkit for Mouse Genome Editing Prokaryote or archaeal CRISPR-Cas systems perform adaptive immunological defence roles to suppress foreign DNA or RNA from viruses or other sources. Systems such as the Streptococcal Cas9 DNA endonuclease have been successfully exploited for genome editing in mammalian cells and subsequently adapted for a range of further applications. Used in combination with short (20 bp) guide RNAs (gRNAs), Cas9 enzymes can be targeted to specific complementary sequences within a genome. The fully active Cas9 endonuclease consists of two nuclease subunits, HNH and RuvC. These cleave opposing DNA strands to induce a blunt end double strand break (DSB), although cleavage is dependent on the presence of a protospacer adjacent motif (PAM), a nucleotide sequence which must be present at the targeted locus. Repair of DSBs through non-homologous end joining (NHEJ) leads to insertions or deletions (indels) at the target site, or template-mediated

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homology-directed repair (HDR) can be performed using a co-delivered repair template. Studies on the incidence of CRISPR-Cas9 off-target endonuclease activity in microinjected mouse embryos revealed that off-target DSBs can be greatly reduced by using nickase versions of Cas9. These nickase Cas9 enzymes are referred to as Cas9n and have amino acid substitutions which render one of their nuclease subunits inactive (RuvC D10A or HNH H840A). Nickases can only induce singlestrand breaks which are efficiently repaired by base excision repair (BER); thus any off-target sites remain as the wild-type sequence. Therefore, when using Cas9n, editing of the target sequence actually requires dual targeting with two gRNAs within close proximity, one for each of the opposing DNA strands. The Cas12A endonuclease offers an alternative to Cas9 gene editing. Cas12a (also known as Cpf1) differs from Cas9 in a number of ways, e.g. it has two RuvC type endonuclease domains and rather than producing a blunt end DSB, it produces staggered overhangs. Furthermore, this endonuclease recognises a different PAM sequence (TTTV vs. NGG for Cas9), which may be useful to target genomic regions that lack Cas9 PAM sequences, thereby expanding the number of loci able to be subjected to CRISPR-Cas. In addition, Cas12a appears to have reduced off-target effects compared to the Cas9 endonuclease, potentially making it more appealing for use in both research and any future clinical applications.

3 Delivery of the CRISPR-Cas Toolbox The application of CRISPR-Cas technologies for germline engineering of mouse genomes has offered the promise of a more rapid and efficient approach for generating useful genetically engineered mouse models (GEMMs). This usually involves the delivery of Cas9 and gRNAs as DNA vectors or as combined Cas9-gRNA ribonucleoprotein complexes into zygotes or ES cells using microinjection, electroporation or viral delivery methods. A standard approach tried in many laboratories using mice involves the CRISPR-Cas-mediated knock-in of LoxP (flox) sites flanking a target sequence, usually an exon. This now common technique is achieved by microinjection of mouse zygotes or ES cells with Cas9-gRNA ribonucleoprotein complexes targeting two genomic loci, along with co-delivery of single-stranded oligo DNA nucleotides (ssODNs). ssODNs are utilised by the HR DNA repair mechanism to repair the dsDNA breaks induced by Cas9 activity. Mice born from this approach are then screened to ensure the desired locus has been floxed. Once established, conditional deletion of the floxed region is later achieved by mating mice with the floxed allele with mice which express Cre recombinase (Cre recombinase driver strains). Offspring which receive both the floxed allele and an allele which facilitates the expression of the Cre recombinase will have the DNA sequence between the two LoxP sites excised. This can be used to facilitate excision in all cells, or just in a particular tissue or cell type if the right Cre recombinase driver strain is selected.

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Gurumurthy et al. (2019a, b) recently reviewed the efficacy and reproducibility of producing conditional gene knockouts using the two-donor floxing approach. This is when two gRNAs and two donor ssODNs are microinjected into zygotes with the aim of introducing two LoxP sites, a technique which had a reported 16% success rate (Yang et al. 2013). Surprisingly, an analysis of data compiled for 56 different genomic targets from multiple research institutions revealed the efficacy of the method to be much lower than initially reported ( T), allowing the molecular and physiological characterisation of this genetic disease with a high degree of relevance to those affected individuals (Rauch et al. 2018). Alternatively, a different approach was used to generate genetically relevant mouse models of Duchenne muscular dystrophy (DMD), a recessive X-linked form of muscular dystrophy caused by mutations in the human DMD gene that alter the reading frame, introduce stop codons or are pathogenic substitutions. Mutations in this gene compromise myofiber integrity and drive the deterioration of muscles, leading to muscle weakness and muscle wasting. To create a single base pair substitution in the murine Dmd gene and generate a mouse model of DMD, Kim et al. (2017a, b) used a base editor composed of a cytidine deaminase fused to a nuclease-inactive CRISPR-Cas9 enzyme, facilitating the conversion of cytosine to uracil at a specific locus. This resulted in a C > T nucleotide conversion after erroneous repair or DNA replication. The authors delivered mRNA encoding their base editor or ribonucleoproteins targeting the Dmd gene into mouse zygotes and were successful in both cases (Kim et al. 2017b). Mutations in the transcription factor SRY-box 9 (SOX9) gene cause campomelic dysplasia (CD) and acampomelic campomelic dysplasia (ACD), developmental disorders of cartilage. ACD is yet another example where the use of CRISPRCas9 has been applied to create a mouse model of a human genetic disease. In this case, a mouse model of this disease was generated by deleting the rib cage specific Sox9 enhancer via the microinjection of zygotes with Cas9 mRNA and gRNA (Mochizuki et al. 2018). The study of this mouse line has provided a wealth of information regarding the spatiotemporal regulation of Sox9 transcription, while also demonstrating that disruption of the rib cage-specific Sox9 enhancer causes the development of a narrower and shorter ribcage in mice, a feature of ACD in humans. Mouse models for human cancer research have proven to be of utmost importance as tumour biology shares similar genomic and physiological characteristics in mice and humans. Platt et al. (2014) established a Cre-dependent Cas9 knock-in mouse and simultaneously modelled the dynamics of p53 and Lkb1 loss of function and Kras gain of function mutations. These three genes are the top three most frequently mutated genes in lung adenocarcinoma in humans (Collisson et al. 2014). Other in vivo cancer mouse models generated by CRISPR-Cas9 tools include medulloblastoma, glioblastoma and several leukaemias, to name just a few. Zuckermann et al. (2015) targeted the tumour suppressor genes Ptch1, Trp53, Pten and Nf1 in the

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mouse brain via electroporation of Cas9-gRNA vectors into the cerebrum, resulting in the knockout of these genes and the subsequent development of glioblastoma. Results from this research have demonstrated that CRISPR-Cas9 tools can be successfully applied to the challenge of generating appropriate animal models of human cancers. Aside from diseases of the neurological or muscular systems, bone or blood disorders and cancer, mouse models are also highly valuable in the study of metabolic diseases. The use of CRISPR-Cas9 by Roh et al. (2018), whereby they produced knockout mice for the leptin and leptin receptor genes, is a good example. Leptin is a hormone that acts within the hypothalamus by binding to its aptly named receptor, functioning to regulate blood sugar concentration, fat synthesis inhibition and appetite suppression. The study of this hormone and its signalling pathway has been of great interest, partly due to the worldwide increase in levels of population obesity and the association of leptin with appetite and metabolism. The mice generated by Roh et al. (2018) showed phenotypic disorders very similar to those observed in humans who possess genetic variants in the leptin, or leptin signalling pathway genes such as an increase in body weight, hyperglycaemia, hepatic steatosis and diabetes.

6 Humanised Mouse Models of Disease Mouse models of human diseases have been essential in biomedical research and have contributed immeasurably to the development of treatments for diseases. However, limitations exist due to differences between mice and humans in several aspects. One such difference can be observed in the DNA sequence homology between orthologues or their regulatory elements. Thus, there are always questions which arise regarding how translatable to human diseases the results of mouse studies are. One cannot merely assume that findings from mouse models of human diseases will be exactly the same in human patients; the same can be said for the development of treatments, including gene therapies. A possible solution to this issue is the generation of humanised mice (Fig. 2.1). These are mice in which their cells have undergone humanisation, i.e. they are engrafted with functional human cells or tissues, or a human gene or regulatory element has been knocked into a genomic locus as a substitute for the endogenous mouse sequence, which often has been removed from the genome (Devoy et al. 2011; Fujiwara 2018). The two main approaches which have been extensively used over the years to deliver human genes into the mouse genome are cDNA constructs and human genecontaining bacterial artificial chromosomes (BACs) (Hosur et al. 2017). BAC transgenesis is a useful approach when large insertions up to 300 kb are required, for example, when it is necessary to retain introns and regulatory elements as part of the transgene. In recent years, the CRISPR-Cas9 system has been used to generate humanised mouse models because it allows a more precise insertion into the genome

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Fig. 2.1 The CRISPR-Cas9 system can be used to generate humanised mice by the replacement of mouse genes with their human orthologues. It is also possible to use this system to edit specific genomic loci or regulatory elements outside of protein-coding sequences. The human RNA and protein (if applicable) are then expressed in the mouse. (The figure was prepared using the Motifolio Illustration Toolkit)

and is more efficient, cost-effective and experimentally faster than some traditional alternative methods (Jin and Li 2016). When utilising CRISPR-Cas to create humanised mouse models of disease, one strategy is to insert human cDNA into the mouse genome while concomitantly knocking out the endogenous mouse gene at that locus. This method has been demonstrated to introduce a human gene at the desired locus which will be under the control of the endogenous regulatory elements, which therefore is likely to have an expression pattern similar to that of the endogenous sequence (Fig. 2.1) (Miura et al. 2015). A second strategy is to insert cDNA into a genomic ‘safe-harbour’ locus, which allows the expression of the human gene at a site away from the endogenous mouse gene. These safe-harbour loci are integration sites which are chosen because transgene insertion does not impact upon surrounding mouse genes. Gt(ROSA)26Sor, usually referred to as Rosa26, is the most common example of this safe-harbour locus in mice. Rosa26 produces a long non-coding RNA (lncRNA) under the control of a constitutive promoter in the mouse (Zambrowicz et al. 1997). The use of CRISPR-Cas9 to integrate a transgene into a safe-harbour locus like Rosa26 is performed when the aim is to insert a human gene without replacing the mouse orthologue. In this case, the human transgene may have minimal unintended phenotypic consequences, as the endogenous mouse gene is still present (Irion et al. 2007; Chu et al. 2016). A third strategy for generating humanised mouse models of disease using CRISPR-Cas is to generate point mutations in the endogenous mouse gene. If there is a high level of nucleotide sequence homology between the murine

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and human genes, this may be the most appropriate approach to take as smaller genomic changes are made and there is less risk of unintended disruption to nearby sequence elements. Alternatively, individual exons in a mouse gene can be exchanged with those from a human orthologue, if only particular exons deviate in their DNA sequence homology (Singh et al. 2015). The use of humanised mice in research is a significant step towards the translation of CRISPR-Cas9 gene editing from experimental tool to therapeutic agent. In Duchenne muscular dystrophy (DMD), a disease caused by variants in the X-linked Dystrophin (DMD) gene, deletions flanking exon 51 disrupt the reading frame and are the most common cause of this condition. Several studies have successfully restored the correct reading frame in different mouse models of DMD using CRISPR-Cas technologies to create small indels or perform exon skipping (Amoasii et al. 2017; Min et al. 2020; Zhang et al. 2022b). However, the nucleotides which Cas9 gRNAs complementary base pair with in these studies are not conserved between the mouse and human genomes, preventing translation of these approaches as gene therapies into clinical practice. For this reason, in an additional study, researchers opted to generate a humanised DMD mouse model by replacing exon 51 of the murine Dmd gene with human exon 51, while also deleting exon 50 to cause ORF disruption (Zhang et al. 2022b). CRISPR-Cas9-mediated correction of the reading frame restored the level of dystrophin protein expression by 20–25% and ameliorated the pathologic hallmarks of DMD. Grip strength was increased, muscle degeneration decreased and improved histopathological markers of the disease such as less inflammatory infiltration and necrosis were observed. CRISPR-Cas9 has also been utilised to create a humanised mouse model of glycosylphosphatidylinositol (GPI) biosynthesis deficiency (de los Santos et al. 2021). GPI is a phosphoglyceride which can be post-translationally added to the C-terminus of a protein. PIGV encodes an enzyme essential for this process, and mutations in the PIGV gene compromise the function of GPI-linked proteins, leading to intellectual disability, global developmental delay, renal anomalies and, in some cases, epilepsy (Horn et al. 2014). De los Santos et al. (2021) introduced into mice the most prevalent hypomorphic missense mutation in European patients with GPI deficiency: Pigv:c.1022C > A (p.A341E). This locus is conserved in mice, and therefore only a single base change was required to generate the pathogenic variant. The humanised Pigv341E mice successfully mirrored the human pathology of GPI deficiency and have allowed deeper study of the pathophysiology underlying this condition. Research using this mouse line revealed reduced synaptophysin immunoreactivity and decreased hippocampal synaptic transmission as characteristics underlying the disease, leading to impaired memory formation. The G-protein-coupled receptor GPRC6A regulates energy metabolism, and in the human population, the GPRC6A gene displays a KGKY polymorphism. Until recently, the influence of the KGKY polymorphism in the third intracellular loop (ICL3) was still unclear. Pi et al. (2020) applied CRISPR-Cas9 technology to replace the RKLP sequence in the third intracellular loop of gprc6a in mice with the human KGKY sequence, to create humanised Gprc6aKGKY mice. These mice presented with reduced basal blood glucose levels and both increased circulating serum insulin and

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Fgf21 concentration. Furthermore, Gprc6aKGKY mice had improved glucose tolerance and enhanced pyruvate-mediated gluconeogenesis. This study demonstrates that CRISPR-Cas9 can be used to successfully generate mice harbouring human sequence polymorphisms, allowing the investigation of such polymorphisms to determine whether there are any biological consequences or advantages. The authors concluded that the human GPRC6A KGKY polymorphism is a gain-of-function variant which likely positively controls energy metabolism, as it does in mice. Haemophilia A is an X-linked blood disorder caused by a deficiency in blood clotting factor VIII, leading to spontaneous bleeding in joints or organs and excessive bleeding during trauma which can be life-threatening. Chen et al. (2019) generated mice which have a permanent chromosomal integration of a modified human clotting factor VIII sequence within the albumin (Alb) locus in liver cells. To do so, mice received tail vein injections of separate AAVs containing Cas9-gRNA to target Alb intron 13 and also deliver the repair template. In FVIII KO mice, the modified factor VIII sequence was inserted at the Alb locus, and human FVIII protein expression was detected in the liver of treated mice. This gene therapy approach resulted in loss of the disease phenotype, increased FVIII activity for 7 months and no liver toxicity, demonstrating therapeutic benefits for those individuals with haemophilia A. Humanised mice have also been generated by CRISPR-Cas to study the human immune system, e.g. to investigate mechanisms that contribute to transplant rejection. One example is the NOG B2m KO mouse generated by Ka et al. (2021), which possesses a nonsense mutation in exon 1 of the B2m gene and was generated using Cas9-gRNA ribonucleoprotein microinjection into zygotes. B2m encodes a component of the class I major histocompatibility complex (MHC I). Mating these mice with ones who are null for a component of the MHC II complex created a modified NOG-major histocompatibility complex class I/II double knockout (dKO-em) mouse model. When human peripheral blood mononuclear cells (PBMC) were transplanted into the dKO-em mice, a high engraftment efficiency was observed with no noticeable graft-versus-host disease. dKO-em mice offer an advantage in the study of PBMC engraftment compared to previous mouse models, due to this lack of graft-versus-host disease. Furthermore, when comparing the modified dKO-em mice with previously reported dKO mice, it was observed that data reproducibility was improved. Thus, these humanised dKO-em mice will help facilitate the development of PBMC-based novel therapeutics. Amyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD) are two neurodegenerative diseases which lie on the same disease spectrum. The lifetime risk of developing ALS is 1 in 300 in the UK, so understanding its aetiology could benefit many people if research leads to clinical advancements in ALS disease management/treatment (Alonso et al. 2009). Mutations in several genes have been associated with enhanced risk of developing these diseases, one of which is TARDBP. Devoy et al. (2021) generated Tardbp humanised mice using CRISPRCas9 for the study of ALS and FTD. These humanised mice express the human orthologue at physiological levels in several tissues, with correct splicing of transcripts and normal human protein biochemistry. To generate the mice, a plasmid

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encoding CRISPR-Cas9 and gRNA was electroporated into embryonic stem cells alongside a bacterial artificial chromosome (BAC), harbouring the human genomic sequence. HDR replaced the murine genomic sequence with that of the human orthologue. These mice will be of huge benefit for the assessment of therapeutic agents in the treatment of ALS and FTD, overcoming issues associated with using transgenic models of human diseases.

7 Testing of Gene Therapies in Mouse Models of Human Disease While the contribution of CRISPR-Cas9 technology to improvements in the generation of mouse models for better understanding of human diseases cannot be understated, its uses are rapidly extending beyond the study of disease aetiology. Many studies have also demonstrated the use of CRISPR-Cas genome editing in gene therapy. The combination of Cas9 enzyme, guide RNA and DNA repair template enables precise genome editing and, consequently, accurate alteration of genetic variants associated with disease. In other words, the ability to change a known pathogenic variant at a specific genomic locus into a wild-type, or otherwise non-pathogenic sequence of DNA. One can imagine that such an approach may be able to prevent the onset of disease, halt/delay its progression and, in an ideal world, even be able to reverse any effects a disease may have already had. CRISPR-Cas9 has been used to generate a mouse model for the study of Huntington’s disease; this inherited neurodegenerative disorder is fatal and primarily caused by CAG repeat expansion within the Huntingtin gene (HTT), which results in early deterioration of motor functions, cognitive decline and psychological changes (Nopoulos 2016). Not only has CRISPR-Cas been used to generate relevant models of human genetic diseases in mice but also to demonstrate gene therapy approaches. Monteys et al. (2017) developed a strategy based on CRISPR-Cas9 technology that takes advantage of highly prevalent SNPs in the HTT gene. They identified SNPs that generate new PAM sequences in pathogenic alleles, therefore making them susceptible to targeting with Cas9. gRNAs were designed to direct Cas9 endonuclease activity to such PAM sequences in a HTT transgenic mouse model of this disease; delivery of Cas9 and gRNAs via AAVs reduced transcription of the pathogenic allele in mouse brain tissue. Using a related but alternative approach, Tabebordbar et al. (2016) induced exon deletion via CRISPR-Cas9 with the aim of halting and recovering muscle function in a mouse model of DMD (the mdx mouse). For this purpose, CRISPR-Cas9 coupled with a pair of guide RNAs flanking exon 23 of the pathogenic Dmd allele was delivered by AAV, which resulted in successful excision of the intervening DNA sequence. Importantly, this restored the reading frame of dystrophin in myofibers, cardiomyocytes and muscle stem cells, with dystrophin protein expression able to be detected. A separate study which used a very similar approach in both adult and neonatal mice also reported promising results. Partial recovery of functional

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Dystrophin protein in skeletal myofibers and cardiac muscle was observed, as well as significant enhancement of muscle force (Nelson et al. 2016). CRISPR-Cas9-loaded nanocomplexes have been shown to be an effective delivery system for the combined delivery of Cas9 mRNA with gRNA or Cas9-gRNA ribonucleoproteins. A notable use of this delivery system is the targeting of Betasecretase 1 (Bace1) in neurons of the adult mouse brain to treat Alzheimer’s disease. Bace1 is necessary for the formation of Aβ peptides, of which intraneuronal accumulation is a typical characteristic of this disease, alongside synaptic dysfunction and memory impairment. CRISPR-Cas-targeting of Bace1 has been shown to reduce the onset of amyloid beta (Aβ)-associated pathologies and cognitive defects in two mouse models of Alzheimer’s disease (Park et al. 2019). Findings such as these raises hope for the development of a proactive treatment for this progressive neurological condition. AAV delivery of CRISPR-Cas9 in mice has been used to deplete Vegfr2 expression in vascular endothelial cells, with a reduction in protein expression observed (Huang et al. 2017). VEGFR2 is crucial for normal angiogenesis in humans; however abnormal angiogenesis is associated with retinopathies such as proliferative diabetic retinopathy and other conditions like neovascular age-related macular degeneration; this process also has a major role in tumour growth and metastasis. Research by Huang et al. (2017) demonstrated the abrogation of angiogenesis in a mouse model of oxygen-induced retinopathy that had intravitreal injection of Cas9 and gRNA AAVs, with a 30% reduction in Vegfr2 protein level in retinal lysates. Another member of the Vegf family which has been subjected to CRISPR-Cas as a therapeutic target is Vegfa. Kim et al. (2017a) targeted the Vegfa gene with the aim of treating age-related macular degeneration (AMD) in a mouse model of this human disease, which is a leading cause of blindness in adults. In order to deliver Vegfa gene-specific Cas9 ribonucleoproteins into the adult mouse eye, a subretinal injection was performed to target the correct site in the retinal pigment epithelium. This resulted in a decrease in Vegfa protein level and a reduction in the area of laserinduced choroidal neovascularization. Such studies have expanded the scope of applications for CRISPR-Cas9 in potential gene therapies, as they have demonstrated the potential of the CRISPR-Cas toolbox in the local treatment of degenerative ocular diseases. Haemophilia B is an X-linked blood clotting disorder caused by mutations in the coagulation factor IX gene. Individuals with severe cases of this condition require regular intravenous administration of the coagulation factor, and without this routine treatment, they experience bleeding of joints, inflammation and pain, frequent nosebleeds and bruise easily. The application of CRISPR-Cas in a mouse model of this disease and the induction of HDR successfully ablated the disease phenotype (Ohmori et al. 2017). Treated mice had increased expression of factor IX protein in hepatocytes; these mice presented normal haemostasis after treatment, including faster blood coagulation and reduced bleeding time. Such demonstrations of successful gene therapy offer exciting prospects for human patients who have the disease(s) in question, although great care should be taken to consider the ease, efficacy and ethical considerations of these methods.

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As we mentioned previously, CRISPR-Cas9 technology is being utilised in the mouse to generate mouse models of human metabolic diseases. Similar to the other mouse models of disease discussed here, CRISPR-Cas is increasingly being applied to tackle the challenge and promise of gene therapy, which also includes metabolic diseases. A recent example is the study by Zhu et al. (2021), which performed in situ genome editing in a monogenic obesity mouse model (Ob/ob) to correct a pathogenic C to T variant in exon 2 of the Lep (leptin) gene. This variant causes congenital Leptin protein deficiency, a rare autosomal recessive obesity syndrome. Conversion of this variant to the wild-type DNA sequence in preadipocytes restored expression of the normal Leptin protein when they were differentiated into mature adipocytes. Promisingly, treated mice displayed suppression of appetite and insulin resistance was no longer observed, demonstrating that CRISPR-Cas-based gene therapy may hold promise for individuals suffering from genetic metabolic conditions, as it does for many other diseases. Tyrosinemia type I is another metabolic disorder which has shown positive results when subjected to CRISPR-Cas genome editing. This condition results from an error of metabolism in which the amino acid tyrosine is not metabolised completely due to a deficiency of the enzyme fumarylacetoacetate hydrolase (FAH). FAH deficiency causes liver problems due to an accumulation of toxic metabolites, affected individuals fail to thrive and food intolerances are observed. Yin et al. (2016) generated Fah-positive hepatocytes in a mouse model of tyrosinemia via Cas9-mediated correction of the pathogenic splicing disrupting mutation. The authors of this study utilised a combination of AAV and nanoparticle-mediated delivery of Cas9 mRNA, gRNA and DNA repair template. Treated mice presented with significantly less liver damage and increased Fah mRNA expression. More recently, another team of researchers also treated tyrosinemia type I in a mouse model using CRISPR-Cas-based gene therapy (Song et al. 2020). Rather than using Cas9 endonuclease activity, they performed a tail-vein injection of plasmid DNA encoding an adenine base editor (ABE) with a gRNA. This corrected the A > G splice site mutation present in their mouse model of tyrosinemia, partially restored correct splicing of Fah mRNA and, importantly, generated Fah-positive hepatocytes. Treated mice had normal body weight and appeared to be healthy. While CRISPR-Cas-based gene therapy approaches have clearly had success in mouse models of human genetic diseases, the CRISPR-Cas9 system has also been used as a treatment for AIDS (acquired immune deficiency syndrome). In a proof-ofconcept study performed by Dash et al. (2019), viral clearance was observed in HIV-infected humanised mice after long-acting slow-effective release antiviral therapy (LASER ART) in conjunction with CRISPR-Cas9. Cas9 endonuclease activity cleaved HIV proviral DNA within the long terminal repeats (LTRs) and the Gag gene, regions which are highly conserved. Disruption of integrated HIV sequences by excision of the intervening nucleotides between cleavage sites was calculated to be 60–80%. The efficiency of viral replication suppression was further enhanced by LASER ART treatment, with an increase in CD4+ T-cells and a drop in plasma viral load below the detection limit observed. These results suggest that the use of CRISPR-Cas in gene therapy may one day extend to eradicating integrated viral genomes like HIV, which is an alluring proposition for affected individuals and clinicians alike.

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8 Prime Editing and Transcriptional Regulation Using CRISPR-Cas Other applications of CRISPR-Cas technology include the fusion of additional functional domains to a nuclease-deficient version of Cas9, known as dCas9. This has been applied to gene activation (CRISPRa) or inhibition (CRISPRi) through the use of transcriptional repression or activation domains fused to dCas9. Domains from transcriptional activators such as p300 and VP64, or repressors like KRAB, induce specific epigenetic modifications of chromatin, influencing gene expression. There are several examples of dCas9 fusions being used to regulate the transcription of targeted genes in mice, but here we will discuss just a few select reports. Liao et al. (2017) successfully used CRISPR-Cas to activate target gene transcription in mice via tail vein injections of AAVs containing a dCas9 fusion protein and specific gRNAs. In a mouse model of acute kidney injury, they managed to demonstrate improved renal function, reduced histological features associated with kidney injury and also increased survival time. The same approach was able to produce insulinsecreting cells in a mouse model of type-I diabetes, increasing serum insulin levels and reducing blood glucose concentration, simply by changing the gRNAs to direct the transcriptional activation activity of their dCas9 complex to Pdx1, a gene necessary for the differentiation of liver cells into pancreatic insulin-secreting ones. Similarly, increasing transcription of the Klotho gene in the mdx mouse model of Duchenne muscular dystrophy improved muscle strength, partially ameliorating the disease phenotype. The use of conditional Cre dCas9 transgenes at the Rosa26 locus has also been demonstrated (Gemberling et al. 2021). Conditional expression of dCas9-p300 or dCas9-KRAB have both been shown to successfully regulate the transcription of target genes in mouse tissues, including CD4+ T-cells. dCas9-KRAB expression by AAV delivery and targeting of this synthetic transcriptional regulator complex to Pcsk9, a gene involved in the control of serum cholesterol levels, repressed its expression and reduced serum cholesterol concentration for 24 weeks after only a single treatment (Thakore et al. 2018). This demonstrates the potential for long-lasting effects with the use of CRISPR-Cas technologies for transcriptional regulation of disease-associated genes. Further research is clearly needed to explore the therapeutic potential of these approaches and how efficacies may vary across tissue types, investigate the specificity of activator/repressor complexes and identify any long-term implications. Cytosine or adenine base editors such as APOBECs or AIDs fused to dCas9 have also been developed to enable single nucleotide editing and have been successfully utilised to generate GEMMs. These can be used to recapitulate clinically relevant mutations, e.g. as performed for the Hoxd13 and androgen receptor (Ar) genes, or to demonstrate gene therapy such as correction of nonsense mutations in the Duchenne muscular dystrophy gene dmd (Kim et al. 2017b; Liu et al. 2018; Ryu et al. 2018). Base editing technologies enable specific editing of single nucleotides without the need for a DSB and the induction of DNA repair pathways, circumventing the risks associated with DNA cleavage, many of which still remain a potential concern for the clinical application of gene editing technologies.

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Another novel Cas9 technology that avoids DSBs and therefore theoretically reduces the likelihood of off-target effects is prime editing. This consists of a Cas9 nickase fused at the C-terminus to a reverse transcriptase, used in conjunction with an edit-containing guide RNA referred to as a prime editing guide RNA, or pegRNA. The nickase-pegRNA complex induces a nick in the non-complementary DNA strand precisely 3 bp upstream of the PAM sequence, exposing a DNA ‘flap’ which hybridises to the pegRNA and acts as a primer (Scholefield and Harrison 2021). The fused reverse transcriptase then extends the 3’-OH flap using the pegRNA as a template, thus incorporating the desired edit into the DNA, and the 5’ flap is preferentially removed by the endogenous endonuclease FEN1. Mismatch repair then resolves mismatch(es) between the edited and opposite DNA strands to either give the desired precise edit or revert back to the wild-type sequence. However, wild-type alleles are susceptible to further prime editing events which are likely to result in the eventual incorporation of the alternative nucleotide sequence. Mutation of the PAM sequence can be built into the experimental design to prevent further change once the desired edit is achieved. Optimised prime editing strategies that reduce the incidence of unwanted by-products are still in development for use in mouse embryos, as undesired outcomes such as large deletions have been reported to have a high frequency (Aida et al. 2020). Nevertheless, prime editing has been successfully used in mice to create clinically relevant variants in several genes, plus correct pathogenic mutations in both genetic eye and liver diseases (Liu et al. 2020; Jang et al. 2021; Böck et al. 2022). Prime editing in mice is currently in its infancy, and no doubt this technique will become more heavily used and develop further over time.

9 Genomic Safe-Harbour Loci as Sites for Stable Integration and Expression Specific loci within a genome where stable DNA insertions can be made and these insertions do not appear to be detrimental to cells are referred to as genomic safeharbour loci (or safe-harbour sites). A requirement for these sites is that they ensure the stable and robust transcription of the transgene, usually incorporating an artificial promoter within the inserted expression cassette, although this is not always necessary and depends on experimental design. They can be either intergenic or intragenic, however it is important that they do not influence the transcription of other genes, i.e. they should not alter the transcriptome. Many primary cells, embryonic stem cells, iPSCs and stable cell lines have been subjected to transgene insertion at a safe-harbour locus. These loci have also been utilised for the generation of desired transgenic mammals, most notably the mouse. Since these genomic loci offer the stable integration of transgenes and the knock-in is usually performed via microinjection at the single cell zygote stage, transgenes can be passed on to future generations as they will exist within the gametes of genetically modified adults. All the genomic safe-harbour loci described in this chapter have been successfully targeted with CRISPR-Cas9.

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Probably the best-known example in the mouse genome is the Rosa26 safeharbour site. This popular choice is used due to its ability to express constitutively throughout embryonic development and into adulthood and was identified via a gene trap experiment in mouse embryonic stem cells in the 1990s (Zambrowicz et al. 1997). Since then, hundreds of transgenic mouse strains have been created using this locus, alongside a vast number of genetically edited cells. With Rosa26, researchers have the option to make use of the endogenous promoter or include an artificial one with the transgene. Orthologues of the murine Rosa26 locus have been identified in humans, rabbits, rats, ferrets, pigs, sheep and cows (Irion et al. 2007; Wu et al. 2016; Yu et al. 2019; Yuan et al. 2021). Research has demonstrated that CRISPR-Cas9 can be utilised to knock-in transgenes into the murine Rosa26 locus, with up to 20% of live pups having the correct insertion (Chu et al. 2016). They microinjected pronuclear stage zygotes with Cas9 mRNA, gRNA and a donor template for HDR, although it is possible that using Cas9 protein rather than mRNA may enhance the chance of correct construct knock-in. Several mouse lines with insertions at the Rosa26 locus have been developed as reporters or even express the Cas9 endonuclease themselves (Platt et al. 2014; Chu et al. 2016). Those which act as reporters can provide information on cellular processes such as visual confirmation of Cre recombinase activity and help answer questions regarding cell lineage, cell migration during development (perhaps by live cell imaging) and spatial analysis of gene expression (Hasegawa et al. 2013; Rovira et al. 2021). A lesser-used genomic safe-harbour locus is Hipp11, which has been found and successfully utilised in mice, pigs and humans (Hippenmeyer et al. 2010; Tasic et al. 2011; Zhu et al. 2014; Ruan et al. 2015; Browning et al. 2020). This particular locus does not contain an endogenous promoter, is intergenic and can be targeted for knock-in with ease using CRISPR-Cas (Ruan et al. 2015; Li et al. 2019; Browning et al. 2020). Transgenic mice which express Cas9 or gRNAs themselves can be of great benefit, as they can be mated with other transgenic lines to produce genetically edited offspring. For example, mouse strains which carry a Cas9 transgene at the Rosa26 locus can be crossed with gRNA-expressing mice that contain the desired gRNA also at the Rosa26 locus. Mating homozygotes of these produces offspring that express both Cas9 and the gRNA, resulting in genome editing at the desired locus (Platt et al. 2014; Zhang et al. 2022a). Using this method, researchers almost completely eliminated the wild-type mRNA transcript in bone marrow and blood samples, with over 99% of next-generation sequencing reads from the targeted locus indicating gene editing had occurred (Zhang et al. 2022a). Alternatively, delivery of the gRNA to Cas9-expressing mice can also successfully induce gene editing. These powerful mouse strains provide the ability to perform genome editing in vivo with relative ease and speed, as Cas9-expressing males can be mated with a range of different gRNA-expressing females. It also allows the maintenance of separate colonies where genetically edited mice are only produced when needed. Moreover, it circumvents the problem of trying to maintain and expand colonies where compound or multiple genetic variants make this difficult. Despite concerns within the scientific community, the constitutive expression of Cas9 does not appear to cause

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any health issues in mice (Bond et al. 2021). Such mice exist thanks to the use of the safe-harbour locus, Rosa26, and these mice have so far been used in studies of cancer and immune system dysfunction. One major drawback however is that these organisms are not provided with a donor repair template; therefore any dsDNA breaks at the targeted loci by Cas9 will most likely be repaired by NHEJ, leading to erroneous repairs. This is acceptable if the aim is to disrupt the coding sequence of the targeted gene; however such an approach will not enable specific, tailored edits at that locus via HDR. Consequently, consideration must be given to whether Cas9- or gRNAexpressing mice are suitable for meeting your experimental objectives. Safe-harbour loci have been used in mice by the research community to facilitate overexpression of genes of interest, as a method of studying gene function in development but also to investigate their role in disease onset and progression, e.g. cancer and neurodegenerative diseases. Examples include a mouse strain expressing a Cre-inducible Wnt3a transgene at the Rosa26 locus for study of the Wnt/β-catenin signalling pathway during embryonic development or the overexpression of IL-7Rɑ to elucidate its contribution to the phenotype of T-cell acute lymphoblastic leukaemia and effect upon known cell signalling pathways during tumourigenesis (Chalamalasetty et al. 2016; Silva et al. 2021). Researchers have also made use of this safe-harbour locus to study immunity. Mouse strains harbouring a transgene encoding the diphtheria toxin at the Rosa26 locus have been used to study immune system function and the contribution of cell types to autoimmune conditions such as lupus (Voehringer et al. 2008; Tchen et al. 2022). Mating of the Rosa26 diphtheria toxin mice with specific Cre driver mouse strains ensures expression of the toxin in the desired cell types and causes cell death. The immune system of mice differs somewhat to that of humans, and therefore studying human infections is not always practical in the mouse. There’s also the added issue that some viruses which cause human disease are unable to infect mice. To create mouse models of human viral infections, scientists have integrated the hepatitis B virus genome into the Rosa26 locus which can be excised by Cre recombinase and knocked-in a human cell-surface receptor necessary for cell infection by the virus that causes hand, foot and mouth disease (Kruse et al. 2021; Jin et al. 2021). It is hoped that mouse strains such as these will bring science closer to enabling the detailed study of viral mechanisms of infection and pathogenesis in human diseases, with the added ability to investigate therapeutic approaches. The Rosa26 safeharbour locus has not been spared in the study of neurodegenerative conditions. The overexpression of transgenic met receptor tyrosine kinase from this locus in neural cells in mouse models of amyotrophic lateral sclerosis (ALS) was found to delay the onset of ALS and prolong lifespan (Genestine et al. 2011). In other research, neuronal Rosa26 transgene overexpression of Glo1 is associated with increased anxiety-like behaviour in mice, and injection of this enzyme’s substrate ablated the anxiety (McMurray et al. 2016). These examples demonstrate that in addition to the aforementioned research areas, this genomic safe-harbour locus can be utilised for the study of gene function in the nervous system and neurodegenerative disease.

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Researchers have harnessed the power of the Rosa26 locus to investigate the potential for transgene overexpression to overcome genetic disease. As an example, knock-in of human alpha-1-antitrypsin using CRISPR-Cas9 resulted in high levels of this protein in blood plasma, something which was maintained over a long time period (Stephens et al. 2018). If this can be repeated for other proteins, it will be of interest to those working on protein deficiencies. A mouse model of tyrosinaemia stopped showing symptoms of the disease, and medication was able to be withdrawn, after human fumarylacetoacetate hydrolase (FAH) cDNA was expressed from the Rosa26 safe-harbour locus (Junge et al. 2018). There was no apparent loss of the transgene over the lifespan of the mice in this study, and their corrected phenotype was stable with no adverse effects. It remains to be seen if such an approach can also be applied to other metabolic liver diseases. In some diseases, misexpression of a gene is the principal cause, and attempting to antagonise the effect of overexpression (whether directly or indirectly) may be an avenue for exploration. Tamoxifen-inducible Cre can facilitate expression from a Rosa26 transgene that has the potential to mimic the phenotype of a human disease due to protein overexpression. Such a mouse model of the human disease facioscapulohumeral muscular dystrophy exists, which has allowed for the testing of therapies to overcome the harmful activity of Dux4 derepression, with success (Giesige et al. 2018). Studies like these support the idea that gene therapy using safeharbour loci may be of therapeutic benefit in cases where a disease is caused by a lack of functional gene expression. The mouse is an ideal organism in which to show proof-of-concept for this type of biomedical research, especially as both mice and humans possess the Rosa26 safe-harbour locus and recent improvements in gene editing techniques make it faster and easier to create transgenic mice. As more precise mouse models are created using CRISPR-Cas which accurately recapitulate human genetic diseases, scientists will be able to further investigate whether gene editing using safe-harbour loci holds a promising direction for genetic intervention in the treatment of disease. At the very least, improved testing of therapies using accurate mouse models of disease should be achievable. While genomic safe-harbour loci are highly convenient if you wish to overexpress a construct and you want to know exactly where in the genome it will integrate after delivery. These particular genomic loci are associated with a much higher level of transcription when compared with random integration of expression cassettes, alongside more consistent expression both within the same cell type and over time (Smith et al. 2008). Unfortunately, they are not perfect and do have their pitfalls. As with any genetic editing, care should be taken not to disturb neighbouring genes, although improvements in design and precision including the use of CRISPRCas have made this much simpler over the last decade. It has been shown that chromatin architecture can influence the ability to successfully knock-in a transgene at a safe-harbour locus; those which are located within a more transcriptionally active region of chromatin have a higher success rate, e.g. AAVS1 (van Rensburg et al. 2012). This has been documented as a general observation for any gene editing, not just safe-harbour loci knock-in and in all ZFN, TALEN and CRISPR-Cas methods. There have been reports of methylation of transgenes silencing their

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expression either over time or upon cell differentiation, although the choice of promoter which is inserted as part of the transgene may be able to provide some protection against this, with stronger promoters suggested to be able to overcome that repression (Ordovás et al. 2015; Bhagwan et al. 2019). AAVS1 (also known as PPP1R12C) is a safe-harbour locus within the human genome; however it does not exist within the genome of mice, partially limiting its wide-spread use in biomedical research despite its ease of transgene integration. This intragenic site is well characterised in humans due to its identification as a frequent integration site in cells infected with adeno-associated virus (Samulski et al. 1991; Heister et al. 2002). The CCR5 safe-harbour locus is intragenic and encodes a HIV co-receptor found on the surface of human immune cells; disruption of this genomic locus is associated with resistance to HIV infection but also some increased susceptibility to other viral diseases such as the West Nile Virus. It is located within a less transcriptionally active region of chromatin than AAVS1, resulting in a lower rate of success for knock-ins (van Rensburg et al. 2012). Disruption of the same locus in mice leads to some immune system abnormalities, and it is for these reasons that CCR5 is not considered a true safe-harbour locus in this organism. CCR5 should therefore be avoided when choosing a region of the mouse genome for knock-in of transgenes. It may be that no or only very few select genomic safe-harbour loci truly meet the requirements of their definition in all cell and tissue types or even can be used in the same way across different mammalian species; however Rosa26 is generally accepted to be the best fit across many species at this present time. We have chosen to describe the most common safe-harbour loci used, with a focus on Rosa26; however there are others which have been recently described. These will require further characterisation to justify their suitability if they are to become more widely used in the future. Many of the required DNA constructs for generation of safe-harbour loci knock-ins via CRISPR-Cas9 in mice are available to the scientific community via non-profit repositories, including plasmids for in vitro Cas9 transcription and donor templates for HDR which can be modified to contain your transgene. Well-established protocols are also obtainable from the scientific literature. Alternatively, many commercial reagents are now available from a variety of scientific suppliers for the generation of your own Rosa26 transgenic mice. Timescales for the generation of mouse strains harbouring desired transgenes at the Rosa26 locus are comparable to those created by gene editing using CRISPRCas at other loci. A major benefit to using a genomic safe-harbour locus such as Rosa26 is that this technique can be scaled up and many different transgenic mice can be created using the same Cas9 and gRNAs. The difference from a technical view is the modification of the donor template for HDR, which would need to be modified to suit each transgene independently. Hence, there is a simple experimental workflow which can be harnessed to produce a variety of transgenic mouse strains with relative ease and in a relatively short time frame.

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Uses of CRISPR-Cas in Mice for Non-clinical Purposes

When CRISPR-Cas9 was first utilised in mammalian cells and produced the first genetically edited mice in 2013, it heralded a new era of genome editing research both in vitro and in vivo, quickly becoming a favoured technique now used by many across a variety of scientific disciplines. Within the decade since then, several other Cas enzymes have been characterised, and the original Cas9/crRNA has been altered in many ways to improve characteristics such as cleavage efficiency and reduce the likelihood of off-target effects by enhancing target specificity. To fully understand how useful edited or new members of the CRISPR-Cas toolbox are for genome editing in vivo, there is a requirement for suitable animal reporter strains which can provide such data. Recently, two EGFP mouse strains have been developed precisely for this purpose; these harbour frameshifts in an Egfp transgene expressed from the Rosa26 locus due to deletions generated by Cas9 and NHEJ repair (Miura et al. 2021). Using these mouse strains, researchers can assess their gene editing tools, their methods of delivery and NHEJ/HDR activity using zygotes, isolated populations of primary cells or adult mice. The genome-editing reporter strains mentioned here have advantages over alternative earlier attempts to develop similar reporter mice, due to Egfp being strongly expressed in every tissue from the safeharbour locus (once the frameshift is corrected) and no requirement for fixation protocols that may impact upon a researcher’s ability to visualise EGFP. Furthermore, EGFP is stable, well characterised and does not need specialised equipment outside of what most researchers will have access to in order to detect its presence. Targeting of the Topoisomerase 1 (Top1) gene by CRISPR-Cas9 and subsequent erroneous repair is lethal in pre-implantation mouse embryos. This has been harnessed to produce single-sex litters of pups with an efficiency of 100%, using a mouse strain that constitutively expresses Cas9 from a transgene at the Rosa26 genomic safe-harbour locus (Douglas et al. 2021). Mating males carrying an X chromosome-integrated Top1 gRNA with females homozygous for the Cas9 transgene results in male only litters; however due to concern over the continued expression of the Cas9 endonuclease in offspring, this method was improved further. Mating homozygous females for the Top1 gRNA transgene integrated at the Hipp11 safe-harbour locus, with males carrying a Cas9 insertion on the X chromosome resulted in entirely female litters. Alternatively, mating the same females with males carrying Cas9 on the Y chromosome resulted in entirely female litters. In both mating types, none of the offspring possess the Cas9 transgene as those have been selected against, and therefore it is no longer a factor of concern in downstream procedures. For research projects where only a single sex is necessary and the alternative sex is surplus to requirements, the use of Cas9 in vivo in this manner will enable researchers to reduce the number of mice and matings required, likely also saving valuable time and money. Such approaches are in line with the 3Rs – Replacement, Reduction and Refinement – of animals used in scientific research (Russell and Burch 1960).

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The Mouse for Actively Recording Cells 1 (MARC1) mouse lines have been developed to allow cell lineage tracing in vivo and utilises homing CRISPR guide RNAs (hgRNAs), a modification of CRISPR-Cas9 gRNAs. These mice contain up to 60 germline-transmitted hgRNAs scattered throughout their genomes which were transposed into mES cells (Kalhor et al. 2017, 2018; Leeper et al. 2021). Mating MARC1 mice with a mouse line that constitutively expresses Cas9 (Rosa26-Cas9 knock-in) produces offspring composed of barcoded cells. The design of the hgRNAs is such that in the presence of Cas9, they can be used like gRNAs and the Cas9-hgRNA complexes will result in dsDNA breaks at the integrated hgRNA genomic loci. Subsequent NHEJ activity is likely to change the DNA code of the targeted locus, thus creating genetic variants. Any daughter cells will also possess these variants and accumulate additional variants at other hgRNA loci as Cas9 activity and cell division continues. Over many iterations of the cell cycle, the genomic barcodes within cell populations will deviate and become distinct. The use of next-generation sequencing on barcoded cells allows for the calculation of cell lineage trees, migration and fate. This particular method facilitates Cas9-mediated barcoding on a whole-organism level; however mating MARC1 mice with another line expressing Cas9 under the control of a tissue-specific promoter would allow tissue-specific editing of hgRNA loci in their progeny, if desired. Other methodologies which similarly make use of CRISPR-Cas9 in vivo for the generation of unique genetic variants in different cell lineages also exist, of which many also provide transcriptomic data from barcoded cells e.g. CARLIN (Bowling et al. 2020). Incorporating transcriptomics has the added benefit of being able to identify specific cell types within mixed populations, potentially identifying new ones and giving further details regarding cell lineage. It should however be noted that not all approaches have yet been trialled in mice even though they have been successfully used in zebrafish, e.g. LINNAEUS and scGESTALT (Raj et al. 2018; Spanjaard et al. 2018). Approaches like the ones described here demonstrate how the CRISPRCas toolbox can be applied for a non-clinical purpose yet will deepen our understanding of mammalian development, and knowledge gained from these studies may ultimately lend a hand to fields of research such as regenerative medicine.

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CRISPR-Cas-Editing of Other Mammalian Genomes

An alternative to microinjection of zygotes with Cas9/gRNA/donor templates is to perform the knock-in using isolated somatic cells, usually foetal fibroblasts, and then carry out somatic cell nuclear transfer (SCNT) of the validated genetically edited nucleus into an enucleated oocyte. Pigs and cattle harbouring transgenes at a genomic safe-harbour locus have been successfully generated using the latter method, while microinjection of mouse, rat and rabbit zygotes with constructs destined for safe-harbour loci integration is commonplace (Wang et al. 2017; Xie et al. 2018; Yuan et al. 2021; You et al. 2021). The process is more laborious and

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complicated for pigs and cattle due to the low survival rate of in vitro matured oocytes. Other mammalian species used in scientific research aside from mice have also been genetically engineered at their Rosa26 locus, e.g. a Cre reporter strain of rat which enables the expression of mCherry in cell types where Cre has been expressed alongside a gene of interest; this allows the determination of gene expression patterns in tissues in vivo and well as cell tracing (Ma et al. 2017). Yang et al. (2016) generated a similar transgenic rabbit strain. CRISPR-Cas9 has also been used to generate Cre reporter rabbits via knock-in to the Rosa26 locus in pronuclear stage zygotes, with 35% of full-term kits harbouring the desired knock-in (Yang et al. 2016). Targeting the rat Rosa26 locus in zygotes gave a 29–44% success rate in screened newborns, while in ferret zygotes, an efficiency of 22% has been reported (Ma et al. 2017; Yu et al. 2019). In addition, success has been demonstrated using sheep zygotes, where CRISPR-Cas9 has been shown to successfully knock-in turboGFP into the Rosa26 locus in 12.5% of lambs (Wu et al. 2016). Cas9expressing pigs have also been generated using the Rosa26 locus (Rieblinger et al. 2021). It is envisaged that the use of this locus for the insertion of transgenes in pigs and cattle may lead to benefits for agriculture or biomedicine (Lee et al. 2021). Indeed, one example in cattle is the insertion of NRAMP1, a metal ion transporter into the Rosa26 locus in bovine foetal fibroblasts using CRISPR-Cas9, followed by SCNT; the cattle produced using this method had increased resistance to bovine tuberculosis, a disease which poses a serious threat to agriculture globally as well as public health (Yuan et al. 2021). Reduced cases of bovine tuberculosis in cattle will likely consequently lessen the chance of pathogen transmission to humans. A somewhat similar approach was carried out by Xie et al. (2018), whereby CRISPR-Cas9 was used to insert antiviral shRNAs, targeting classical swine fever virus into the porcine genome at the Rosa26 locus. Classical swine fever virus does not affect human health; however it can often be fatal in pigs and is highly contagious, so is of great concern within the agricultural industry. Insertion of the antiviral shRNAs into the genome of porcine foetal fibroblasts, followed by SCNTgenerated pigs which were resistant to infection with this virus, a trait that was also passed on to their offspring. Genome editing of livestock certainly comes with many ethical and practical considerations; however eliminating the need for vaccine development, vaccination programmes and their associated costs while protecting the health of agricultural animals is certainly a positive. It should be anticipated that CRISPR-Cas technologies will almost certainly be put to use within the agricultural industry, although for what purposes and to what extent that is deemed acceptable remains to be decided.

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References Acosta S, Fiore L, Carota IA, Oliver G (2018) Use of two gRNAs for CRISPR/Cas9 improves bi-allelic homologous recombination efficiency in mouse embryonic stem cells. Genesis 56. https://doi.org/10.1002/DVG.23212 Aida T, Wilde JJ, Yang L et al (2020) Prime editing primarily induces undesired outcomes in mice. bioRxiv:2020.08.06.239723. https://doi.org/10.1101/2020.08.06.239723 Alonso A, Logroscino G, Jick SS, Hernán MA (2009) Incidence and lifetime risk of motor neuron disease in the United Kingdom: a population-based study. Eur J Neurol 16:745–751. https://doi. org/10.1111/J.1468-1331.2009.02586.X Amoasii L, Long C, Li H et al (2017) Single-cut genome editing restores dystrophin expression in a new mouse model of muscular dystrophy. Sci Transl Med:9. https://doi.org/10.1126/ SCITRANSLMED.AAN8081 Bhagwan JR, Denning C, Collins E et al (2019) Variable expression and silencing of CRISPR-Cas9 targeted transgenes identifies the AAVS1 locus as not an entirely safe harbour. F1000Res 8. https://doi.org/10.12688/F1000RESEARCH.19894.2 Böck D, Rothgangl T, Villiger L et al (2022) In vivo prime editing of a metabolic liver disease in mice. Sci Transl Med:14. https://doi.org/10.1126/scitranslmed.abl9238 Bond ST, Zhuang A, Yang C et al (2021) Tissue-specific expression of Cas9 has no impact on whole-body metabolism in four transgenic mouse lines. Mol Metab 53. https://doi.org/10.1016/ J.MOLMET.2021.101292 Bowling S, Sritharan D, Osorio FG et al (2020) An engineered CRISPR/Cas9 mouse line for simultaneous readout of lineage histories and gene expression profiles in single cells. Cell 181: 1410. https://doi.org/10.1016/J.CELL.2020.04.048 Browning J, Rooney M, Hams E et al (2020) Highly efficient CRISPR-targeting of the murine Hipp11 intergenic region supports inducible human transgene expression. Mol Biol Rep 47: 1491–1498. https://doi.org/10.1007/S11033-019-05204-9 Canalis E, Yu J, Schilling L et al (2018) The lateral meningocele syndrome mutation causes marked osteopenia in mice. J Biol Chem 293:14165–14177. https://doi.org/10.1074/JBC.RA118. 004242 Chalamalasetty RB, Ajima R, Garriock RJ et al (2016) A new gain-of-function mouse line to study the role of Wnt3a in development and disease. Genesis 54:497–502. https://doi.org/10.1002/ DVG.22959 Chen H, Shi M, Gilam A et al (2019) Hemophilia A ameliorated in mice by CRISPR-based in vivo genome editing of human factor VIII. Sci Rep 9:1–15. https://doi.org/10.1038/s41598-01953198-y Chu VT, Weber T, Graf R et al (2016) Efficient generation of Rosa26 knock-in mice using CRISPR/ Cas9 in C57BL/6 zygotes. BMC Biotechnol 16:1–15. https://doi.org/10.1186/S12896-0160234-4/TABLES/3 Collisson EA, Campbell JD, Brooks AN et al (2014) Comprehensive molecular profiling of lung adenocarcinoma. Nature 511:543–550. https://doi.org/10.1038/nature13385 Dash PK, Kaminski R, Bella R et al (2019) Sequential LASER ART and CRISPR treatments eliminate HIV-1 in a subset of infected humanized mice. Nat Commun 10:1–20. https://doi.org/ 10.1038/s41467-019-10366-y de los Santos MR, Rivalan M, David FS et al (2021) A CRISPR-Cas9-engineered mouse model for GPI-anchor deficiency mirrors human phenotypes and exhibits hippocampal synaptic dysfunctions. Proc Natl Acad Sci U S A 118:e2014481118. https://doi.org/10.1073/pnas.2014481118 Devoy A, Bunton-Stasyshyn RKA, Tybulewicz VLJ et al (2011) Genomically humanized mice: technologies and promises. Nat Rev Genet 13:14–20. https://doi.org/10.1038/NRG3116 Devoy A, Price G, de Giorgio F et al (2021) Generation and analysis of innovative genomically humanized knockin SOD1, TARDBP (TDP-43), and FUS mouse models. iScience 24:103463. https://doi.org/10.1016/J.ISCI.2021.103463

36

C. J. Monteiro et al.

Douglas C, Maciulyte V, Zohren J et al (2021) CRISPR-Cas9 effectors facilitate generation of single-sex litters and sex-specific phenotypes. Nat Commun 12:1–12. https://doi.org/10.1038/ s41467-021-27227-2 Duan X, Liu J, Zheng X et al (2016) Deficiency of ATP6V1H causes bone loss by inhibiting bone resorption and bone formation through the TGF-β1 pathway. Theranostics 6:2183. https://doi. org/10.7150/THNO.17140 Dumeau CE, Monfort A, Kissling L et al (2019) Introducing gene deletions by mouse zygote electroporation of Cas12a/Cpf1. Transgenic Res 28:525–535. https://doi.org/10.1007/S11248019-00168-9/FIGURES/4 Fujiwara S (2018) Humanized mice: a brief overview on their diverse applications in biomedical research. J Cell Physiol 233:2889–2901. https://doi.org/10.1002/JCP.26022 Gemberling MP, Siklenka K, Rodriguez E et al (2021) Transgenic mice for in vivo epigenome editing with CRISPR-based systems. Nat Methods 18:965. https://doi.org/10.1038/S41592021-01207-2 Genestine M, Caricati E, Fico A et al (2011) Enhanced neuronal met signalling levels in ALS mice delay disease onset. Cell Death Dis 2:e130. https://doi.org/10.1038/CDDIS.2011.11 Giesige CR, Rodino-Klapac LR, Harper SQ (2018) AAV-mediated follistatin gene therapy improves functional outcomes in the TIC-DUX4 mouse model of FSHD. https://doi.org/10. 1172/jci.insight.123538 Gurumurthy CB, O’Brien AR, Quadros RM et al (2019a) Reproducibility of CRISPR-Cas9 methods for generation of conditional mouse alleles: a multi-center evaluation. Genome Biol 20:171. https://doi.org/10.1186/s13059-019-1776-2 Gurumurthy CB, Sato M, Nakamura A et al (2019b) Creation of CRISPR-based germline-genomeengineered mice without ex vivo handling of zygotes by i-GONAD. Nat Protoc 14:2452–2482. https://doi.org/10.1038/S41596-019-0187-X Hasegawa Y, Daitoku Y, Sekiguchi K et al (2013) Novel ROSA26 Cre-reporter knock-in C57BL/ 6N mice exhibiting green emission before and red emission after Cre-mediated recombination. Exp Anim 62:295–304. https://doi.org/10.1538/expanim.62.295 Hashimoto M, Takemoto T (2015) Electroporation enables the efficient mRNA delivery into the mouse zygotes and facilitates CRISPR/Cas9-based genome editing. Sci Rep 5:1–8. https://doi. org/10.1038/srep11315 Heister T, Heid I, Ackermann M, Fraefel C (2002) Herpes simplex virus type 1/adeno-associated virus hybrid vectors mediate site-specific integration at the adeno-associated virus preintegration site, AAVS1, on human chromosome 19. J Virol 76:7163–7173. https://doi.org/10.1128/JVI. 76.14.7163-7173.2002 Hippenmeyer S, Youn YH, Moon HM et al (2010) Genetic mosaic dissection of Lis1 and Ndel1 in neuronal migration. Neuron 68:695–709. https://doi.org/10.1016/j.neuron.2010.09.027 Horn D, Wieczorek D, Metcalfe K et al (2014) Delineation of PIGV mutation spectrum and associated phenotypes in hyperphosphatasia with mental retardation syndrome. Eur J Hum Genet 22:762. https://doi.org/10.1038/EJHG.2013.241 Hosur V, Low BE, Avery C et al (2017) Development of humanized mice in the age of genome editing. J Cell Biochem 118:3043–3048. https://doi.org/10.1002/JCB.26002 Huang X, Zhou G, Wu W et al (2017) Genome editing abrogates angiogenesis in vivo. Nat Commun:8. https://doi.org/10.1038/S41467-017-00140-3 Irion S, Luche H, Gadue P et al (2007) Identification and targeting of the ROSA26 locus in human embryonic stem cells. Nat Biotechnol 25:1477–1482. https://doi.org/10.1038/NBT1362 Jang H, Jo DH, Cho CS et al (2021) Application of prime editing to the correction of mutations and phenotypes in adult mice with liver and eye diseases. Nat Biomed Eng 6(26):181–194. https:// doi.org/10.1038/s41551-021-00788-9 Jin LF, Li JS (2016) Generation of genetically modified mice using CRISPR/Cas9 and haploid embryonic stem cell systems. Dongwuxue Yanjiu 37:205–213. https://doi.org/10.13918/J. ISSN.2095-8137.2016.4.205

2

Modern Approaches to Mouse Genome Editing Using the CRISPR-Cas Toolbox. . .

37

Jin Y, Sun T, Zhou G et al (2021) Pathogenesis study of enterovirus 71 using a novel human SCARB2 Knock-in mouse model. mSphere 6. https://doi.org/10.1128/MSPHERE.01048-20 Junge N, Yuan Q, Vu TH et al (2018) Homologous recombination mediates stable Fah gene integration and phenotypic correction in tyrosinaemia mouse-model. World J Hepatol 10: 277–286. https://doi.org/10.4254/WJH.V10.I2.277 Ka Y, Katano I, Nishinaka E et al (2021) Improved engraftment of human peripheral blood mononuclear cells in NOG MHC double knockout mice generated using CRISPR/Cas9. Immunol Lett 229:55–61. https://doi.org/10.1016/J.IMLET.2020.11.011 Kalhor R, Mali P, Church GM (2017) Rapidly evolving homing CRISPR barcodes. Nat Methods 14:195–200. https://doi.org/10.1038/nmeth.4108 Kalhor R, Kalhor K, Mejia L et al (2018) Developmental barcoding of whole mouse via homing CRISPR. Science 361. https://doi.org/10.1126/SCIENCE.AAT9804 Kim K, Park SW, Kim JH et al (2017a) Genome surgery using Cas9 ribonucleoproteins for the treatment of age-related macular degeneration. Genome Res 27:419–426. https://doi.org/10. 1101/GR.219089.116/-/DC1 Kim K, Ryu SM, Kim ST et al (2017b) Highly efficient RNA-guided base editing in mouse embryos. Nat Biotechnol 35:435–437. https://doi.org/10.1038/NBT.3816 Kim YS, Kim GR, Park M et al (2020) Electroporation of AsCpf1/RNP at the zygote stage is an efficient genome editing method to generate Knock-out mice deficient in leukemia inhibitory factor. Tissue Eng Regen Med 17:45. https://doi.org/10.1007/S13770-019-00225-8 Kruse RL, Barzi M, Legras X et al (2021) A hepatitis B virus transgenic mouse model with a conditional, recombinant, episomal genome. JHEP Report 3:100252. https://doi.org/10.1016/J. JHEPR.2021.100252 Lee B, Lee K, Panda S et al (2018) Nanoparticle delivery of CRISPR into the brain rescues a mouse model of fragile X syndrome from exaggerated repetitive behaviours. Nat Biomed Eng 2:497. https://doi.org/10.1038/S41551-018-0252-8 Lee C, Yum S-Y, Choi W et al (2021) Identification of the safe harbor locus, AAVS1, from porcine genome and site-specific integration of recombinase-mediated cassette exchange system in porcine fibroblasts using CRISPR/Cas9. bioRxiv:2021.09.15.460532. https://doi.org/10.1101/ 2021.09.15.460532 Leeper K, Kalhor K, Vernet A et al (2021) Lineage barcoding in mouse with homing CRISPR. Nat Protoc 16:2088. https://doi.org/10.1038/S41596-020-00485-Y Li YS, Meng RR, Chen X et al (2019) Generation of H11-albumin-rtTA transgenic mice: a tool for inducible gene expression in the liver. G3: Genes|Genomes|Genetics 9:591. https://doi.org/10. 1534/G3.118.200963 Liao HK, Hatanaka F, Araoka T et al (2017) In vivo target gene activation via CRISPR/Cas9mediated trans-epigenetic modulation. Cell 171:1495. https://doi.org/10.1016/J.CELL.2017. 10.025 Liu Z, Lu Z, Yang G et al (2018) Efficient generation of mouse models of human diseases via ABEand BE-mediated base editing. Nat Commun 9(19):1–8. https://doi.org/10.1038/s41467-01804768-7 Liu Y, Li X, He S et al (2020) Efficient generation of mouse models with the prime editing system. Cell Discov 6:1–6. https://doi.org/10.1038/s41421-020-0165-z Ma Y, Yu L, Pan S et al (2017) CRISPR/Cas9-mediated targeting of the Rosa26 locus produces Cre reporter rat strains for monitoring Cre–loxP-mediated lineage tracing. FEBS J 284:3262–3277. https://doi.org/10.1111/FEBS.14188 Maruyama T, Dougan SK, Truttmann MC et al (2015) Inhibition of non-homologous end joining increases the efficiency of CRISPR/Cas9-mediated precise [TM: inserted] genome editing. Nat Biotechnol 33:538. https://doi.org/10.1038/NBT.3190 McMurray KMJ, Du X, Brownlee M, Palmer AA (2016) Neuronal overexpression of Glo1 or amygdalar microinjection of methylglyoxal is sufficient to regulate anxiety-like behavior in mice. Behav Brain Res 301:119–123. https://doi.org/10.1016/J.BBR.2015.12.026

38

C. J. Monteiro et al.

Min YL, Chemello F, Li H et al (2020) Correction of three prominent mutations in mouse and human models of Duchenne muscular dystrophy by single-cut genome editing. Mol Ther 28: 2044. https://doi.org/10.1016/J.YMTHE.2020.05.024 Miura H, Gurumurthy CB, Sato T et al (2015) CRISPR/Cas9-based generation of knockdown mice by intronic insertion of artificial microRNA using longer single-stranded DNA. Sci Rep 5:1–11. https://doi.org/10.1038/srep12799 Miura H, Imafuku J, Kurosaki A et al (2021) Novel reporter mouse models useful for evaluating in vivo gene editing and for optimization of methods of delivering genome editing tools. Mol Ther Nucleic Acids 24:325–336. https://doi.org/10.1016/j.omtn.2021.03.003 Mochizuki Y, Chiba T, Kataoka K et al (2018) Combinatorial CRISPR/Cas9-approach to elucidate a far-upstream enhancer complex for tissue-specific Sox9 expression. Dev Cell 46:794. https:// doi.org/10.1016/J.DEVCEL.2018.07.024 Monteys AM, Ebanks SA, Keiser MS, Davidson BL (2017) CRISPR/Cas9 editing of the mutant huntingtin allele in vitro and in vivo. Mol Ther 25:12. https://doi.org/10.1016/J.YMTHE.2016. 11.010 Nelson CE, Hakim CH, Ousterout DG et al (2016) In vivo genome editing improves muscle function in a mouse model of Duchenne muscular dystrophy. Science 351:403–407. https:// doi.org/10.1126/SCIENCE.AAD5143 Nopoulos PC (2016) Huntington disease: a single-gene degenerative disorder of the striatum. Dialogues Clin Neurosci 18:91. https://doi.org/10.31887/DCNS.2016.18.1/PNOPOULOS Ohmori T, Nagao Y, Mizukami H et al (2017) CRISPR/Cas9-mediated genome editing via postnatal administration of AAV vector cures haemophilia B mice. Sci Rep 7(17):1–11. https://doi.org/10.1038/s41598-017-04625-5 Ordovás L, Boon R, Pistoni M et al (2015) Efficient recombinase-mediated cassette exchange in hPSCs to study the hepatocyte lineage reveals AAVS1 locus-mediated transgene inhibition. Stem Cell Report 5:918–931. https://doi.org/10.1016/j.stemcr.2015.09.004 Park H, Oh J, Shim G et al (2019) In vivo neuronal gene editing via CRISPR-Cas9 amphiphilic nanocomplexes alleviates deficits in mouse models of Alzheimer’s disease. Nat Neurosci 22: 524–528. https://doi.org/10.1038/S41593-019-0352-0 Pi M, Xu F, Ye R et al (2020) Humanized GPRC6AKGKY is a gain-of-function polymorphism in mice. Sci Rep 10:1–10. https://doi.org/10.1038/s41598-020-68113-z Platt RJ, Chen S, Zhou Y et al (2014) CRISPR-Cas9 Knockin mice for genome editing and cancer modeling. Cell 159:440. https://doi.org/10.1016/J.CELL.2014.09.014 Raj B, Wagner DE, McKenna A et al (2018) Simultaneous single-cell profiling of lineages and cell types in the vertebrate brain. Nat Biotechnol 36:442. https://doi.org/10.1038/NBT.4103 Rauch F, Geng Y, Lamplugh L et al (2018) Crispr-Cas9 engineered osteogenesis imperfecta type V leads to severe skeletal deformities and perinatal lethality in mice. Bone 107:131–142. https:// doi.org/10.1016/J.BONE.2017.11.013 Rieblinger B, Sid H, Duda D et al (2021) Cas9-expressing chickens and pigs as resources for genome editing in livestock. Proc Natl Acad Sci 118:e2022562118. https://doi.org/10.1073/ pnas.2022562118 Roh JI, Lee J, Park SU et al (2018) CRISPR-Cas9-mediated generation of obese and diabetic mouse models. Exp Anim 67:229. https://doi.org/10.1538/EXPANIM.17-0123 Rovira M, Maestro MA, Grau V, Ferrer J (2021) Hnf1b-CreER causes efficient recombination of a Rosa26-RFP reporter in duct and islet δ cells. Islets 13:134. https://doi.org/10.1080/19382014. 2021.1955088 Ruan J, Li H, Xu K et al (2015) Highly efficient CRISPR/Cas9-mediated transgene knockin at the H11 locus in pigs. Sci Rep 5(15):1–10. https://doi.org/10.1038/srep14253 Russell WMS, Burch RL (1960) The principles of humane experimental technique. Med J Aust 1: 500–500. https://doi.org/10.5694/J.1326-5377.1960.TB73127.X Ryu SM, Koo T, Kim K et al (2018) Adenine base editing in mouse embryos and an adult mouse model of Duchenne muscular dystrophy. Nat Biotechnol 36:536–539. https://doi.org/10.1038/ NBT.4148

2

Modern Approaches to Mouse Genome Editing Using the CRISPR-Cas Toolbox. . .

39

Samulski RJ, Zhu X, Xiao X et al (1991) Targeted integration of adeno-associated virus (AAV) into human chromosome 19. EMBO J 10:3941–3950. https://doi.org/10.1002/J.1460-2075.1991. TB04964.X Scholefield J, Harrison PT (2021) Prime editing – an update on the field. Gene Ther 28:396–401. https://doi.org/10.1038/s41434-021-00263-9 Silva A, Almeida ARM, Cachucho A et al (2021) Overexpression of wild-type IL-7Rα promotes T-cell acute lymphoblastic leukemia/lymphoma. Blood 138:1040–1052. https://doi.org/10. 1182/BLOOD.2019000553 Singh P, Schimenti JC, Bolcun-Filas E (2015) A mouse geneticist’s practical guide to CRISPR applications. Genetics 199:1–15. https://doi.org/10.1534/GENETICS.114.169771 Smith JR, Maguire S, Davis LA et al (2008) Robust, persistent transgene expression in human embryonic stem cells is achieved with AAVS1-targeted integration. Stem Cells 26:496–504. https://doi.org/10.1634/STEMCELLS.2007-0039 Song CQ, Jiang T, Richter M et al (2020) Adenine base editing in an adult mouse model of tyrosinemia. Nat Biomed Eng 4:125. https://doi.org/10.1038/S41551-019-0357-8 Spanjaard B, Hu B, Mitic N et al (2018) Simultaneous lineage tracing and cell-type identification using CRISPR/Cas9-induced genetic scars. Nat Biotechnol 36:469. https://doi.org/10.1038/ NBT.4124 Stephens CJ, Kashentseva E, Everett W et al (2018) Targeted in vivo knock-in of human alpha-1antitrypsin cDNA using adenoviral delivery of CRISPR/Cas9. Gene Ther 25:139–156. https:// doi.org/10.1038/S41434-018-0003-1 Tabebordbar M, Zhu K, Cheng JKW et al (2016) In vivo gene editing in dystrophic mouse muscle and muscle stem cells. Science 351:407. https://doi.org/10.1126/SCIENCE.AAD5177 Tasic B, Hippenmeyer S, Wang C et al (2011) Site-specific integrase-mediated transgenesis in mice via pronuclear injection. Proc Natl Acad Sci U S A 108:7902–7907. https://doi.org/10.1073/ pnas.1019507108 Tchen J, Simon Q, Chapart L et al (2022) CT-M8 mice: a new mouse model demonstrates that basophils have a nonredundant role in lupus-like disease development. Front Immunol 13:1. https://doi.org/10.3389/FIMMU.2022.900532 Thakore PI, Kwon JB, Nelson CE et al (2018) RNA-guided transcriptional silencing in vivo with S. aureus CRISPR-Cas9 repressors. Nat Commun 9:1–9. https://doi.org/10.1038/s41467-01804048-4 Tröder SE, Ebert LK, Butt L et al (2018) An optimized electroporation approach for efficient CRISPR/Cas9 genome editing in murine zygotes. PloS One 13:e0196891. https://doi.org/10. 1371/JOURNAL.PONE.0196891 van Rensburg R, Beyer I, Yao XY et al (2012) Chromatin structure of two genomic sites for targeted transgene integration in induced pluripotent stem cells and hematopoietic stem cells. Gene Ther 20:201–214. https://doi.org/10.1038/gt.2012.25 Voehringer D, Liang H-E, Locksley RM (2008) Homeostasis and effector function of lymphopeniainduced ‘memory-like’ T cells in constitutively T cell-depleted mice. J Immunol 180:4742. https://doi.org/10.4049/JIMMUNOL.180.7.4742 Wang H, Yang H, Shivalila CS et al (2013) One-step generation of mice carrying mutations in multiple genes by CRISPR/Cas-mediated genome engineering. Cell 153:910. https://doi.org/10. 1016/J.CELL.2013.04.025 Wang K, Jin Q, Ruan D et al (2017) Cre-dependent Cas9-expressing pigs enable efficient in vivo genome editing. Genome Res 27:2061–2071. https://doi.org/10.1101/GR.222521.117/-/DC1 Wu M, Wei C, Lian Z et al (2016) Rosa26-targeted sheep gene knock-in via CRISPR-Cas9 system. Sci Rep 6:1–7. https://doi.org/10.1038/srep24360 Xie Z, Pang D, Yuan H et al (2018) Genetically modified pigs are protected from classical swine fever virus. PLoS Pathog:14. https://doi.org/10.1371/JOURNAL.PPAT.1007193 Yang H, Wang H, Shivalila CS et al (2013) One-step generation of mice carrying reporter and conditional alleles by CRISPR/Cas-mediated genome engineering. Cell 154:1370. https://doi. org/10.1016/J.CELL.2013.08.022

40

C. J. Monteiro et al.

Yang D, Song J, Zhang J, et al (2016) Identification and characterization of rabbit ROSA26 for gene knock-in and stable reporter gene expression. Sci Rep 2016 6:1–8. doi:https://doi.org/10.1038/ srep25161 Yin H, Song CQ, Dorkin JR et al (2016) Therapeutic genome editing by combined viral and non-viral delivery of CRISPR system components in vivo. Nat Biotechnol 34:328. https://doi. org/10.1038/NBT.3471 You W, Li M, Qi Y et al (2021) CRISPR/Cas9-mediated specific integration of Fat-1 and IGF-1 at the pRosa26 locus. Genes 12:1027. https://doi.org/10.3390/GENES12071027 Yu M, Sun X, Tyler SR et al (2019) Highly efficient Transgenesis in ferrets using CRISPR/Cas9mediated homology-independent insertion at the ROSA26 locus. Sci Rep 9:1–13. https://doi. org/10.1038/s41598-018-37192-4 Yuan M, Zhang J, Gao Y et al (2021) HMEJ-based safe-harbor genome editing enables efficient generation of cattle with increased resistance to tuberculosis. J Biol Chem 296:100497. https:// doi.org/10.1016/j.jbc.2021.100497 Zambrowicz BP, Imamoto A, Fiering S et al (1997) Disruption of overlapping transcripts in the ROSA βgeo 26 gene trap strain leads to widespread expression of β-galactosidase in mouse embryos and hematopoietic cells. Proc Natl Acad Sci 94:3789–3794. https://doi.org/10.1073/ pnas.94.8.3789 Zhang L, Li W, Liu Z et al (2022a) sgRNA knock-in mouse provides an alternative approach for in vivo genetic modification. Front Cell Dev Biol 9:3869. https://doi.org/10.3389/fcell.2021. 769673 Zhang Y, Li H, Nishiyama T et al (2022b) A humanized knockin mouse model of Duchenne muscular dystrophy and its correction by CRISPR-Cas9 therapeutic gene editing. Mol Ther Nucleic Acids 29:525. https://doi.org/10.1016/J.OMTN.2022.07.024 Zhu F, Gamboa M, Farruggio AP et al (2014) DICE, an efficient system for iterative genomic editing in human pluripotent stem cells. Nucleic Acids Res 42:e34–e34. https://doi.org/10.1093/ NAR/GKT1290 Zhu L, Yang X, Li J et al (2021) Leptin gene-targeted editing in ob/ob mouse adipose tissue based on the CRISPR/Cas9 system. J Genet Genomics 48:134–146. https://doi.org/10.1016/J.JGG. 2021.01.008 Zuckermann M, Hovestadt V, Knobbe-Thomsen CB et al (2015) Somatic CRISPR/Cas9-mediated tumour suppressor disruption enables versatile brain tumour modelling. Nat Commun 6. https:// doi.org/10.1038/NCOMMS8391

Chapter 3

Long Non-coding RNAs and CRISPR-Cas Edition in Tumorigenesis Cristiana Libardi Miranda Furtado, Renan da Silva Santos, Sarah Leyenne Alves Sales, Louhana Pinheiro Rodrigues Teixeira, and Claudia do Ó Pessoa

1 Long Non-coding RNAs: Biogenesis and Mechanisms of Action Most of the mammalian genome is comprised of DNA elements that are transcribed into non-coding RNAs (ncRNAs) that control gene expression in a cell-specific manner (Yao et al. 2019). Among these non-protein coding transcripts, long ncRNAs (lncRNAs) are a class of transcripts greater than 200 nucleotides (Yao et al. 2019), which are the most common ncRNAs in the human genome, and account for 30.87% of the total number of genes (61,852), with an estimate of 19,095 lncRNA genes, almost similar to protein coding genes (19,370–31.31%) (GENCODE 2022) (https://www.gencodegenes.org, accessed on 30/09/2022). The lncRNAs can be transcribed by different DNA elements, as primary gene transcripts, overlapping protein-coding gene transcripts, as well as upstream promoter regions, enhancers and intergenic regions, with different RNA processing mechanisms and functions in the cell fate (Wu et al. 2017). The majority of lncRNAs are transcribed by RNA polymerase II although their biogenesis is complex and directly related to their roles in cell function. Like proteinC. L. M. Furtado (✉) University of Fortaleza, Experimental Biology Center, Fortaleza, Ceara, Brazil Drug Research and Development Center, Postgraduate Program in Translational Medicine, Federal University of Ceara, Fortaleza, Brazil e-mail: [email protected] R. da Silva Santos · S. L. A. Sales · C. Ó. Pessoa Department of Physiology and Pharmacology, Drug Research and Development Center, Federal University of Ceara, Fortaleza, Brazil L. P. R. Teixeira University of Fortaleza, Experimental Biology Center, Fortaleza, Ceara, Brazil © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_3

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coding messenger RNAs (mRNAs), some lncRNAs can be processed by the addition of a methyl group on the guanosine (7-metil-guanosine, m7G) at their 5′-end (capping) and polyadenylation at their 3′-end (poly(A) tail). However, unlike mRNAs, in most lncRNAs, the removal of introns at splicing is not a well-ordered process, as the splicing regulator’s signals are not efficient (Statello et al. 2021). These lncRNAs usually are transcribed by dysregulated phosphorylation at the C-terminal domain (CTD) of Pol II. In the absence of post-transcriptional modifications initiated by the co-transcription splicing, capping and polyA, and nuclear export signals to the cytoplasm, many lncRNAs remain in the nucleus linked to chromatin (Wu et al. 2017; Statello et al. 2021). For other lncRNAs, the processing and stabilization can be achieved by other mechanisms, including RNase P cleavage, capping in one (5′) or both ends by small nucleolar ribonucleoproteins (snoRNPs), and by forming protective circular structures (Statello et al. 2021; Salzman et al. 2012; Yin et al. 2012). LncRNAs participate in an intricate network of interaction, targeting important molecular markers transcriptionally and post-transcriptionally, with important roles in several biological processes during cell growth and differentiation. The function of lncRNAs is still unclear given the complex genomic architecture of lncRNA locus, being distributed through the genome in intergenic, antisense, and intronic, which may lack identified regulatory elements, such as promoter regions, transcription start sites, and enhancers (Batagov et al. 2013). Also, intergenic, cis-antisense, and intronic lncRNAs have increased genomic instability given the short half-life time (80%) comprises papillary thyroid cancer (PTC), which is considered a differentiated type of cancer (i.e., one that maintains thyroid cell function) and is often curable with standard care, involving surgery and radioiodine treatment (Haugen et al. 2016). The second most common type is follicular thyroid carcinoma (FTC), comprising ~15% of cases. However, there is a rare form of thyroid cancer (~2–3% of cases), the anaplastic thyroid cancer (ATC), that is very aggressive and

C. S. Fuziwara · E. T. Kimura (✉) Department of Cell and Developmental Biology, Institute of Biomedical Sciences, University of São Paulo, São Paulo, SP, Brazil e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_5

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lethal, for which clinical care is complicated (Bible et al. 2021). Therefore, searching for new approaches to treat this disease is imperative. To do so, we need to understand better the molecular mechanisms involved in cancer cell aggressiveness in vitro using, for example, gene editing technologies, such as CRISPR/Cas9. CRISPR/Cas9 programmed gene editing emerged as a potent tool to manipulate a genome sequence without demanding time-consuming bioengineering as the TALEN and ZNFs (Jinek et al. 2012). The possibility of permanent modification of the genome in vitro leads to a better understanding of a gene’s role in cancer cell biology and makes possible a potential therapeutic application. In this context, thyroid cancer cell lines have been extensively used to develop basic cancer research (Schweppe et al. 2008; Zhao et al. 2011) and as a tool for drug validation in vitro and in animal models. Nevertheless, Schweppe et al. raised concerns regarding the cell line identity. They reported that several human thyroid cancer cell lines were crosscontaminated with other cancer types (Schweppe et al. 2008), leading to a shared effort to authenticate the uniqueness of thyroid cancer cell lines using short tandem repeat (STR) profile sequencing. Apart from that issue, currently, our field has access to more than twenty unique thyroid cancer cell lines (Table 5.1), including the papillary, follicular, and anaplastic histotypes (Zhao et al. 2011; Schweppe et al. 2008). In this chapter, we will explore recent advances in thyroid cancer research using CRISPR/Cas9 methodology as a tool to understand thyroid cancer biology in vitro (Fig. 5.1).

Table 5.1 Unique human thyroid cancer cell lines authenticated by STR profiling Histotype Papillary thyroid cancer (PTC) Follicular thyroid cancer (FTC) Anaplastic thyroid cancer (ATC)

Cell lines TPC1, BCPAP, K1, KTC1 TT2609-CO2, FTC133, ML1, WRO82-1 8505C, SW1736, Cal-62, T235, T238, Uhth-104, ACT-1, HTh74, KAT18, TTA1, FRO81-2, HTh7, C643, BHT101

References Schweppe et al. (2008) and Zhao et al. (2011) Schweppe et al. (2008) and Zhao et al. (2011) Schweppe et al., (2008) and Zhao et al. (2011)

Fig. 5.1 Applications of CRISPR/Cas9 to target coding and noncoding genes in the investigation of thyroid cancer biology

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2 Targeting MAPK Thyroid Cancer Genes It is well established that the oncogenesis of thyroid cancer is driven by MAPK signaling. Since the first descriptions of RAS mutations (Lemoine et al. 1989), the RET/PTC rearrangements (Grieco et al. 1990), and the BRAFV600E mutation (Kimura et al. 2003), cumulative evidence has indicated that thyroid cancer is essentially an MAPK disease (Fagin and Wells Jr. 2016). The most frequent genetic alteration in PTC is BRAFV600E mutation followed by nonoverlapping RAS mutations and RET/PTC rearrangements, which altogether account more than 80% of the cases (Kimura et al. 2003). Although researchers have studied thyroid cancer oncogenesis and genetics for three decades, there are several gaps in the understanding of the molecular pathway of cancer progression. Recent advances in high-throughput sequencing methodology unveiled a panel of additional genetic alterations in aggressive-thyroid-cancer patients, such as alterations in PI3K signaling, protein synthesis components, chromatin modifiers, and other genes (Landa et al. 2016; Ricarte-Filho et al. 2009). Nevertheless, functional validation of these alterations’ role in aggressive thyroid cancer biology and the potential for targeted therapy is still under investigation. Thyroid follicular cell lines have been useful in elucidating the role of MAPKsignaling-associated oncogenes. Seminal studies have shown that activation of thyroid oncogenes, such as RASG12D (Saavedra et al. 2000), BRAFV600E (Mitsutake et al. 2005), and RET/PTC3 (Wang et al. 2003), induces cell proliferation and loss of thyroid differentiation in normal rat thyroid follicular cell PCCl3. These data were further corroborated with the establishment of transgenic mouse models with targeted expression of BRAFV600E, RET/PTC1-3, and RAS in thyroid follicular cells that developed thyroid cancer (Knauf et al. 2005; Jhiang et al. 1996; Vitagliano et al. 2006). Murine mouse models are a potent tool for in vivo studies (Kirschner et al. 2016); however, the use of human-derived thyroid cancer cell lines prevails as the most used tool in thyroid cancer studies. Blocking MAPK signaling with multi-kinase inhibitors (sorafenib and lenvatinib), MEK inhibitors (selumetinib and trametinib), or mutant BRAF inhibitor (vemurafenib and dabrafenib) is currently indicated as a therapy for advanced thyroid cancer patients (Shonka Jr. et al. 2022; Cabanillas et al. 2019). Indeed, PTC and ATC patients may already benefit from a combination of target therapies (Brose et al. 2016; Wang et al. 2019). Therefore, drug testing in thyroid cancer cell lines is imperative to evaluate cell response and understand the molecular mechanisms of resistance. For example, in vitro testing of the BRAFV600E inhibitor vemurafenib showed that thyroid cancer cells with BRAFV600E acquire resistance to this drug by induction of the HER3 gene and reactivation of MAPK signaling (Montero-Conde et al. 2013). In a transgenic mouse model, BRAF-mutated ATC lines were resistant to BRAF inhibitors, and recurrent tumors usually developed amplifications in the Met gene that reactivated MAPK signaling (Knauf et al. 2018). With all these background reports, in recent studies, researchers have tested the combinatory treatment of dabrafenib and trametinib, which blocks mutant BRAF

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and MEK, respectively, with promising effects in ATC patients, in some cases leading to complete resection of the tumor (Wang et al. 2019; Subbiah et al. 2018). The combinatorial effect of BRAF inhibitors and immunotherapy (i.e., blockage of PD1 with pembrolizumab) also promoted positive effects in ATC treatment (Cabanillas et al. 2018). Nevertheless, the thyroid cancer main oncogenic drivers RET, RAS, and BRAF genes have not yet been successfully targeted with CRISPR/Cas9 gene editing. Below, we pinpoint some studies in which researchers have employed CRISPR/ Cas9 to disrupt MAPK pathway components. Targeting EGFR MAPK signaling may also be activated by the ligation of epidermal growth factor (EGF) to its receptor EGFR, and in several types of cancer, such as lung cancer, EGFR mutations are responsible for oncogenesis (Uribe et al. 2021). However, no mutation is detected in ATC, but EGFR is overexpressed (Schiff et al. 2004). The temporary blockage of the EGFR function with the tyrosine kinase inhibitor gefitinib resulted in antitumoral effects, such as induction of apoptosis as well as reduction of cell proliferation and tumor growth in xenotransplant tumors in mice (Schiff et al. 2004). The use of CRISPR/Cas9 to blunt EGFR gene expression permanently is important to investigate the role of EGF-mediated signaling in thyroid cancer. Targeting EGFR with specific sgRNA in a lentiviral particle that also carries the Cas9 gene resulted in efficient reduction of EGFR protein levels and blocked MAPK/ERK and AKT signaling phosphorylation in ATC cell lines (Huang et al. 2018). Moreover, EGFR gene editing induced antitumoral effects such as impairment of colony formation, cell growth arrest, and reduction of EMT gene expression in vitro. Targeting NF1 and NF2 MAPK signaling is negatively regulated by the tumor suppressor genes NF1 and NF2, which block RAS activation. The analysis of largescale genetic data in RAS-mutant PTC and poorly differentiated thyroid cancer (PDTC) shows ch22q loss of heterozygosity where the NF2 gene is located (Garcia-Rendueles et al. 2015). In thyroid cancer cell lines, a low incidence of NF2 mutations is observed, but NF2 was repressed in some lines. Therefore, the overexpression of NF2 in NF2-null cell lines inhibited cell growth and attenuated Ras signaling. On the other hand, NF2 knock-down resulted in induction of RAS expression (Garcia-Rendueles et al. 2015). With the use of CRISPR/Cas9 to blunt the NF2 gene in BRAFV600E thyroid cancer cell lines, no effect in cell growth was observed (You et al. 2019), probably due to the prominent role of NF2 only in Ras-mutated lines. NF1 also plays a role in Ras-mutated thyroid tumors. Using a murine model that develops PDTC (Hras- (Tpo-Cre/HrasG12V/p53flox/flox)), researchers showed that mice treated with tipifarnib for a long period develop resistance due to inactivation of the Nf1 gene (Untch et al. 2018). Indeed, Nf1 gene editing with CRISPR/Cas9 in a murine PDTC cell line resulted in a 5-fold increase in the resistance to tipifarnib. Moreover, a similar effect was observed when NF1 was knocked down in RAS-mutant human ATC cell line, leading to the development of resistance to tipifarnib.

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3 Targeting Noncoding RNA Genes (microRNA and lncRNA) The completion of human euchromatic genome sequencing in 2003 brought to our attention that only a small proportion of human DNA encodes proteins (with an estimated ~20,000–25,000 protein-coding genes at that time) (International Human Genome Sequencing 2004). However, in 2022 the telomere-to-telomere (T2T) Consortium revealed only 19,969 protein coding genes and closed the gaps related to heterochromatic-region sequencing (Nurk et al. 2022). Most of our genome comprises noncoding DNA, which encompasses regulatory sequences for gene expression as cis-regulatory elements (cCRE) and another more than 50,000 genes for noncoding RNAs. In this context, the Encyclopedia of DNA Elements (ENCODE) is a consortium project that aims to map and decipher the functional elements in the human genome (The ENCODE Project Consortium 2012). microRNAs (miRNAs) are a class of small noncoding RNAs ~18nt long that regulate gene expression post-transcriptionally (Gebert and MacRae 2019). On the other hand, long noncoding RNAs are a class of long RNAs (>200nt in length) that regulate gene expression in several ways, including by interacting with miRNAs (Statello et al. 2021). Currently, the miRNA database miRBase release 22.1 contains 1973 human miRNA-annotated genes (Kozomara et al. 2019). For long noncoding RNAs, the number of annotated genes is far superior with 49,372 genes in the fifth release of the LNCipedia database (Volders et al. 2019). The miRNA gene is transcribed into long primary miRNAs (pri-miRNA) that contains a 5′Cap and poli-A tail, as well as a region of a hairpin secondary structure that is recognized by the microprocessor complex (DROSHA+DGCR8) (Ha and Kim 2014). This complex excises the miRNA precursor (pre-miRNA), which is an ~80-nt RNA folded into a hairpin that Exportin-5 exports to the cytoplasm. In the cytoplasm, DICER endonuclease processes the precursor to form a miRNA/miRNA duplex. Then, only one strand of this duplex is retained and loaded into an AGO protein to form the RNA-induced silencing complex (RISC), which directs the posttranscriptional effect in the 3′-UTR of target mRNA (Ha and Kim 2014). The mechanism of action is associated with the blockage of mRNA translation and the destabilization of mRNA by targeting poli-A tail and 5′ Cap regions, leading to reduction of protein levels (Gebert and MacRae 2019). It is estimated that miRNAs may regulate more than 60% of human coding genes by interacting with the 3′-UTR of mRNAs (Friedman et al. 2009). Indeed, when looking for potential targets of a single miRNA, such as let-7 family, we find 1207 potential mRNA targets in the human TargetScan database (McGeary et al. 2019; Lewis et al. 2003). As miRNA target mRNAs may be protooncogenes and tumor suppressor genes, any dysregulation in the physiological expression within a cell line may lead to oncogenic effects (Esquela-Kerscher and Slack 2006). For example, the overexpression of the miR-17-92 cluster is oncogenic in lymphoma because it reduces PTEN (He et al. 2005; Olive et al. 2009), and down-regulation of let-7 is oncogenic

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in lung cancer because it de-represses RAS oncogene (Johnson et al. 2005; Takamizawa et al. 2004). Our group has focused on investigating miRNAs’ role in thyroid cancer biology (Fuziwara and Kimura 2017). In this context, modulating miRNAs that are overexpressed in thyroid cancer would affect thyroid cancer treatment. Among these miRNAs, miR-146b-5p and miR-17-92 are associated with a worst thyroid cancer prognosis. miR-146b is one of the most highly expressed miRNAs in thyroid cancer, and it has a positive correlation with poor prognosis characteristics of the disease, such as the presence of extrathyroidal invasion, tumor recurrence, and distant metastasis in patients (Chou et al. 2010; Yip et al. 2011). Several studies have shown that the transient blockage of miR-146b function with antimiRs exerts antitumoral effects in thyroid cancer cells and in vivo and de-represses miR-146b-5p target genes, such as SMAD4, PTEN, and E-cadherin (Geraldo et al. 2012; Lima et al. 2016; RamirezMoya et al. 2018). AntimiRs are antisense oligonucleotides that contain LNA modifications that promote a stable interaction with miRNAs and block its function. However, a permanent modulation (blockage) is difficult to achieve using antimiR (antagomiRs) methodology. Therefore, in gene editing, CRISPR/Cas9 emerged as a potential tool to delete a miRNA segment in the genome and promote a permanent down-regulation of miRNAs. The MIR146B gene is located in chromosome 10 (10q24.32) and is surrounded by the CUEDC2 gene in the 5′ region and C10orf95-AS1 in the 3′ region. To blunt (disrupt) permanently the expression of miR-146b-5p in anaplastic thyroid cancer, we searched for PAM sequences (NGG) that flanked the precursor sequence of mir-146b (Santa-Inez et al. 2021), targeting this region with Cas9n. The use of Cas9n (mutant nickase) is associated with a reduced off-targeting possibility and employs a double-sgRNA strategy that requires targeting of opposite strands of DNA at a defined distance from each other (Ran et al. 2013). As expected, the CRISPR/Cas9n promoted deletions in the precursor loop within the double-sgRNAtargeted region of MIR146B gene and resulted in altered miRNA processing and a >80% reduction of mature miR-146b-5p levels in edited clones of KTC2 cell line (Santa-Inez et al. 2021). Functionally, this reduction of miR-146b-5p led to antitumoral effects in the ATC cell line, such as reduction of cell viability, cell migration, and colony formation and also tumor growth in vivo. Another oncogenic miRNA is the cluster miR-17-92, which is located in the MIR17HG gene in chromosome 13 (13q31.3) and is surrounded by the LIN00379 gene in the 5′ region and GPC5 gene in the 3′ region. miR-17-92 is over-expressed in anaplastic thyroid cancer, the most aggressive and lethal form of thyroid cancer (Takakura et al. 2008). The temporary inhibition of miR-17-92 due to antimiR treatment resulted in anti-tumoral effects such as reduction of cell growth, as well as induction of apoptosis and senescence in ATC cell lines (Calabrese et al. 2019; Takakura et al. 2008). Moreover, it de-repressed PTEN and Rb, predicted targets of the cluster (Takakura et al. 2008). Indeed, the literature shows that miR-17-92 targets several tumor suppressor genes, such as PTEN, SMAD4, TGFBR2, and CDKN1 (p21) (Fuziwara and Kimura 2015).

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Aiming to edit miR-17-92 cluster gene in ATC cells, we relied on studies that showed primary mir-17-92, which is a long ~800nt RNA, forms a globular tertiary structure that is differentially processed by microprocessor, a component of miRNA machinery (Du et al. 2015; Chakraborty et al. 2012). In this tertiary structure, the region located in the 5′ portion of the miR-17 sequence is important for the correct folding and processing of all miRNAs due to additional binding to splicing machinery components (Du et al. 2015). Therefore, we designed sgRNAs to target the 3′ region in the pre-mir-17 using CRISPR/Cas9n-double nicking (Fuziwara et al. 2020). As a result, the targeted region was edited with specific small deletions, and miR-17 levels were blunted due to the proximity of the gene-editing target site, but we observed a global down-regulation in the other five miRNAs (miR-18a, miR-19a, miR-20a, miR-19b, and miR-92a). Functionally, gene editing of miR-1792 resulted in antitumoral effects in the ATC cell line KTC2 such as reduction of migration and proliferation while inducing cell differentiation. These studies on miRNA (miR-146b and miR-17-92) corroborate CRISPR/ Cas9n system’s effectiveness in targeting miRNA noncoding genes and reducing their expression in thyroid cancer cell lines (Fuziwara et al. 2022). The discovery of a large number of noncoding genes, especially lncRNAs, associated with a better understanding of the mechanism of noncoding action by recent publications, explicit these noncoding sequences’ role in the regulation of coding genes and genomic stability (Statello et al. 2021). In this context, lncRNAs are a class of noncoding RNAs with >200t that is very heterogeneous in terms of size, resulting in different mechanisms of action. For example, Air has 108 Kb (Sleutels et al. 2002) and Kcnq1ot1 has 91 Kb (Pandey et al. 2008). lncRNAs have a broader spectrum of effects, ranging from regulation of a gene’s transcriptional activity by mediating the interactions of proteins with chromatin to posttranscriptional effects by regulating the interactions between miRNAs and mRNAs (Statello et al. 2021). A well-known lncRNA is the human XIST, an lncRNA ~19kb in size that is involved in the X chromosome inactivation by coating and recruiting chromatinremodeling proteins (Wutz 2011). The longer the lncRNA, the more interactions are predicted to occur with other RNAs or proteins. For example, XIST shows predicted interactions with more than 200 miRNAs and 29 RNA-binding proteins in the Starbase v2.0 database (Li et al. 2014) In the field of thyroid cancer, an increasing number of publications have reported lncRNA deregulation’s effects on tumor cell biology. However, the literature is still scarce regarding editing an lncRNA with CRISPR/Cas9. Targeting an lncRNA is possible but doing so involves intrinsic difficulties due to (1) lncRNA transcripts’ size; (2) their noncoding nature, meaning that a slight change in the genomic sequence might not alter a lncRNA’s structure or function of a lncRNA as it would with a coding-gene (change of protein frame); and (3) the absence of a conserved regulatory region due to the variability of lncRNA size and function, such as what is observed in miRNA precursor structure. In one study, the role of the small nucleolar host gene 3 (SNHG3), a 2346-bp lncRNA that is differentially expressed in thyroid cancer, was investigated (Duan

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Table 5.2 Thyroid cancer cells edited with CRISPR/Cas9 Cell line SW579 Mouse-derived Tpo-Cre/HrasG12V/p53flox/flox KTC1 KTC2

Cancer ATC PDTC

Gene EGFR Nf1

Type of gene Coding Coding

References Huang et al. (2018) Untch et al. (2018)

PTC ATC

NF2 MIR17HG

You et al. (2019) Fuziwara et al. (2020)

KTC2

ATC

MIR146B

BCPAP and TPC1

PTC

SNHG3

Coding Noncoding (miRNA) Noncoding (miRNA) Noncoding (lncRNA)

Santa-Inez et al. (2021) Duan et al. (2020)

et al. 2020). When the researchers compared PTC samples to normal thyroid tissue, they found a reduction in the expression of SNHG3, and lower levels of SNHG3 were associated with a shorter recurrence-free survival period and poor clinical prognosis. Indeed, targeting SNHG3 with CRISPR/Cas9 in PTC cell lines BCPAP and TPC1 increased cell proliferation and colony formation and induced tumor growth in nude mice (Duan et al. 2020), indicating a tumor suppressor role for this lncRNA in thyroid cancer.

4 Final Considerations Gene editing with CRISPR/Cas9 is a methodology that allows for permanent modification of genes and was successfully used to study thyroid cancer biology, as Table 5.2 shows. Moreover, it became an alternative method to silence coding and noncoding genes. Importantly, CRISPR/Cas9-mediated gene editing provides a new potential therapeutic perspective for aggressive thyroid cancer where current clinical approaches have failed. Therefore, cell line studies are essential to validate further this methodology until we reach a safe point to implement it as therapy and to uncover the molecular mechanisms of tumor progression and resistance to drugs.

References Bible KC, Kebebew E, Brierley J, Brito JP, Cabanillas ME, Clark TJ Jr, Di Cristofano A, Foote R, Giordano T, Kasperbauer J, Newbold K, Nikiforov YE, Randolph G, Rosenthal MS, Sawka AM, Shah M, Shaha A, Smallridge R, Wong-Clark CK (2021) 2021 American Thyroid Association guidelines for management of patients with anaplastic thyroid cancer. Thyroid 31:337–386 Brose MS, Cabanillas ME, Cohen EE, Wirth LJ, Riehl T, Yue H, Sherman SI, Sherman EJ (2016) Vemurafenib in patients with BRAF(V600E)-positive metastatic or unresectable papillary thyroid cancer refractory to radioactive iodine: a non-randomised, multicentre, open-label, phase 2 trial. Lancet Oncol 17:1272–1282

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Cabanillas ME, Ferrarotto R, Garden AS, Ahmed S, Busaidy NL, Dadu R, Williams MD, Skinner H, Gunn GB, Grosu H, Iyer P, Hofmann MC, Zafereo M (2018) Neoadjuvant BRAF- and immune-directed therapy for anaplastic thyroid carcinoma. Thyroid 28:945–951 Cabanillas ME, Ryder M, Jimenez C (2019) Targeted therapy for advanced thyroid cancer: Kinase inhibitors and beyond. Endocr Rev 40:1573–1604 Calabrese G, Dolcimascolo A, Caruso G, Forte S (2019) miR-19a is involved in progression and malignancy of anaplastic thyroid cancer cells. Onco Targets Ther 12:9571–9583 Chakraborty S, Mehtab S, Patwardhan A, Krishnan Y (2012) Pri-miR-17-92a transcript folds into a tertiary structure and autoregulates its processing. RNA 18:1014–1028 Chou CK, Chen RF, Chou FF, Chang HW, Chen YJ, Lee YF, Yang KD, Cheng JT, Huang CC, Liu RT (2010) miR-146b is highly expressed in adult papillary thyroid carcinomas with high risk features including extrathyroidal invasion and the BRAF(V600E) mutation. Thyroid 20: 489–494 Du P, Wang L, Sliz P, Gregory RI (2015) A biogenesis step upstream of microprocessor Controls miR-17 approximately 92 Expression. Cell 162:885–899 Duan Y, Wang Z, Xu L, Sun L, Song H, Yin H, He F (2020) lncRNA SNHG3 acts as a novel tumor suppressor and regulates tumor proliferation and metastasis via AKT/mTOR/ERK pathway in papillary thyroid carcinoma. J Cancer 11:3492–3501 Esquela-Kerscher A, Slack FJ (2006) Oncomirs – microRNAs with a role in cancer. Nat Rev Cancer 6:259–269 Fagin JA, Wells SA Jr (2016) Biologic and clinical perspectives on thyroid cancer. N Engl J Med 375:1054–1067 Friedman RC, Farh KK, Burge CB, Bartel DP (2009) Most mammalian mRNAs are conserved targets of microRNAs. Genome Res 19:92–105 Fuziwara CS, Kimura ET (2015) Insights into regulation of the miR-17-92 cluster of miRNAs in cancer. Front Med (Lausanne) 2:64 Fuziwara CS, Kimura ET (2017) MicroRNAs in thyroid development, function and tumorigenesis. Mol Cell Endocrinol 456:44–50 Fuziwara CS, Saito KC, Kimura ET (2020) Thyroid follicular cell loss of differentiation induced by microRNA miR-17-92 cluster is attenuated by CRISPR/Cas9n gene silencing in anaplastic thyroid cancer. Thyroid 30:81–94 Fuziwara CS, De Mello DC, Kimura ET (2022) Gene editing with CRISPR/Cas methodology and thyroid cancer: where are we? Cancers (Basel) 14:844 Garcia-Rendueles ME, Ricarte-Filho JC, Untch BR, Landa I, Knauf JA, Voza F, Smith VE, Ganly I, Taylor BS, Persaud Y, Oler G, Fang Y, Jhanwar SC, Viale A, Heguy A, Huberman KH, Giancotti F, Ghossein R, Fagin JA (2015) NF2 loss promotes oncogenic RAS-induced thyroid cancers via YAP-dependent transactivation of RAS proteins and sensitizes them to MEK inhibition. Cancer Discov 5:1178–1193 Gebert LFR, Macrae IJ (2019) Regulation of microRNA function in animals. Nat Rev Mol Cell Biol 20:21–37 Geraldo MV, Yamashita AS, Kimura ET (2012) MicroRNA miR-146b-5p regulates signal transduction of Tgf-beta by repressing Smad4 in thyroid cancer. Oncogene 31:1910–1922 Grieco M, Santoro M, Berlingieri MT, Melillo RM, Donghi R, Bongarzone I, Pierotti MA, Della Porta G, Fusco A, Vecchio G (1990) PTC is a novel rearranged form of the ret proto-oncogene and is frequently detected in vivo in human thyroid papillary carcinomas. Cell 60:557–563 Ha M, Kim VN (2014) Regulation of microRNA biogenesis. Nat Rev Mol Cell Biol 15:509–524 Haugen BR, Alexander EK, Bible KC, Doherty GM, Mandel SJ, Nikiforov YE, Pacini F, Randolph GW, Sawka AM, Schlumberger M, Schuff KG, Sherman SI, Sosa JA, Steward DL, Tuttle RM, Wartofsky L (2016) 2015 American Thyroid Association Management guidelines for adult patients with thyroid nodules and differentiated thyroid cancer: the American Thyroid Association guidelines task force on thyroid nodules and differentiated thyroid cancer. Thyroid 26: 1–133

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C. S. Fuziwara and E. T. Kimura

He L, Thomson JM, Hemann MT, Hernando-Monge E, Mu D, Goodson S, Powers S, CordonCardo C, Lowe SW, Hannon GJ, Hammond SM (2005) A microRNA polycistron as a potential human oncogene. Nature 435:828–833 Huang LC, Tam KW, Liu WN, Lin CY, Hsu KW, Hsieh WS, Chi WM, Lee AW, Yang JM, Lin CL, Lee CH (2018) CRISPR/Cas9 genome editing of epidermal growth factor receptor sufficiently abolished oncogenicity in anaplastic thyroid cancer. Dis Markers 2018:3835783 International Human Genome Sequencing Consortium (2004) Finishing the euchromatic sequence of the human genome. Nature 431:931–945 Jhiang SM, Sagartz JE, Tong Q, Parker-Thornburg J, Capen CC, Cho JY, Xing S, Ledent C (1996) Targeted expression of the ret/PTC1 oncogene induces papillary thyroid carcinomas. Endocrinology 137:375–378 Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337:816–821 Johnson SM, Grosshans H, Shingara J, Byrom M, Jarvis R, Cheng A, Labourier E, Reinert KL, Brown D, Slack FJ (2005) RAS is regulated by the let-7 microRNA family. Cell 120:635–647 Kimura ET, Nikiforova MN, Zhu Z, Knauf JA, Nikiforov YE, Fagin JA (2003) High prevalence of BRAF mutations in thyroid cancer: genetic evidence for constitutive activation of the RET/PTC-RAS-BRAF signaling pathway in papillary thyroid carcinoma. Cancer Res 63: 1454–1457 Kirschner LS, Qamri Z, Kari S, Ashtekar A (2016) Mouse models of thyroid cancer: a 2015 update. Mol Cell Endocrinol 421:18–27 Knauf JA, Ma X, Smith EP, Zhang L, Mitsutake N, Liao XH, Refetoff S, Nikiforov YE, Fagin JA (2005) Targeted expression of BRAFV600E in thyroid cells of transgenic mice results in papillary thyroid cancers that undergo dedifferentiation. Cancer Res 65:4238–4245 Knauf JA, Luckett KA, Chen KY, Voza F, Socci ND, Ghossein R, Fagin JA (2018) Hgf/Met activation mediates resistance to BRAF inhibition in murine anaplastic thyroid cancers. J Clin Invest 128:4086–4097 Kozomara A, Birgaoanu M, Griffiths-Jones S (2019) miRBase: from microRNA sequences to function. Nucleic Acids Res 47:D155–D162 Landa I, Ibrahimpasic T, Boucai L, Sinha R, Knauf JA, Shah RH, Dogan S, Ricarte-Filho JC, Krishnamoorthy GP, Xu B, Schultz N, Berger MF, Sander C, Taylor BS, Ghossein R, Ganly I, Fagin JA (2016) Genomic and transcriptomic hallmarks of poorly differentiated and anaplastic thyroid cancers. J Clin Invest 126:1052–1066 Lemoine NR, Mayall ES, Wyllie FS, Williams ED, Goyns M, Stringer B, Wynford-Thomas D (1989) High frequency of ras oncogene activation in all stages of human thyroid tumorigenesis. Oncogene 4:159–164 Lewis BP, Shih IH, Jones-Rhoades MW, Bartel DP, Burge CB (2003) Prediction of mammalian microRNA targets. Cell 115:787–798 Li JH, Liu S, Zhou H, Qu LH, Yang JH (2014) starBase v2.0: decoding miRNA-ceRNA, miRNAncRNA and protein-RNA interaction networks from large-scale CLIP-Seq data. Nucleic Acids Res 42:D92–D97 Lima CR, Geraldo MV, Fuziwara CS, Kimura ET, Santos MF (2016) MiRNA-146b-5p upregulates migration and invasion of different Papillary Thyroid Carcinoma cells. BMC Cancer 16:108 Mcgeary SE, Lin KS, Shi CY, Pham TM, Bisaria N, Kelley GM, Bartel DP (2019) The biochemical basis of microRNA targeting efficacy. Science 366:eaav1741 Mitsutake N, Knauf JA, Mitsutake S, Mesa C Jr, Zhang L, Fagin JA (2005) Conditional Brafv600E expression induces DNA synthesis, apoptosis, dedifferentiation, and chromosomal instability in thyroid PCCL3 cells. Cancer Res 65:2465–2473 Montero-Conde C, Ruiz-Llorente S, Dominguez JM, Knauf JA, Viale A, Sherman EJ, Ryder M, Ghossein RA, Rosen N, Fagin JA (2013) Relief of feedback inhibition of HER3 transcription by RAF and MEK inhibitors attenuates their antitumor effects in BRAF-mutant thyroid carcinomas. Cancer Discov 3:520–533

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Nurk S, Koren S, Rhie A, Rautiainen M, Bzikadze AV, Mikheenko A, Vollger MR, Altemose N, Uralsky L, Gershman A, Aganezov S, Hoyt SJ, Diekhans M, Logsdon GA, Alonge M, Antonarakis SE, Borchers M, Bouffard GG, Brooks SY, Caldas GV, Chen NC, Cheng H, Chin CS, Chow W, De Lima LG, Dishuck PC, Durbin R, Dvorkina T, Fiddes IT, Formenti G, Fulton RS, Fungtammasan A, Garrison E, Grady PGS, Graves-Lindsay TA, Hall IM, Hansen NF, Hartley GA, Haukness M, Howe K, Hunkapiller MW, Jain C, Jain M, Jarvis ED, Kerpedjiev P, Kirsche M, Kolmogorov M, Korlach J, Kremitzki M, Li H, Maduro VV, Marschall T, Mccartney AM, Mcdaniel J, Miller DE, Mullikin JC, Myers EW, Olson ND, Paten B, Peluso P, Pevzner PA, Porubsky D, Potapova T, Rogaev EI, Rosenfeld JA, Salzberg SL, Schneider VA, Sedlazeck FJ, Shafin K, Shew CJ, Shumate A, Sims Y, Smit AFA, Soto DC, Sovic I, Storer JM, Streets A, Sullivan BA, Thibaud-Nissen F, Torrance J, Wagner J, Walenz BP, Wenger A, Wood JMD, Xiao C, Yan SM, Young AC, Zarate S, Surti U, Mccoy RC, Dennis MY, Alexandrov IA, Gerton JL, O’neill RJ, Timp W, Zook JM, Schatz MC, Eichler EE, Miga KH, Phillippy AM (2022) The complete sequence of a human genome. Science 376:44–53 Olive V, Bennett MJ, Walker JC, Ma C, Jiang I, Cordon-Cardo C, Li QJ, Lowe SW, Hannon GJ, He L (2009) miR-19 is a key oncogenic component of mir-17-92. Genes Dev 23:2839–2849 Ortiga-Carvalho TM, Chiamolera MI, Pazos-Moura CC, Wondisford FE (2016) Hypothalamuspituitary-thyroid axis. Compr Physiol 6:1387–1428 Pandey RR, Mondal T, Mohammad F, Enroth S, Redrup L, Komorowski J, Nagano T, ManciniDinardo D, Kanduri C (2008) Kcnq1ot1 antisense noncoding RNA mediates lineage-specific transcriptional silencing through chromatin-level regulation. Mol Cell 32:232–246 Ramirez-Moya J, Wert-Lamas L, Santisteban P (2018) Microrna-146b promotes PI3K/AKT pathway hyperactivation and thyroid cancer progression by targeting Pten. Oncogene 37:3369–3383 Ran FA, Hsu PD, Lin CY, Gootenberg JS, Konermann S, Trevino AE, Scott DA, Inoue A, Matoba S, Zhang Y, Zhang F (2013) Double nicking by RNA-guided CRISPR Cas9 for enhanced genome editing specificity. Cell 154:1380–1389 Ricarte-Filho JC, Ryder M, Chitale DA, Rivera M, Heguy A, Ladanyi M, Janakiraman M, Solit D, Knauf JA, Tuttle RM, Ghossein RA, Fagin JA (2009) Mutational profile of advanced primary and metastatic radioactive iodine-refractory thyroid cancers reveals distinct pathogenetic roles for BRAF, PIK3CA, and AKT1. Cancer Res 69:4885–4893 Saavedra HI, Knauf JA, Shirokawa JM, Wang J, Ouyang B, Elisei R, Stambrook PJ, Fagin JA (2000) The RAS oncogene induces genomic instability in thyroid PCCL3 cells via the Mapk pathway. Oncogene 19:3948–3954 Santa-Inez DC, Fuziwara CS, Saito KC, Kimura ET (2021) Targeting the Highly Expressed microRNA miR-146b with CRISPR/Cas9n Gene Editing System in Thyroid Cancer. Int J Mol Sci 22:7992 Schiff BA, Mcmurphy AB, Jasser SA, Younes MN, Doan D, Yigitbasi OG, Kim S, Zhou G, Mandal M, Bekele BN, Holsinger FC, Sherman SI, Yeung SC, El-Naggar AK, Myers JN (2004) Epidermal growth factor receptor (EGFR) is overexpressed in anaplastic thyroid cancer, and the EGFR inhibitor gefitinib inhibits the growth of anaplastic thyroid cancer. Clin Cancer Res 10: 8594–8602 Schweppe RE, Klopper JP, Korch C, Pugazhenthi U, Benezra M, Knauf JA, Fagin JA, Marlow LA, Copland JA, Smallridge RC, Haugen BR (2008) Deoxyribonucleic acid profiling analysis of 40 human thyroid cancer cell lines reveals cross-contamination resulting in cell line redundancy and misidentification. J Clin Endocrinol Metab 93:4331–4341 Shonka DC Jr, Ho A, Chintakuntlawar AV, Geiger JL, Park JC, Seetharamu N, Jasim S, Abdelhamid Ahmed AH, Bible KC, Brose MS, Cabanillas ME, Dabekaussen K, Davies L, Dias-Santagata D, Fagin JA, Faquin WC, Ghossein RA, Gopal RK, Miyauchi A, Nikiforov YE, Ringel MD, Robinson B, Ryder MM, Sherman EJ, Sadow PM, Shin JJ, Stack BC Jr, Tuttle RM, Wirth LJ, Zafereo ME Jr, Randolph GW (2022) American Head and Neck Society Endocrine Surgery Section and International Thyroid Oncology Group consensus statement on mutational testing in thyroid cancer: Defining advanced thyroid cancer and its targeted treatment. Head Neck 44:1277–1300

84

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Siegel RL, Miller KD, Fuchs HE, Jemal A (2022) Cancer statistics, 2022. CA Cancer J Clin 72: 7–33 Sleutels F, Zwart R, Barlow DP (2002) The non-coding air RNA is required for silencing autosomal imprinted genes. Nature 415:810–813 Statello L, Guo CJ, Chen LL, Huarte M (2021) Gene regulation by long non-coding RNAs and its biological functions. Nat Rev Mol Cell Biol 22:96–118 Subbiah V, Kreitman RJ, Wainberg ZA, Cho JY, Schellens JHM, Soria JC, Wen PY, Zielinski C, Cabanillas ME, Urbanowitz G, Mookerjee B, Wang D, Rangwala F, Keam B (2018) Dabrafenib and trametinib treatment in patients with locally advanced or metastatic BRAF V600-mutant anaplastic thyroid cancer. J Clin Oncol 36:7–13 Takakura S, Mitsutake N, Nakashima M, Namba H, Saenko VA, Rogounovitch TI, Nakazawa Y, Hayashi T, Ohtsuru A, Yamashita S (2008) Oncogenic role of miR-17-92 cluster in anaplastic thyroid cancer cells. Cancer Sci 99:1147–1154 Takamizawa J, Konishi H, Yanagisawa K, Tomida S, Osada H, Endoh H, Harano T, Yatabe Y, Nagino M, Nimura Y, Mitsudomi T, Takahashi T (2004) Reduced expression of the let-7 microRNAs in human lung cancers in association with shortened postoperative survival. Cancer Res 64:3753–3756 The ENCODE Project Consortium (2012) An integrated encyclopedia of DNA elements in the human genome. Nature 489:57–74 Untch BR, Dos Anjos V, Garcia-Rendueles MER, Knauf JA, Krishnamoorthy GP, Saqcena M, Bhanot UK, Socci ND, Ho AL, Ghossein R, Fagin JA (2018) Tipifarnib inhibits HRAS-driven dedifferentiated thyroid cancers. Cancer Res 78:4642–4657 Uribe ML, Marrocco I, Yarden Y (2021) EGFR in cancer: signaling mechanisms, drugs, and acquired resistance. Cancers (Basel) 13:2748 Vanderpump MP (2011) The epidemiology of thyroid disease. Br Med Bull 99:39–51 Vitagliano D, Portella G, Troncone G, Francione A, Rossi C, Bruno A, Giorgini A, Coluzzi S, Nappi TC, Rothstein JL, Pasquinelli R, Chiappetta G, Terracciano D, Macchia V, Melillo RM, Fusco A, Santoro M (2006) Thyroid targeting of the N-ras(Gln61Lys) oncogene in transgenic mice results in follicular tumors that progress to poorly differentiated carcinomas. Oncogene 25: 5467–5474 Volders PJ, Anckaert J, Verheggen K, Nuytens J, Martens L, Mestdagh P, Vandesompele J (2019) LNCipedia 5: towards a reference set of human long non-coding RNAs. Nucleic Acids Res 47: D135–D139 Wang J, Knauf JA, Basu S, Puxeddu E, Kuroda H, Santoro M, Fusco A, Fagin JA (2003) Conditional expression of RET/PTC induces a weak oncogenic drive in thyroid PCCL3 cells and inhibits thyrotropin action at multiple levels. Mol Endocrinol 17:1425–1436 Wang JR, Zafereo ME, Dadu R, Ferrarotto R, Busaidy NL, Lu C, Ahmed S, Gule-Monroe MK, Williams MD, Sturgis EM, Goepfert RP, Gross ND, Lai SY, Gunn GB, Phan J, Rosenthal DI, Fuller CD, Morrison WH, Iyer P, Cabanillas ME (2019) Complete surgical resection following neoadjuvant dabrafenib plus trametinib in BRAF(V600E)-mutated anaplastic thyroid carcinoma. Thyroid 29:1036–1043 Wutz A (2011) Gene silencing in X-chromosome inactivation: advances in understanding facultative heterochromatin formation. Nat Rev Genet 12:542–553 Yip L, Kelly L, Shuai Y, Armstrong MJ, Nikiforov YE, Carty SE, Nikiforova MN (2011) MicroRNA signature distinguishes the degree of aggressiveness of papillary thyroid carcinoma. Ann Surg Oncol 18:2035–2041 You MH, Jeon MJ, Kim TY, Kim WB, Shong YK, Kim WG (2019) Expression of NF2 modulates the progression of BRAF(V600E) mutated thyroid cancer cells. Endocrinol Metab (Seoul) 34: 203–212 Zhao M, Sano D, Pickering CR, Jasser SA, Henderson YC, Clayman GL, Sturgis EM, Ow TJ, Lotan R, Carey TE, Sacks PG, Grandis JR, Sidransky D, Heldin NE, Myers JN (2011) Assembly and initial characterization of a panel of 85 genomically validated cell lines from diverse head and neck tumor sites. Clin Cancer Res 17:7248–7264

Chapter 6

Genome Editing for Engineering the Next Generation of Advanced Immune Cell Therapies Sarah Caroline Gomes de Lima, Daianne Maciely Carvalho Fantacini, Izadora Peter Furtado, Rafaela Rossetti, Roberta Maraninchi Silveira, Dimas Tadeu Covas, and Lucas Eduardo Botelho de Souza

1 Introduction The revolutionary wave in cancer immunotherapy came with a better understanding of how to re-awake and enhance the immune system to fight cancer. Although the first evidence of harnessing the host immune system to eradicate cancer could trace back to a century ago with William Bradley Coley’s findings (McCarthy 2006), cancer immunotherapy has blossomed into fruition only in recent years after significant advances in basic and clinical investigations (Zhang and Zhang 2020; Dobosz and Dzieciątkowski 2019; Waldman et al. 2020). Consequently, in 2013, Science named cancer immunotherapy the Breakthrough of the Year due to remarkable progress in two fields: immune modulation using antibodies that block immune regulatory checkpoints and the adoptive transfer of chimeric antigen receptor (CAR)-modified T cells (Couzin-Frankel 2013). Immune checkpoint (IC) inhibitors as well as adoptive cell therapy have emerged as core pillars of immunotherapy. ICs are immune-cell surface receptors that control immune homeostasis and are particularly relevant to T-cell functionality (Johnson et al. 2022). The best-described checkpoint molecules include the cytotoxic T

S. C. G. de Lima · I. P. Furtado · R. Rossetti · R. M. Silveira · D. T. Covas · L. E. B. de Souza (✉) Blood Center of Ribeirão Preto – Ribeirão Preto School of Medicine, University of São Paulo, Ribeirão Preto, SP, Brazil e-mail: [email protected]; [email protected]; [email protected]; [email protected]; [email protected]; [email protected] D. M. C. Fantacini Biotechnology Nucleus of Ribeirão Preto – Butantan Institute, Ribeirão Preto, SP, Brazil e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_6

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lymphocyte-associated molecule-4 (CTLA-4), programmed cell death receptor-1 (PD-1), and programmed cell death ligand-1 (PD-L1). Both CTLA-4 and PD-1 are upregulated on the surface of active T cells to prevent excessive stimulation by the T-cell receptor (TCR). In contrast, the tumor microenvironment (TME) is characterized by overexpression of PD-L1 by cancer cells (Marin-Acevedo et al. 2021). The PD-1 pathway drives T cells into a dysfunctional state known as T-cell exhaustion. Molecules that block these pathways have been developed and become the standard of care in the treatment of many malignancies, such as melanoma, nonsmall-cell lung cancer (NSCLC), urothelial cancer, and renal cell cancer. However, one of the related toxicities of this treatment that impairs the overall survival is the manifestation of immune-related Adverse Events (irAEs) (Franzin et al. 2020). Unlike IC inhibitor therapy, in which monoclonal antibodies target immune checkpoint proteins, adoptive cell therapy is based on genetic modification of T cells isolated from the patient to express a specific TCR or CAR, followed by ex vivo cell expansion and infusion back into the patient to mediate an antitumor function. In TCR gene therapy, genes encoding the α- and β-chains of the TCR that recognize tumor-specific targets are cloned into viral or nonviral vectors and transferred into patient’s T cells (Dossett et al. 2009; Peng et al. 2009). Such TCR-modified T cells are redirected to recognize tumor-specific epitopes in a major histocompatibility complex (MHC)-dependent manner. For this reason, one practical limitation is that the most available TCR T-cell therapies in clinical trials are restricted to MHC proteins of human leukocyte antigen (HLA)-A*02:01 allele, as this is present at most in 40–45% of patients of Caucasian descent (Song et al. 2013). Moreover, chain mispairing between endogenous and introduced α- and β-chains counterparts can occur and lead to not only a reduced expression of the engineered TCR but also introduce the significant risk of autoimmunity since a new TCR with unknown specificity can be formed (Shao et al. 2010). CAR T cells, however, recognize the antigen in an MHC-unrestricted way. Briefly, CARs are synthetic receptors consisting of four elements: an extracellular domain, typically a single-chain variable fragment (scFv) derived from tumor antigen-reactive antibody, a transmembrane domain, and an intracellular T-cell activation and co-stimulation signaling domain commonly composed of CD3ζ, CD28, and/or 4-1BB. CAR T designs have been extensively reviewed elsewhere and will not be further elaborated on this chapter (Sadelain et al. 2013; Jayaraman et al. 2020). The successful use of CAR T cells was a breakthrough in the treatment of hematological malignancies. There are currently six CAR T-cell products approved by the Food and Drug Administration (FDA), which are available against B-cell lymphomas, B-cell acute lymphoblastic leukemia (B-ALL), and multiple myeloma (MM). However, due to its personalized feature, autologous cell therapy is endowed with limitations, such as a time-consuming and costly process with an inherent risk of production failure as a consequence of the quality and quantity of patient-derived T cells (Zhao et al. 2018; Ceppi et al. 2018). Additionally, there is a major concern about the use of retroviral vectors as a gene carrier to encode CARs due to their random integration in the host genome (Zhang et al. 2017a).

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While CAR T cells have shown impressive results against B-cell hematological cancers, the same has not been true for solid tumors. This is explained by the barriers imposed by the tumor microenvironment to restrict T-cell function, such as physical barriers to T-cell infiltration, the presence of immunosuppressive cytokines (such as transforming growth factor β (TGFβ) and Interleukin (IL)-10), and overexpression of IC molecules (Hou et al. 2021; Kankeu Fonkoua et al. 2022). Another bottleneck associated with CAR T-cell therapy is the development of an efficient therapy against T-cell malignancies, mainly because effector T cells and malignant T cells tend to share the expression of target antigens, which results in fratricide, limiting the therapeutic potential (Cooper et al. 2018). Off-the-shelf, allogeneic approaches, in contrast, may potentially change the treatment landscape since it is possible to generate from a single batch of engineered CAR T cells high amounts of functional cells enough to treat hundreds of patients (Depil et al. 2020). However, the administration of allogeneic T cells has a potential risk of inducing graft-versus-host disease (GvHD). In addition, the host immune system might, in turn, induce allorejection, which will lead to an inefficient therapy against the tumor. To overcome these challenges, genome editing tools are opening a new era in the engineering of T cells to be used as universal cellular products with limited or no potential for GvHD. Furthermore, the site-specific genetic editing provides an opportunity to direct therapeutic transgenes to specific genomic loci and therefore, generate safer cell products. Additionally, genome editing can be employed to efficiently knock out genes that work as immune checkpoint regulators and eliminate the expression of shared target antigens to avoid fratricide. Genome editing tools are mostly based on the use of engineered nucleases that can modify the DNA through the creation of a double-strand break (DSB) at a specific genomic site (Li et al. 2020). Once the DSB occurs, cellular DNA repair mechanisms will be activated, which may occur via homology-directed repair (HDR) or nonhomologous end-joining (NHEJ). HDR is a precise repair mechanism that needs a homologous DNA template to repair the missing DNA at DSB. In contrast, NHEJ is an error-prone mechanism and often results in the formation of insertions or deletions (indels) (O’Driscoll and Jeggo 2006). The three major generations of genome editing tools are zinc finger nucleases (ZFNs), transcription activator-like endonucleases (TALENs), and clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated Cas9 endonucleases (CRISPR/Cas9). ZFNs were the first gene editors to enter the clinic (Ledford 2011). They are artificial endonucleases that consist of a designed zinc finger protein (ZFP) that recognizes three specific base pairs of DNA fused to the cleavage domain of the FokI endonuclease (Urnov et al. 2010). Similar to ZFNs, TALENs are another type of engineered nuclease that comprise a nonspecific DNA cleavage domain fused to a customizable sequence-specific DNA-binding domain to induce DSBs into specific DNA sites (Joung and Sander 2012). Differently from ZFNs and TALENS, which rely on protein-DNA interaction for specificity determination, CRISPR/Cas9 uses a single-guide RNA (gRNA) to recognize the DNA sequence and redirect Cas

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endonuclease activity (Cho et al. 2013). More recently, a newer form of genome editing technology that does not induce DSBs has been developed, eliminating safety concerns related to the emergence of chromosomal aberrations as demonstrated elsewhere (Stadtmauer et al. 2020; Nahmad et al. 2022). DNA base editing tools break only a single strand of the target DNA sequence and install targeted point mutations (converting C to T or A to G) through a Cas9 catalytically impaired fused to a deaminase. This mechanism does not rely on HDR, thereby mitigating the genome instability resulting from a DSB (Komor et al. 2016; Gaudelli et al. 2017). Each one of these genome editing technologies has different characteristics that allow target editing of specific sequences within the genome. However, the further description concerning the detailed mechanism of their function and pros and cons is beyond the scope of this chapter, and thereby audiences are referred to some excellent reviews (Li et al. 2020; Urnov et al. 2010; Joung and Sander 2012; Porteus and Carroll 2005; Anzalone et al. 2020). This burst of innovation has led to the current research efforts focusing on the precise engineering of T cells to optimize the manufacturing process, improve therapeutic efficiency, and broaden the repertoire of possible targets for cancer immunotherapies. This chapter addresses why and how genome editing has been applied to advanced immune cell therapies (Fig. 6.1) and discusses several ongoing and future directions toward the establishment of the next generation of cell-based immunotherapies.

2 Exploring Genome Editing Tools for Advanced CAR T-Cell Manufacturing Most clinical trials and all commercially available CAR T-cell products require gamma-retroviral or lentiviral vectors for CAR transgene delivery into T cells (Labbé et al. 2021), which is a highly efficient process and provides long-term CAR expression. Despite the overall safety pattern of retroviral-mediated modification of CAR T cells in a long-term evaluation (Scholler et al. 2012), concerns still arise from the retroviral profile of random integration into the genome and the possibility of insertional mutagenesis. In a case report, Fraietta et al. described a lentiviral-mediated integration of the CAR coding sequence into the TET2 (Tet methylcytosine dioxygenase 2) tumor-suppressor gene in CAR T cells produced for a patient with chronic lymphocytic leukemia (Fraietta et al. 2018a). Surprisingly, the TET2 dysfunction, resulting from both insertion-mediated disruption in one allele and the patient’s inherent mutation in the second one, gave rise to more potent CAR T cells with enhanced expansion and persistence without signs of malignant transformation. However, this event illustrates the potential risks of the random pattern of viral vector integration, as observed previously for other human gene therapy (Hacein-Bey-Abina et al. 2008).

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Fig. 6.1 Advanced genome editing tools for improved T-cell immunotherapies. Genome editing technologies are addressed in this chapter as tools for boosting four aspects of T-cell immunotherapies. (I) Advanced manufacturing strategies: The sequence-specific DNA cleavage by endonucleases coupled with the availability of a donor template can redirect homology-directed repair (HDR) machinery toward the precise insertion of a new sequence into the genome. Engineered homing endonucleases (HM), megaTAL (mT), TALEN (T), and CRISPR/Cas systems (C) are among the genome editing tools explored for the targeted insertion of an artificial receptor (CAR or artificial TCR) into the TRAC or PDCD1 locus, resulting in depletion of the corresponding proteins (TCR and PD-1, respectively) and inserted gene expression controlled by an endogenous promoter. In this strategy, the HDR donor template can be delivered through an adeno-associated virus (AAV) or as a free DNA molecule (single- or double-stranded) co-electroporated with the editing machinery. (II) Increased efficiency: TALEN and CRISPR/Cas9 were used for disruption of negative cell regulators, boosting T-cell activity against tumors. Negative regulators are immune checkpoint inhibitors and other molecules with a known impact on T-cell functionality or others screened by CRISPR/Cas9. (III) An allogeneic CAR T-cell therapy manufactured from healthy donors’ T cells is feasible through the disruption of key elements that mediate cell alloreactivity. Zinc finger endonucleases (Z), TALEN, and CRISPR/Cas9 have been explored to knock out the TRAC gene as a strategy to deplete the TCR from CAR T cells, avoiding the graft (allogeneic CAR T cell) against patient’s healthy cells response that otherwise could lead to the graft-versus-host disease (GvHD). Through a multiplex genome editing, TCR and HLA-I can be depleted from CAR T cells to eliminate the patient’s T-cell response against CAR T cells; at the same time, HLA-E can be expressed in CAR T cells to prevent natural killer (NK) cell-mediated cytotoxicity. (IV) Engineered T cells share the target antigen with the tumor T cell, leading to self-killing (fratricide). To produce fratricide-resistant cells, CRISPR/Cas9 and base editors (BE) were used to abrogate the expression of shared proteins (e.g., CD7 and CD3) on CAR T cells, expanding the repertoire of targets for the therapies against T-cell malignancies. The genome editing tools were represented by their initial letter after a scissor illustration. BE base editor, C CRISPR/Cas system, HE homing endonuclease, mT megaTAL nuclease, T transcription activator-like effector nuclease (TALEN), zinc finger nuclease (ZFN), AAV adeno-associated virus; artificial receptor: CAR or artificial TCR, GvHD graft-versus-host disease, HDR homology-directed repair, HLA human leukocyte antigen, PD-1 programmed cell death 1, PDCD1 PD-1 coding gene, TRAC T-cell receptor alpha constant, TCR T-cell receptor

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A promising alternative to random CAR transgene integration is its targeted insertion driven by genome editing tools. Through the induction of DNA DSBs by sequence-specific nucleases, a given donor template harboring the CAR sequence can guide the HDR machinery toward the new sequence insertion into the cut site (Li et al. 2020). In innovative studies, CAR insertion was directed into the endogenous TCR locus using engineered homing endonuclease, megaTAL nuclease, TALEN, or CRISPR/Cas9 system (MacLeod et al. 2017; Hale et al. 2017; Jo et al. 2022; Eyquem et al. 2017). All those strategies rely on adeno-associated virus (AAV)-mediated delivery of the HDR donor template, whose demonstrated high transduction efficiency and non-integrative nature contribute to a high site-specific insertion (Wang et al. 2016). The resulting combination of TCR knockout and CAR knock-in into one process streamlined the generation of allogeneic CAR T cells with a potent antitumor effect both in vitro and in murine models. Also, by placing the CAR transgene under the control of the TCR promoter, Eyquem and colleagues reported CAR T cells with homogeneous expression of the transgene, decreased tonic signaling, and internalization and re-expression dynamics of the receptor similar to that observed for the endogenous TCR, which contributed to delayed T-cell terminal differentiation and exhaustion (Eyquem et al. 2017). Moving forward, in a one-step process, an anti-CD22 CAR knock-in into the TRAC (T cell receptor alpha constant) locus coupled with PD-1 gene disruption was achieved using an mRNA-based CRISPR/Cpf1 system (Dai et al. 2019). PD-1 is a negative regulator of T-cell activity that will be discussed later in this chapter. CAR knock-in frequency was 81.88%, and PD-1 gene knockout reached 90.39% after CAR T-cell sorting. Using the same loci for multiple targeted-insertion based on the CRISPR/Cpf1 system, double knock-in rates of anti-CD22 and anti-CD19 CAR reached 85%, producing bi-specific CAR T cells with higher frequency, similar functional activity, and decreased exhaustion profile compared to that obtained with the ribonucleoprotein (RNP)-based CRISPR/Cas9 system (Dai et al. 2019). Cpf1 is a class 2 type V CRISPR system nuclease, which has a demonstrated high specificity profile in human cells (Kleinstiver et al. 2016). Its RNA guidance occurs through a CRISPR RNA (crRNA) independently of the trans-activating crRNA (tracrRNA), introducing DSB with a 5′ overhang at the target site (Zetsche et al. 2015). In the described strategy, crRNAs for each locus were encoded in an AAV vector in an array, which facilitates multiplex genome editing after their expression and autonomously processing by the Cpf1 nuclease (Zetsche et al. 2017). The HDR donor template was also included in the AAV vector. Despite the non-integrative nature of AAV vectors, the use of any viral vector is associated with increased costs due to the required good manufacturing practice (GMP)-grade production and extensive quality testing (Gándara et al. 2018). Then, further optimized approaches for targeted insertion have been proposed to eliminate the use of viral vectors. In a proof-of-concept, Roth and colleagues demonstrated that the co-electroporation of the editing machinery (Cas9 RNP) and an HDR donor template (double-stranded DNA) sustained up to 50% knock-in rates of fluorescent proteins while preserving T-cell viability (Roth et al. 2018). Besides that, bi-allelic or multiple modifications of two or three genes at once were possible using HDR donor

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templates encoding different fluorescent proteins. Moving to therapeutically relevant constructs, the authors replaced the endogenous TCR with an artificial NY-ESO-1specific TCR, producing reprogrammed T cells with robust antitumor activity (Roth et al. 2018). The mispairing between artificial and natural TCR is a point of concern since it might build a receptor with unknown specificity. Although mispairing tendency varies between TCRs with different specificity, artificial TCR knock-in into the TCR α locus coupled with TCR β chain disruption using a nonviral CRISPR/ Cas9 methodology reduced mispairing events (Schober et al. 2019). In the CAR context, virus-free CRISPR/Cas9 strategies have been explored for targeted integration of different CAR constructs into varying loci. In a comprehensive characterization, anti-GD2 CAR T cells generated through CAR knock-in into the TRAC locus displayed increased expression of memory-associated proteins and decreased expression of exhaustion markers compared to retrovirally produced CAR T cells (Mueller et al. 2022). At the same time, CAR T cells produced through the virus-free protocol showed robust cytotoxic activity against neuroblastoma tumor cells both in vitro and in vivo. Further optimizations of the technique are underway (Nguyen et al. 2020; Odé et al. 2020; Kath et al. 2022), showing the increasing interest in this promising area. Translating into the clinic, anti-CD19 CAR T cells were produced by knocking in the CAR transgene into the PD-1 locus and used to treat patients with relapsed/ refractory B cell non-Hodgkin lymphoma (r/r B-NHL) (ClinicalTrials.gov, NCT04213469). Complete remission was observed in seven (87.5%) out of eight patients, with durable responses in five of them 12 months later. Infusion product analysis showed an increased expression of memory-associated genes and decreased expression of dysfunction-related markers, which is consistent with the above descriptions for targeted insertion into the TRAC locus (Zhang et al. 2022a). Combined, these data show the feasibility of using a nonviral manufacturing process based on genome editing techniques to produce potent CAR T cells for a clinicalscale application.

3 Genome Editing for Increased Efficiency of Advanced T-Cell-Based Immunotherapies 3.1

Disrupting IC Inhibitors Through Genome Editing

IC molecules have a central role in regulating immunological homeostasis by repressing T-cell activity. Examples of well-described IC molecules include PD-1, CTLA-4, lymphocyte-activation gene 3 (LAG-3), and T-cell immunoglobulin and mucin 3 (TIM-3). By overexpressing IC molecules, tumors subvert this physiological mechanism to repress the activity of tumor-reactive immune cells, thereby evading immunological surveillance. Additionally, upregulation of IC molecules has been reported in genetically engineered T-cell products after injection (Moon

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et al. 2014, 2016), indicating that the immunosuppressive mechanisms triggered by the tumor microenvironment will likely restrict the efficacy of this therapy. Therefore, genome editing has been employed to knock out IC genes to generate more potent cell products that are resistant to tumor immunosuppression. Sun and colleagues demonstrated that knocking out PD-1 through CRISPR/Cas9 editing in primary T cells improved their cytotoxic activity (Su et al. 2016). Disruption of PD-1 in anti-CD19 CAR T increased its antitumor activity in vitro and in tumor xenograft models (Rupp et al. 2017). Also, a phase 1 clinical trial with refractory cancer patients evaluated the safety of engineering T cells to express an affinity-enhanced anti-NY-ESO-1 TCR and abrogate both chains of the endogenous TCR (TRAC and TRBC deletions) and PD-1 by CRISPR/Cas9 (ClinicalTrials.gov, NCT03399448). In general, the results demonstrated few off-target edits induced by CRISPR/Cas9 and no clinical toxicities associated with the edited TCR-T cells (Stadtmauer et al. 2020). In addition, PD-1 knockout mediated by CRISPR/Cas9 increased the therapeutic efficacy of CAR T cells against other tumor-associated antigens, such as mesothelin (Hu et al. 2019a), glypican-3 (Guo et al. 2018), and CD133 (Hu et al. 2019b). Similar results were obtained for TALEN-mediated PD-1 knockout, as edited tumor-reactive T cells displayed enhanced antitumor activity and persistence upon administration (Menger et al. 2016). Disruption of other IC molecules has been explored to increase the therapeutic efficacy of adoptively transferred T cells. Zhang and colleagues described a 45–70% efficiency of LAG-3 knockout in anti-CD19 CAR T cells using CRISPR/Cas9. However, there was no improvement in the antitumor activity of CAR T cells (Zhang et al. 2017b). The adenosine receptor expressed on T cells is also associated with immune suppression, and its depletion using CRISPR/Cas9 on human CAR T cells increased proinflammatory cytokines secretion, such as interferon-γ (IFN-γ) and tumor necrosis factor (TNF), in addition to improve mice survival. Furthermore, the observed low levels of off-target cut sites (only three predicted) did not elicit safety concerns (Giuffrida et al. 2021). The CRISPR/Cas9 system allows disruption of multiple genes simultaneously. This strategy has been applied for the generation of allogeneic CAR T cells via TCR and HLA-I depletions (a topic that will be detailed later in this chapter) concomitant with the knockout of the IC molecules PD-1 and CTLA-4. Although the authors succeeded in this multiplexed genome editing, they described that increasing the number of genes to be edited decreased the knockout efficiency, providing inferior results for this quadruple gene ablation (Ren et al. 2017a). Other studies have invested efforts in the generation of allogeneic CAR T cells edited by CRISPR/ Cas9 to delete PD-1 and thus improve its antitumor activity. In general, the data obtained demonstrate an improvement in antitumor activity upon PD-1 disruption in CAR T cells (Ren et al. 2017b). Together, these studies suggest that it is possible to efficiently generate functionenhanced engineered T cells through deletion of IC molecules using genome editing technologies. The improvement in the antitumor potential seems to depend on the edited IC, and, in this sense, interesting results have been described mainly for PD-1, both alone or associated with deletion of other ICs. Furthermore, new clinical studies

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may provide more detailed information regarding the safety of these edited cells. This type of genome edition strategy may provide an effective alternative to overcome the negative effects often associated with long-term use of IC inhibitors.

3.2

Targeting Other Regulators of Immune Activity to Enhance the Safety and Therapeutic Efficacy of Advanced T-Cell-Based Immunotherapies

For several years, a strong effort has been made to engineer immune cells to improve their antitumor activity. Besides knocking out IC molecules, interfering with other regulators of immune cell activity might be important to optimize the efficacy of Tcell-mediated tumor control. In this context, Shifrut and colleagues developed a screening platform in which they were able to perform single guide RNA (sgRNA) lentiviral infection combined with Cas9 protein electroporation in primary human T cells (Shifrut et al. 2018). This genome-wide loss-of-function screening revealed novel molecules that help T cells counteract cancer cells. Selected hits from this screening were knocked out in T cells engineered to express an anti-NY-ESO-1 TCR (Robbins et al. 2008). They found that knocking out TCEB2 (transcription elongation factor B (SIII), polypeptide 2), SOCS1 (suppressor of cytokine signaling 1), CBLB (Cbl proto-oncogene B), or RASA2 (RAS P21 protein activator 2) significantly enhanced T-cell proliferation and A375 tumor cell clearance (Shifrut et al. 2018). Among them, CBLB is an intracellular immune checkpoint (Hinterleitner et al. 2012). Since the depletion of either SOCS1, a primary suppressor of JAK/STAT signaling in activated T cells (Kamura et al. 1998; Liau et al. 2018), or TCEB2, a binding partner of SOCS1 (Liau et al. 2018; Kamizono et al. 2001; Ilangumaran et al. 2017), improves T-cell antitumor activity (Shifrut et al. 2018), it is reasonable to conclude that SOCS1/ TCEB2 complex precludes T-cell antitumor response. Although RASA2 is a GTPase-activating molecule that promotes the activity of wild-type RAS function (Arafeh et al. 2015), the detailed role of this protein in T-cell antitumor activity has not been described yet. Another whole-genome CRISPR screening study proposed that knocking out TLE4 (TLE family member 4, transcriptional corepressor), IKZF2 (IKAROS family zinc finger 2), EIF5A (eukaryotic translation initiation factor 5A), or TMEM184B (transmembrane protein 184B) genes improves both T-cell activation and CAR antitumor efficacy against patient-derived glioblastoma stem cells (Wang et al. 2021). Specifically, TLE4 or IKZF2 ablations resulted in cytokine upregulation and signaling activation. TLE4-depleted CAR T cells induced T-cell stimulatory and cytotoxic factors, such as IFNγ, CCL4 (C-C motif chemokine ligand 4), CCL3 (C-C motif chemokine ligand 3), and granzyme B, while IKZF2-depleted CAR T cells upregulated proinflammatory cytokines and pathways involving interactions between cytokines and their receptors (Wang et al. 2021). Corroborating these

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results, TLE4 upregulation has been reported in extensively cultured CAR T cells, a process known to induce functional exhaustion (Ghassemi et al. 2018). Another strategy to improve CAR T-cell efficiency is the knockout of key transcription factors regulators of T-cell function. This observation comes from a work that reported that upregulating c-Jun, a member of the activator protein (AP)-1 family, enhances the T-cell activity and decreases terminal differentiation (Lynn et al. 2019). Additionally, CAR T cells knocked out for all three nuclear receptor subfamily 4A (NR4A) transcription factors (NR4A1, NR4A2, and NR4A), partners of AP-1, leads to tumor regression and increased survival of tumor-bearing mice (Chen et al. 2019). Other transcription factors also have been reported to help the T cell in its battle against the tumor. The tumor-infiltrating CAR T cells knocked out for the high-mobility group (HMG)-box transcription factors TOX (thymocyte selection-associated HMG box) and TOX2 presented an increased capacity to suppress tumor growth (Seo et al. 2019). Collectively, these data highlight that the depletion of some transcription factors that regulate T-cell activation might increase CAR T-cell antitumor efficiency. Several groups dedicated efforts to investigate how to overcome the downregulation of T-cell activity through TCR signaling attenuation caused by some molecules, such as diacylglycerol kinases (DGK), a family of enzymes that phosphorylates diacylglycerol (DAG) to phosphatidic acid (PA) (Eichmann and Lass 2015). DGK interacts with essential proteins involved in TCR signal transduction, such as protein kinase C (PKC) and Ras-activating protein (RasGRP1), and this interaction results in the downregulation of TCR distal molecules, including extracellular signal-related kinases 1/2 (ERK1/2) (Quann et al. 2011; Krishna and Zhong 2013). In accordance with these biochemical interactions, disruption of DGK improved TCR-mediated signaling endowing T cells with enhanced secretion of proinflammatory cytokines, such as IL-2 and INFγ (Zhong et al. 2003), greater proliferation capacity, and resistance to anergy (Olenchock et al. 2006). Also, CRISPR/Cas9-mediated knockout DGK increased the resistance of CAR T cells to immunosuppressive factors, such as prostaglandin E2 and TGFβ, and preserved effector functions even after repeated stimulation with tumor cells. In another report, DGK-depleted CAR T cells displayed increased antitumor capacity in a glioblastoma xenograft model (Jung et al. 2018). Recently, the hematopoietic progenitor kinase 1 (HPK1) protein also came to the light in the CAR T-cell field. It is well known that HPK1 expression is restricted to the hematopoietic compartment, and this kinase is a negative regulator of TCR signaling (Shui et al. 2007; Wang et al. 2012). Initially, Alzabin and colleagues reported that T cells lacking HPK1 expression displayed increased capacity to reduce the number and size of tumor foci in mice lungs (Alzabin et al. 2010). Confirming the importance of HPK1 to the T-cell response against the tumor, other work reported that loss of HPK1 kinase activity enhanced T-cell antitumor immune responses in a GL261 glioma model (Hernandez et al. 2018a). Recently, it has been described that knocking out HPK1 in CAR T cells enhanced antitumor function in several preclinical mouse models of hematological and solid malignancies (Si et al. 2020). These findings were already replicated in the clinic since the

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treatment with anti-CD19 CAR T cells knocked out for HPK1 enhanced the number of patients that achieved complete remission or complete remission with incomplete count recovery in young adult patients with r/r B-ALL (Zhang et al. 2022b) (ClinicalTrials.gov, NCT04037566). The TGFβ-receptor II (TGFβRII) also became a target for modulating T-cell therapeutic efficacy. It has been reported that knocking out TGFβRII might increase CAR T-cell activity by directly restricting the immunosuppressive effects of TGFβ in the tumor microenvironment. This could circumvent the side effects associated with systemic administration of TGFβ antagonists (Vong et al. 2017). Indeed, T cells engineered to express the TGFβRII-dominant negative receptor have better tumor eradication properties in a B16 melanoma model (Zhang et al. 2012) and in aggressive human prostate cancer mouse models (Kloss et al. 2018). Consistently, CRISPR/Cas9-based knockout of TGFβRII increases the capacity of CAR T cells to eliminate the tumors in cell line-derived and patient-derived xenograft models of solid tumors (Tang et al. 2020). Deletion of TGFβRII was also shown to diminish T-cell exhaustion after multiple rounds of antigen stimulation (Alishah et al. 2021) and rendered ovarian cancer-derived tumor-infiltrating lymphocytes (TILs) resistant to TGFβ-mediated immunosuppression (Fix et al. 2022). Another molecule shown to modulate T-cell therapeutic efficacy was the ribonuclease REGNASE-1 (regulatory RNase 1) (Akira 2013). Wei and colleagues reported that the depletion of REGNASE-1 mediated by CRISPR/Cas9 in CD8+ CAR T cells enhanced their antitumor function in leukemia and melanoma mouse models (Wei et al. 2019). They demonstrated that the main target of REGNASE-1 is the mRNA-encoding BATF (basic leucine zipper ATF-like transcription factor), which is a key transcription factor involved in the differentiation of effector T cells (Quigley et al. 2010; Kurachi et al. 2014; Kuroda et al. 2011). Ablation of REGNASE-1 resulted in the accumulation of BATF which in turn promoted increased expansion and mitochondrial fitness of CD8+ T cells, leading to a better antitumor function (Wei et al. 2019). Later, the same group reported that REGNASE-1 deficiency enhances CAR T-cell antitumor toxicity in a xenograft model of B-ALL (Zheng et al. 2021). Additionally, knocking out REGNASE-1 promotes T-cell factor 1 (TCF-1) expression, increasing CAR T-cell expansion and memory-like cell formation (Zheng et al. 2021). Further reinforcing these observations, Behrens and colleagues demonstrated that weakening the interaction of ROQUIN-1 with REGNASE-1 improves the antitumor response of cytotoxic T cells in a B16-OVA melanoma model (Behrens et al. 2021). Despite the successful clinical application of CAR T-cell therapy and remission rates in hematologic malignancies, cytokine release syndrome (CRS) and potential toxicities, such as neurotoxicity, are limiting factors. The in vivo expansion of CAR T cells and the massive release of cytokines such as IFNγ, IL-6, monocyte chemoattractant protein 1 (MCP-1), and granulocyte-macrophage colony-stimulating factor (GM-CSF), are considered important triggers of CRS (Neelapu et al. 2017; Maude et al. 2018; Teachey et al. 2016; Park et al. 2018). Recent data suggest that monocytes and macrophages contribute to the development of CRS and neurotoxicity after CAR T-cell infusion. More specifically, GM-CSF serum levels were

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identified as one of the most significant markers of grade 3–4 neurotoxicity (Neelapu et al. 2017). Accordingly, grade 3–4 neurotoxicity correlates with increased CD14+ monocytes in the cerebrospinal fluid of patients (Locke et al. 2017), suggesting the potential role of this CM-CSF and myeloid cells in CRS and neurotoxicity. Considering this, Sterner and colleagues investigated the neutralization of GM-CSF as a way to manage CAR T-cell toxicities. First, they neutralized GM-CSF through lenzilumab, resulting in enhanced CAR T-cell proliferation and reduction of myeloid and T-cell infiltration in the central nervous system in a patient-derived xenograft model of B-AAL. After that, GM-CSF-deficient anti-CD19 CAR T cells were generated through CRISPR/Cas9 and demonstrated enhanced antitumor activity in vivo compared to unedited CAR T cells, which reflected in a better mice survival (Sterner et al. 2019). These findings bring up the disruption of GM-CSF to counteract CRS and neurotoxicity associated with CAR T-cell therapy.

4 Genome Editing Strategies to Make an Allogeneic CAR T-Cell Therapy Feasible The traditional manufacturing process of CAR T-cell therapies relies on patientderived T-cell modification to express the CAR transgene (Levine et al. 2016). Despite the impressive results achieved with such personalized (autologous) products, this way of production limits the number of patients who can potentially benefit from the therapy. In the phase 2 trial of the tisagenlecleucel (tisa-cel) CAR T-cell product, 17 out of the 92 patients (18.5%) could not be treated due to manufacturing and disease-related issues (Maude et al. 2018). Advanced disease and previous cancer treatments influence the quantity and quality of patients’ T cells, which can impair the manufacturing process. For example, repeated cycles of chemotherapy in pediatric patients contribute to a deficit in less differentiated T-cell subpopulations (naïve and stem central memory phenotypes), which is associated with a poor expansion potential and higher risks of production failure (Singh et al. 2016; Das et al. 2019). The cell subpopulations into the starting material and subsequent CAR T-cell product are also important indicators for clinical activity (Fraietta et al. 2018b; Deng et al. 2020). Another factor that hurdles autologous therapy is the time needed for the production, which is not feasible for some patients. In a comprehensive review of CAR T-cell therapy application in the real world, Westin and colleagues found vein-tovein time, starting in the cell collection until the infusion, ranging from 21 to 44 days for axicabtagene ciloleucel (axi-cel) and tisa-cel CAR T-cell products (Westin et al. 2021). This prolonged time might put the patients at risk for disease progression, making them no longer eligible for the therapy (Schuster et al. 2019; Wang et al. 2020). Additionally, the therapy production for each patient increases the cost once manufacturing scale-up is not possible. The expected cost for the treatment with tisa-

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cel and axi-cel is between $400,000 and $500,000 per patient (Hernandez et al. 2018b), which hinders autologous therapy accessibility. A promising solution to overcome those limitations is to use healthy donors’ T cells as a source to produce an allogeneic therapy. This approach can increase the number of doses obtained from a single production batch, offering a ready-to-use product for several patients and potentially reducing costs through manufacturing scale-up (Blache et al. 2022). To face the barriers related to allogeneic interactions, genome editing techniques have been applied for “nonimmunogenic” T-cell development, preventing immune rejection in both graft-versus-host and host-versus-graft directions. GvHD is a common complication post allogeneic transplant. This condition occurs when the donor’s TCR recognizes host antigens and establishes an immune response against the cells harboring them, with subsequent impact in the corresponding tissues (Marino et al. 2016). To address safety concerns related to GvHD, many preclinical studies explored TCR disruption in CAR T cells by knocking out genes encoding the constant regions of TCR α- and β-chains using ZFN, TALEN, and CRISPR/Cas9-based methods (Jo et al. 2022; Eyquem et al. 2017; Ren et al. 2017a; Torikai et al. 2012; Poirot et al. 2015). αβ heterodimer is required for full assembly and activity of the TCR complex, so gene disruption of just one of those chains is enough to deplete the receptor. In contrast to the two genes that encode the TCRβ chain, the α chain is encoded by a single TRAC gene, making it the preferred target for disruption (Osborn et al. 2016). In the first clinical translation of this approach in a special access setting, two B-ALL pediatric patients were treated with TALEN-edited allogeneic (“universal”) CAR T cells (UCART19) (Qasim et al. 2017). mRNA encoding TALENs were introduced into allogeneic cells to mediate TCR and CD52 disruptions, minimizing the risk of GvHD and conferring resistance to the lymphodepleting agent alemtuzumab. Both patients achieved complete remission with mild or no occurrence of GvHD and persisted disease-free 4 years later (Benjamin et al. 2020). In more comprehensive clinical studies, 7 children (ClinicalTrials.gov, NCT02808442) and 14 adults (ClinicalTrials.gov, NCT02746952) with r/r B-ALL were treated with UCART19 (Benjamin et al. 2020). The safety profile of this therapy was highlighted since only two patients (10%) experienced a mild GvHD. Also, allogeneic CAR T cells had a potent antileukemia effect, resulting in high complete response rates. However, the short persistence and the expansion conditioned to alemtuzumabmediated lymphodepletion were both suggestive indicators of host-mediated rejection of allogeneic CAR T cells (Benjamin et al. 2020). To overcome graft rejection and improve persistence, some studies have explored HLA-I depletion in allogeneic CAR T cells to prevent their recognition by patients’ CD8+ T cells. By using the CRISPR/Cas9 platform for multiplex genome editing, Ren and colleagues produced TCR- and HLA-I-depleted anti-CD19 CAR T cells (Ren et al. 2017b). Edited CAR T cells had an expressive antitumor activity in vitro and in a B-cell leukemia murine model, which was comparable to unedited cells. Despite the success in GvHD abrogation in vivo by knocking out the TRAC gene from CAR T cells, HLA-I depletion through β2-microglobulin (B2M) gene

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disruption was not enough to eliminate graft rejection by allogeneic immune cells in vitro (Ren et al. 2017b). Allorejection was detected even when both HLA-I and HLA-II molecules were depleted from activated CAR T-cell surface by B2M and CIITA (class II major histocompatibility complex transactivator) genes disruption (Kagoya et al. 2020). In two patients with r/r diffuse large B-cell lymphoma, an increase in NK cell number in peripheral blood followed by a decrease in the number of TCR/HLA-I-depleted CAR T cells was observed (Guo et al. 2022). This response can be attributed to the capacity of NK cells to recognize and lyse HLA-negative cells (Moretta et al. 1996), which potentially compromise long-term allogeneic CAR T cell persistence in vivo. A recently demonstrated way to inhibit NK cell-mediated cytotoxicity is the expression of the non-polymorphic HLA-E molecule on CAR T cells. Jo and coworkers described the TRAC and B2M disruption through CAR and HLA-Etargeted insertion, respectively, using a TALEN-based methodology (Jo et al. 2022). Double insertion rates were as high as 68%, with no significant observation of off-targets and an expected level of translocation between TRAC and B2M loci. Double-edited cells maintained an antitumor effect comparable to TCR-disrupted CAR T cells both in vitro and in a B-cell leukemia murine model. The HLA-I depletion on CAR T cells successfully prevented their rejection by CD8+ T cells; at the same time, HLA-E expression inhibited NK cell cytolytic activity, with a demonstrated prolonged antitumor activity in a co-challenge with tumor and NK cells (Jo et al. 2022). Despite this effective strategy to overcome persistence issues of allogeneic CAR T cells, a simpler solution is redosing, which is facilitated by the offthe-shelf nature of this therapy. Allogeneic therapies are a growing area of research, which provide a safe alternative to face the challenges of the traditional way of CAR T-cell manufacture (Chen et al. 2022). Besides anti-CD19 CAR, other CAR constructs have been tested in a clinical setting for different tumor targets. A brief overview of these studies was described by Lin and colleagues (Lin et al. 2021).

5 Genome Editing for Generating Fratricide-Resistant Engineered T Cells T-cell malignancies comprise a group of aggressive diseases whose therapeutic options have limited efficacy, resulting in high rates of relapse and mortality for both adult and pediatric patients (Ma and Abdul-Hay 2017; Karrman and Johansson 2017). CAR T-cell therapy for this type of hematological malignancy has a limiting factor: the shared expression of target antigens between normal T effector and malignant T cells. Fratricide is a natural and important mechanism for T-cell homeostasis maintenance (Callard et al. 2003). However, in the context of CAR T-cell therapy against T-cell malignancies, targeting shared proteins such as CD7 and CD3 impedes CAR T-cell expansion and persistence as a consequence of self-

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killing. This observation is also true for immunotherapies using T cells expressing artificial TCRs against survivin, an apoptose inhibitor overexpressed by many tumors and also expressed by T cells (Leisegang et al. 2010). Genome editing tools have been applied as an effective solution for the removal of T-cell surface markers in CAR T cells, thereby sparing CAR T cells from self-killing and redirecting their function against tumor T cells. The transmembrane glycoprotein CD7 is expressed by most T and natural killer (NK) cells, and its expression is also detected in approximately 95% of T-cell leukemias and lymphomas (Scherer et al. 2019). It was previously observed that T cells encoding anti-CD7 CAR underwent fratricide, which hampered their expansion after modification (Cooper et al. 2018; Png et al. 2017; Gomes-Silva et al. 2017). To avoid this phenomenon, CD7-disrupted anti-CD7 CAR T cells were produced using CRISPR/Cas9 (Gomes-Silva et al. 2017). A high CD7 knockout rate was observed, resulting in >80% CD7-negative CAR T cells by the end of the manufacturing protocol. These cells had improved cell viability, robust expansion, and potent cytotoxic activity against CD7+ immortalized and primary tumor cells compared to the unedited CAR T cells. Although CD7 has an important role in T-cell costimulation, its depletion did not compromise the functional activity of CAR T cells (Gomes-Silva et al. 2017). Combined, these data showed that the depletion of CD7 can overcome fratricide in CAR T cells, making CAR T-cell therapy feasible against T-cell malignancies. Using a multiplex CRISPR/Cas9-editing approach to abrogate CD7 and TRAC expressions, Cooper and collaborators produced allogeneic anti-CD7 CAR T cells resistant to fratricide (Cooper et al. 2018). These cells had a robust antitumor activity in vivo and did not induce GvHD, revealing a promising strategy to produce a CAR T-cell therapy from third-party donors for the treatment of patients with T-cell malignancies who have a risk of malignant cell contamination in the starting material for CAR T-cell production (Ruella et al. 2018). However, CRISPR/Cas9-mediated targeting of multiple genes was associated with increased observations of aneuploidies and chromosome truncations, even when a single gene was targeted (Nahmad et al. 2022). Base editors (BE) have shown an encouraging alternative to genome editing approaches that rely on DSB induction. The third generation of BE (BE3) technology comprises a Cas9 nickase fused to the deaminase APOBEC1 and a single uracil glycosylase inhibitor for inducing cytidine to uridine conversion by deamination, leading to the final substitution of cytosine for thymine (Komor et al. 2016). Since the Cas9 specificity is supported by the gRNA base complementarity to the target site, the modification occurs in a precise way, producing stop codons or disrupted splice donor/acceptor sites. Georgiadis and coworkers explored BE3 technology to produce anti-CD7 and anti-CD3 CAR T cells in a preclinical study (Georgiadis et al. 2021). The achieved TCR β-chain and CD7 double disruption were similar between BE3 and Cas9-based editing approaches, with the former having the advantage of inducing lower levels of translocation. Coculture with edited anti-CD3 and anti-CD7 showed an enrichment of the corresponding depleted cells, indicating both a fratricide-resistance property, due to the maintenance of negative cells, and

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elimination of residual unedited cells. BE3-edited cells also had an antitumor potency similar to the Cas9-edited counterparts in vitro, and the combination of the anti-CD7 and anti-CD3 CAR T cells in vivo resulted in the best antitumoral effect against T-cell acute lymphoblastic leukemia (T-ALL) cells (Georgiadis et al. 2021). These results restate the possibility of using advanced genome editing tools to overcome challenges that still arise in CAR T-cell therapy, such as the fratricideassociated limitation for therapies against T-cell malignancies.

6 Conclusion The adoptive transfer of genetically modified T cells expressing artificial tumorspecific receptors such as CARs or engineered TCRs has resulted in outstanding remission rates in relapsed/refractory B-cell malignancies. The next frontiers of the field are to extend these results to solid tumors, which constitute the vast majority of neoplasms, and to improve access to these therapies. In this chapter, we described how genome editing has been applied to unlock the full potential of these therapies through genome-wide functional screens, editing of single or multiple genes during manufacturing, and targeted insertions in engineered T-cell products (Fig. 6.1). This huge effort has resulted in a wide plethora of solutions that paved the way to increase the therapeutic efficacy of engineered T cells, expand the repertoire of treatable diseases with this technology, and develop the next generation of an allogeneic “offthe-shelf” advanced cell therapy with the potential to maximize patient accessibility.

References Akira S (2013) Regnase-1, a ribonuclease involved in the regulation of immune responses. Cold Spring Harb Symp Quant Biol [Internet] 78(1):51–60. Available from: https://pubmed.ncbi.nlm. nih.gov/24163394/ [cited 2022 Sep 16] Alishah K, Birtel M, Masoumi E, Jafarzadeh L, Mirzaee HR, Hadjati J et al (2021) CRISPR/Cas9mediated TGFβRII disruption enhances anti-tumor efficacy of human chimeric antigen receptor T cells in vitro. J Transl Med [Internet] 19(1). Available from: https://pubmed.ncbi.nlm.nih. gov/34838059/ [cited 2022 Sep 16] Alzabin S, Pyarajan S, Yee H, Kiefer F, Suzuki A, Burakoff S et al (2010) Hematopoietic progenitor kinase 1 is a critical component of prostaglandin E2-mediated suppression of the anti-tumor immune response. Cancer Immunol Immunother [Internet] 59(3):419–429. Available from: https://pubmed.ncbi.nlm.nih.gov/19787351/ [cited 2022 Sep 16] Anzalone AV, Koblan LW, Liu DR (2020) Genome editing with CRISPR–Cas nucleases, base editors, transposases and prime editors. Nat Biotechnol [Internet] 38(7):824–844. Available from: https://www.nature.com/articles/s41587-020-0561-9 [cited 2022 Sep 16] Arafeh R, Qutob N, Emmanuel R, Keren-Paz A, Madore J, Elkahloun A et al (2015) Recurrent inactivating RASA2 mutations in melanoma. Nat Genet [Internet] 47(12):1408–1410. Available from: https://pubmed.ncbi.nlm.nih.gov/26502337/ [cited 2022 Sep 16] Behrens G, Edelmann SL, Raj T, Kronbeck N, Monecke T, Davydova E et al (2021) Disrupting Roquin-1 interaction with Regnase-1 induces autoimmunity and enhances antitumor responses.

6

Genome Editing for Engineering the Next Generation of Advanced Immune. . .

101

Nat Immunol [Internet] 22(12):1563–1576. Available from: https://pubmed.ncbi.nlm.nih. gov/34811541/ [cited 2022 Sep 16] Benjamin R, Graham C, Yallop D, Jozwik A, Mirci-Danicar OC, Lucchini G et al (2020) Genomeedited, donor-derived allogeneic anti-CD19 chimeric antigen receptor T cells in paediatric and adult B-cell acute lymphoblastic leukaemia: results of two phase 1 studies. The Lancet [Internet] 396(10266):1885–1894. Available from: http://www.thelancet.com/article/S014067362032334 5/fulltext [cited 2022 Sep 16] Blache U, Popp G, Dünkel A, Koehl U, Fricke S (2022) Potential solutions for manufacture of CAR T cells in cancer immunotherapy. Nat Commun [Internet] 13(1):5225. Available from: https:// pubmed.ncbi.nlm.nih.gov/36064867/ [cited 2022 Sep 16] Callard RE, Stark J, Yates AJ (2003) Fratricide: a mechanism for T memory-cell homeostasis. Trends Immunol 24(7):370–375 CAR T Cells: Engineering immune cells to treat cancer – NCI [Internet]. Available from: https:// www.cancer.gov/about-cancer/treatment/research/car-t-cells [cited 2022 Sep 16] Ceppi F, Rivers J, Annesley C, Pinto N, Park JR, Lindgren C et al (2018) Lymphocyte apheresis for chimeric antigen receptor T-cell manufacturing in children and young adults with leukemia and neuroblastoma. Transfusion (Paris) [Internet] 58(6):1414–1420. Available from: https:// onlinelibrary.wiley.com/doi/full/10.1111/trf.14569 [cited 2022 Sep 16] Chen J, López-Moyado IF, Seo H, Lio CWJ, Hempleman LJ, Sekiya T et al (2019) NR4A transcription factors limit CAR T cell function in solid tumours. Nature [Internet] 567(7749): 530–534. Available from: https://pubmed.ncbi.nlm.nih.gov/30814732/ [cited 2022 Sep 16] Chen S, Zhang Y, Fang C, Zhang N, Wang Y, Chen R et al (2022) Donor-derived and off-the-shelf allogeneic anti-CD19 CAR T-cell therapy for R/R ALL and NHL: a systematic review and meta-analysis. Crit Rev Oncol Hematol [Internet] 103807. Available from: https://pubmed.ncbi. nlm.nih.gov/36087853/ [cited 2022 Sep 16] Cho SW, Kim S, Kim JM, Kim JS (2013) Targeted genome engineering in human cells with the Cas9 RNA-guided endonuclease. Nat Biotechnol [Internet] 31(3):230–232. Available from: https://www.nature.com/articles/nbt.2507 [cited 2022 Sep 16] Cooper ML, Choi J, Staser K, Ritchey JK, Devenport JM, Eckardt K et al (2018) An “off-the-shelf” fratricide-resistant CAR-T for the treatment of T cell hematologic malignancies. Leukemia [Internet] 32(9):1970–1983. Available from: https://www.nature.com/articles/s41375-018-00 65-5 [cited 2022 Sep 16] Couzin-Frankel J (2013) Breakthrough of the year 2013. Cancer immunotherapy. Science [Internet] 342(6165):1432–1433. Available from: https://pubmed.ncbi.nlm.nih.gov/24357284/ [cited 2022 Sep 16] Dai X, Park JJ, Du Y, Kim HR, Wang G, Errami Y et al (2019) One-step generation of modular CAR-T cells with AAV–Cpf1. Nat Methods [Internet] 16(3):247–254. Available from: https:// www.nature.com/articles/s41592-019-0329-7 [cited 2022 Sep 16] O'Driscoll M, Jeggo PA. (2006) The role of double-strand break repair – insights from human genetics. Nat Rev Genet 7(1):45–54. https://doi.org/10.1038/nrg1746 Das RK, Vernau L, Grupp SA, Barrett DM (2019) Naïve T-cell deficits at diagnosis and after chemotherapy impair cell therapy potential in Pediatric cancers. Cancer Discov [Internet] 9(4): 492–499. Available from: https://pubmed.ncbi.nlm.nih.gov/30630850/ [cited 2022 Sep 16] Deng Q, Han G, Puebla-Osorio N, Ma MCJ, Strati P, Chasen B et al (2020) Characteristics of antiCD19 CAR T cell infusion products associated with efficacy and toxicity in patients with large B cell lymphomas. Nat Med [Internet] 26(12):1878–1887. Available from: https://pubmed.ncbi. nlm.nih.gov/33020644/ [cited 2022 Sep 16] Depil S, Duchateau P, Grupp SA, Mufti G, Poirot L (2020) ‘Off-the-shelf’ allogeneic CAR T cells: development and challenges. Nat Rev Drug Discov [Internet] 19(3):185–199. Available from: https://www.nature.com/articles/s41573-019-0051-2 [cited 2022 Sep 16] Dobosz P, Dzieciątkowski T (2019) The intriguing history of cancer immunotherapy. Front Immunol 10:2965

102

S. C. G. de Lima et al.

Dossett ML, Teague RM, Schmitt TM, Tan X, Cooper LJN, Pinzon C et al (2009) Adoptive immunotherapy of disseminated leukemia with TCR-transduced, CD8+ T cells expressing a known endogenous TCR. Mol Ther [Internet] 17(4):742–749. Available from: https://pubmed. ncbi.nlm.nih.gov/19209146/ [cited 2022 Sep 16] Eichmann TO, Lass A (2015) DAG tales: the multiple faces of diacylglycerol–stereochemistry, metabolism, and signaling. Cell Mol Life Sci [Internet] 72(20):3931–3952. Available from: https://pubmed.ncbi.nlm.nih.gov/26153463/ [cited 2022 Sep 16] Eyquem J, Mansilla-Soto J, Giavridis T, van der Stegen SJC, Hamieh M, Cunanan KM et al (2017) Targeting a CAR to the TRAC locus with CRISPR/Cas9 enhances tumour rejection. Nature [Internet] 543(7643):113–117. Available from: https://www.nature.com/articles/nature21405 [cited 2022 Sep 16] Fix SM, Forget MA, Sakellariou-Thompson D, Wang Y, Griffiths TM, Lee M et al (2022) CRISPRmediated TGFBR2 knockout renders human ovarian cancer tumor-infiltrating lymphocytes resistant to TGF-β signaling. J Immunother Cancer [Internet] 10(7). Available from: https:// pubmed.ncbi.nlm.nih.gov/35882447/ [cited 2022 Sep 16] Fraietta JA, Nobles CL, Sammons MA, Lundh S, Carty SA, Reich TJ et al (2018a) Disruption of TET2 promotes the therapeutic efficacy of CD19-targeted T cells. Nature [Internet] 558(7709): 307–312. Available from: https://www.nature.com/articles/s41586-018-0178-z [cited 2022 Sep 16] Fraietta JA, Lacey SF, Orlando EJ, Pruteanu-Malinici I, Gohil M, Lundh S et al (2018b) Determinants of response and resistance to CD19 chimeric antigen receptor (CAR) T cell therapy of chronic lymphocytic leukemia. Nat Med [Internet] 24(5):563–571. Available from: https:// pubmed.ncbi.nlm.nih.gov/29713085/ [cited 2022 Sep 16] Franzin R, Netti GS, Spadaccino F, Porta C, Gesualdo L, Stallone G et al (2020) The use of immune checkpoint inhibitors in oncology and the occurrence of AKI: where do we stand? Front Immunol 11:2619 Gándara C, Affleck V, Stoll EA (2018) Manufacture of third-generation lentivirus for preclinical use, with process development considerations for translation to Good manufacturing practice. Hum Gene Ther Methods [Internet] 29(1):1–15. Available from: https://pubmed.ncbi.nlm.nih. gov/29212357/ [cited 2022 Sep 16] Gaudelli NM, Komor AC, Rees HA, Packer MS, Badran AH, Bryson DI et al (2017) Programmable base editing of a•T to G•C in genomic DNA without DNA cleavage. Nature [Internet] 551(7681):464–471. Available from: https://www.nature.com/articles/nature24644 [cited 2022 Sep 16] Georgiadis C, Rasaiyaah J, Gkazi SA, Preece R, Etuk A, Christi A et al (2021) Base-edited CAR T cells for combinational therapy against T cell malignancies. Leukemia [Internet] 35(12): 3466–3481. Available from: https://pubmed.ncbi.nlm.nih.gov/34035409/ [cited 2022 Sep 16] Ghassemi S, Nunez-Cruz S, O’Connor RS, Fraietta JA, Patel PR, Scholler J et al (2018) Reducing ex vivo culture improves the Antileukemic activity of chimeric antigen receptor (CAR) T cells. Cancer Immunol Res [Internet] 6(9):1100–1109. Available from: https://pubmed.ncbi.nlm.nih. gov/30030295/ [cited 2022 Sep 16] Giuffrida L, Sek K, Henderson MA, Lai J, Chen AXY, Meyran D et al (2021) CRISPR/Cas9 mediated deletion of the adenosine A2A receptor enhances CAR T cell efficacy. Nat Commun [Internet] 12(1):1–18. Available from: https://www.nature.com/articles/s41467-021-23331-5 [cited 2022 Sep 16] Gomes-Silva D, Srinivasan M, Sharma S, Lee CM, Wagner DL, Davis TH et al (2017) CD7-edited T cells expressing a CD7-specific CAR for the therapy of T-cell malignancies. Blood [Internet] 130(3):285–296. Available from: https://pubmed.ncbi.nlm.nih.gov/28539325/ [cited 2022 Sep 16] Guo X, Jiang H, Shi B, Zhou M, Zhang H, Shi Z et al (2018) Disruption of PD-1 enhanced the antitumor activity of chimeric antigen receptor T cells against hepatocellular carcinoma. Front Pharmacol 9:1118

6

Genome Editing for Engineering the Next Generation of Advanced Immune. . .

103

Guo Y, Tong C, Su L, Zhang W, Jia H, Liu Y et al (2022) CRISPR/Cas9 genome-edited universal CAR T cells in patients with relapsed/refractory lymphoma. Blood Adv [Internet] 6(8): 2695–2699. Available from: https://pubmed.ncbi.nlm.nih.gov/35008103/ [cited 2022 Sep 16] Hacein-Bey-Abina S, Garrigue A, Wang GP, Soulier J, Lim A, Morillon E et al (2008) Insertional oncogenesis in 4 patients after retrovirus-mediated gene therapy of SCID-X1. J Clin Invest [Internet] 118(9):3132–3142. Available from: http://www.jci.org [cited 2022 Sep 16] Hale M, Lee B, Honaker Y, Leung WH, Grier AE, Jacobs HM et al (2017) Homology-directed recombination for enhanced engineering of chimeric antigen receptor T cells. Mol Ther Methods Clin Dev [Internet] 4:192–203. Available from: https://pubmed.ncbi.nlm.nih.gov/2 8345004/ [cited 2022 Sep 16] Hernandez S, Qing J, Thibodeau RH, Du X, Park S, Lee HM et al (2018a) The kinase activity of hematopoietic progenitor kinase 1 is essential for the regulation of T cell function. Cell Rep [Internet] 25(1):80–94. Available from: https://pubmed.ncbi.nlm.nih.gov/30282040/ [cited 2022 Sep 16] Hernandez I, Prasad V, Gellad WF (2018b) Total costs of chimeric antigen receptor T-cell immunotherapy. JAMA Oncol [Internet] 4(7):994–996. Available from: https://pubmed.ncbi. nlm.nih.gov/29710129/ [cited 2022 Sep 16] Hinterleitner R, Gruber T, Pfeifhofer-Obermair C, Lutz-Nicoladoni C, Tzankov A, Schuster M et al (2012) Adoptive transfer of siRNA Cblb-silenced CD8+ T lymphocytes augments tumor vaccine efficacy in a B16 melanoma model. PLoS One [Internet] 7(9). Available from: https://pubmed.ncbi.nlm.nih.gov/22962608/ [cited 2022 Sep 16] Hou AJ, Chen LC, Chen YY (2021) Navigating CAR-T cells through the solid-tumour microenvironment. Nat Rev Drug Discov [Internet] 20(7):531–550. Available from: https://www. nature.com/articles/s41573-021-00189-2 [cited 2022 Sep 16] Hu W, Zi Z, Jin Y, Li G, Shao K, Cai Q et al (2019a) CRISPR/Cas9-mediated PD-1 disruption enhances human mesothelin-targeted CAR T cell effector functions. Cancer Immunol Immunother [Internet] 68(3):365–377. Available from: https://pubmed.ncbi.nlm.nih.gov/30 523370/ [cited 2022 Sep 16] Hu B, Zou Y, Zhang L, Tang J, Niedermann G, Firat E et al (2019b) Nucleofection with plasmid DNA for CRISPR/Cas9-mediated inactivation of programmed cell death protein 1 in CD133specific CAR T cells. Hum Gene Ther [Internet] 30(4):446–458. Available from: https:// pubmed.ncbi.nlm.nih.gov/29706119/ [cited 2022 Sep 16] Ilangumaran S, Bobbala D, Ramanathan S (2017) SOCS1: regulator of T cells in autoimmunity and cancer. Curr Top Microbiol Immunol [Internet] 410:159–189. Available from: https://link. springer.com/chapter/10.1007/82_2017_63 [cited 2022 Sep 17] Jayaraman J, Mellody MP, Hou AJ, Desai RP, Fung AW, Pham AHT et al (2020) CAR-T design: elements and their synergistic function. EBioMedicine [Internet]:58. Available from: https:// pubmed.ncbi.nlm.nih.gov/32739874/ [cited 2022 Sep 16] Jo S, Das S, Williams A, Chretien AS, Pagliardini T, le Roy A et al (2022) Endowing universal CAR T-cell with immune-evasive properties using TALEN-gene editing. Nat Commun [Internet] 13(1). Available from: https://pubmed.ncbi.nlm.nih.gov/35773273/ [cited 2022 Sep 16] Johnson DB, Nebhan CA, Moslehi JJ, Balko JM (2022) Immune-checkpoint inhibitors: long-term implications of toxicity. Nat Rev Clin Oncol [Internet] 19(4):254–267. Available from: https:// www.nature.com/articles/s41571-022-00600-w [cited 2022 Sep 16] Joung JK, Sander JD (2012) TALENs: a widely applicable technology for targeted genome editing. Nat Rev Mol Cell Biol [Internet] 14(1):49–55. Available from: https://www.nature.com/articles/ nrm3486 [cited 2022 Sep 16] Jung IY, Kim YY, Yu HS, Lee M, Kim S, Lee J (2018) CRISPR/Cas9-mediated knockout of DGK improves antitumor activities of human T cells. Cancer Res [Internet] 78(16):4692–4703. Available from: https://pubmed.ncbi.nlm.nih.gov/29967261/ [cited 2022 Sep 16] Kagoya Y, Guo T, Yeung B, Saso K, Anczurowski M, Wang CH et al (2020) Genetic ablation of HLA class I, class II, and the T-cell receptor enables allogeneic T cells to be used for adoptive T-cell therapy. Cancer Immunol Res [Internet] 8(7):926–936. Available from: https://pubmed. ncbi.nlm.nih.gov/32321775/ [cited 2022 Sep 16]

104

S. C. G. de Lima et al.

Kamizono S, Hanada T, Yasukawa H, Minoguchi S, Kato R, Minoguchi M et al (2001) The SOCS box of SOCS-1 accelerates ubiquitin-dependent proteolysis of TEL-JAK2. J Biol Chem [Internet] 276(16):12530–12538. Available from: https://pubmed.ncbi.nlm.nih.gov/11278610/ [cited 2022 Sep 17] Kamura T, Sato S, Haque D, Liu L, Kaelin WG, Conaway RC et al (1998) The Elongin BC complex interacts with the conserved SOCS-box motif present in members of the SOCS, ras, WD-40 repeat, and ankyrin repeat families. Genes Dev [Internet] 12(24):3872–3881. Available from: http://genesdev.cshlp.org/content/12/24/3872.full [cited 2022 Sep 17] Kankeu Fonkoua LA, Sirpilla O, Sakemura R, Siegler EL, Kenderian SS (2022) CAR T cell therapy and the tumor microenvironment: current challenges and opportunities. Mol Ther Oncolytics [Internet] 25:69–77. Available from: https://pubmed.ncbi.nlm.nih.gov/35434273/ [cited 2022 Sep 16] Karrman K, Johansson B (2017) Pediatric T-cell acute lymphoblastic leukemia. Genes Chromosomes Cancer [Internet] 56(2):89–116. Available from: https://pubmed.ncbi.nlm.nih.gov/2763 6224/ [cited 2022 Sep 16] Kath J, Du W, Pruene A, Braun T, Thommandru B, Turk R et al (2022) Pharmacological interventions enhance virus-free generation of TRAC-replaced CAR T cells. Mol Ther Methods Clin Dev [Internet] 25:311–330. Available from: https://pubmed.ncbi.nlm.nih.gov/35573047/ [cited 2022 Sep 16] Kleinstiver BP, Tsai SQ, Prew MS, Nguyen NT, Welch MM, Lopez JM et al (2016) Genome-wide specificities of CRISPR-Cas Cpf1 nucleases in human cells. Nat Biotechnol [Internet] 34(8): 869–874. Available from: https://www.nature.com/articles/nbt.3620 [cited 2022 Sep 16] Kloss CC, Lee J, Zhang A, Chen F, Melenhorst JJ, Lacey SF et al (2018) Dominant-negative TGF-β receptor enhances PSMA-targeted human CAR T cell proliferation and augments prostate cancer eradication. Mol Ther [Internet] 26(7):1855–1866. Available from: https://pubmed. ncbi.nlm.nih.gov/29807781/ [cited 2022 Sep 16] Komor AC, Kim YB, Packer MS, Zuris JA, Liu DR (2016) Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature [Internet] 533(7603):420–424. Available from: https://www.nature.com/articles/nature17946 [cited 2022 Sep 16] Krishna S, Zhong XP (2013) Role of diacylglycerol kinases in T cell development and function. Crit Rev Immunol [Internet] 33(2):97–118. Available from: https://pubmed.ncbi.nlm.nih. gov/23582058/ [cited 2022 Sep 16] Kurachi M, Barnitz RA, Yosef N, Odorizzi PM, Diiorio MA, Lemieux ME et al (2014) The transcription factor BATF operates as an essential differentiation checkpoint in early effector CD8+ T cells. Nat Immunol [Internet] 15(4):373–383. Available from: https://pubmed.ncbi. nlm.nih.gov/24584090/ [cited 2022 Sep 16] Kuroda S, Yamazaki M, Abe M, Sakimura K, Takayanagi H, Iwai Y (2011) Basic leucine zipper transcription factor, ATF-like (BATF) regulates epigenetically and energetically effector CD8 T-cell differentiation via Sirt1 expression. Proc Natl Acad Sci U S A [Internet] 108(36): 14885–14889. Available from: https://pubmed.ncbi.nlm.nih.gov/21873234/ [cited 2022 Sep 16] Labbé RP, Vessillier S, Rafiq QA (2021) Lentiviral vectors for T cell engineering: clinical applications, bioprocessing and future perspectives. Viruses [Internet] 13(8):1528. Available from: https://pubmed.ncbi.nlm.nih.gov/34452392/ [cited 2022 Sep 16] Ledford H (2011) Targeted gene editing enters clinic. Nature 471(7336):16 Levine BL, Miskin J, Wonnacott K, Keir C (2016) Global manufacturing of CAR T cell therapy. Mol Ther Methods Clin Dev [Internet] 4:92–101. Available from: https://pubmed.ncbi.nlm.nih. gov/28344995/ [cited 2022 Sep 16] Li H, Yang Y, Hong W, Huang M, Wu M, Zhao X (2020) Applications of genome editing technology in the targeted therapy of human diseases: mechanisms, advances and prospects. Signal Transduct Target Ther [Internet] 5(1):1–23. Available from: https://www.nature.com/ articles/s41392-019-0089-y [cited 2022 Sep 16]

6

Genome Editing for Engineering the Next Generation of Advanced Immune. . .

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Liau NPD, Laktyushin A, Lucet IS, Murphy JM, Yao S, Whitlock E et al (2018) The molecular basis of JAK/STAT inhibition by SOCS1. Nat Commun [Internet] 9:1–14. Available from: https://www.nature.com/articles/s41467-018-04013-1 [cited 2022 Sep 17] Lin H, Cheng J, Mu W, Zhou J, Zhu L (2021) Advances in universal CAR-T cell therapy. Front Immunol [Internet]:12. Available from: https://pubmed.ncbi.nlm.nih.gov/34691052/ [cited 2022 Sep 16] Locke FL, Neelapu SS, Bartlett NL, Lekakis LJ, Jacobson CA, Braunschweig I et al (2017) Preliminary results of prophylactic tocilizumab after axicabtageneciloleucel (axi-cel; KTE-C19) treatment for patients with refractory, aggressive Non-Hodgkin Lymphoma (NHL). Blood 130(Suppl_1):1547 Lynn RC, Weber EW, Sotillo E, Gennert D, Xu P, Good Z et al (2019) c-Jun overexpression in CAR T cells induces exhaustion resistance. Nature [Internet] 576(7786):293–300. Available from: https://pubmed.ncbi.nlm.nih.gov/31802004/ [cited 2022 Sep 16] Ma H, Abdul-Hay M (2017) T-cell lymphomas, a challenging disease: types, treatments, and future. Int J Clin Oncol 22(1):18–51. Available from: https://pubmed.ncbi.nlm.nih.gov/27743148/ [cited 2022 Sep 16] MacLeod DT, Antony J, Martin AJ, Moser RJ, Hekele A, Wetzel KJ et al (2017) Integration of a CD19 CAR into the TCR alpha chain locus streamlines production of allogeneic gene-edited CAR T cells. Mol Ther [Internet] 25(4):949–961. Available from: https://pubmed.ncbi.nlm.nih. gov/28237835/ [cited 2022 Sep 16] Marin-Acevedo JA, Kimbrough EMO, Lou Y (2021) Next generation of immune checkpoint inhibitors and beyond. J Hematol Oncol [Internet] 14(1):1–29. Available from: https:// jhoonline.biomedcentral.com/articles/10.1186/s13045-021-01056-8 [cited 2022 Sep 16] Marino J, Paster J, Benichou G (2016) Allorecognition by T lymphocytes and allograft rejection. Front Immunol [Internet] 7. Available from: https://pubmed.ncbi.nlm.nih.gov/28018349/ [cited 2022 Sep 16] Maude SL, Laetsch TW, Buechner J, Rives S, Boyer M, Bittencourt H et al (2018) Tisagenlecleucel in children and young adults with B-cell lymphoblastic Leukemia. N Engl J Med [Internet] 378(5):439–448. Available from: https://pubmed.ncbi.nlm.nih.gov/29385370/ [cited 2022 Sep 16] McCarthy EF (2006) The toxins of William B. Coley and the treatment of bone and soft-tissue sarcomas. Iowa Orthop J 26:154. Available from: /pmc/articles/PMC1888599/ [cited 2022 Sep 16] Menger L, Sledzinska A, Bergerhoff K, Varga FA, Smith J, Poirot L et al (2016) TALEN-mediated inactivation of PD-1 in tumor-reactive lymphocytes promotes Intratumoral T-cell persistence and rejection of established Tumors. Cancer Res [Internet] 76(8):2087–2093. Available from: https://pubmed.ncbi.nlm.nih.gov/27197251/ [cited 2022 Sep 16] Moon EK, Wang LC, Dolfi DV, Wilson CB, Ranganathan R, Sun J et al (2014) Multifactorial T-cell hypofunction that is reversible can limit the efficacy of chimeric antigen receptor-transduced human T cells in solid tumors. Clin Cancer Res [Internet] 20(16):4262–4273. Available from: https://pubmed.ncbi.nlm.nih.gov/24919573/ [cited 2022 Sep 16] Moon EK, Ranganathan R, Eruslanov E, Kim S, Newick K, O’Brien S et al (2016) Blockade of programmed death 1 augments the ability of human T cells engineered to target NY-ESO-1 to control tumor growth after adoptive transfer. Clin Cancer Res [Internet] 22(2):436–447. Available from: https://pubmed.ncbi.nlm.nih.gov/26324743/ [cited 2022 Sep 16] Moretta A, Bottino C, Vitale M, Pende D, Biassoni R, Mingari MC et al (1996) Receptors for HLA class-I molecules in human natural killer cells. Annu Rev Immunol [Internet] 14:619–648. Available from: https://pubmed.ncbi.nlm.nih.gov/8717527/ [cited 2022 Sep 16] Mueller KP, Piscopo NJ, Forsberg MH, Saraspe LA, Das A, Russell B et al (2022) Production and characterization of virus-free, CRISPR-CAR T cells capable of inducing solid tumor regression. J Immunother Cancer [Internet] 10(9):e004446. Available from: https://jitc.bmj.com/ content/10/9/e004446 [cited 2022 Sep 16]

106

S. C. G. de Lima et al.

Nahmad AD, Reuveni E, Goldschmidt E, Tenne T, Liberman M, Horovitz-Fried M et al (2022) Frequent aneuploidy in primary human T cells after CRISPR–Cas9 cleavage. Nat Biotechnol [Internet] 1–7. Available from: https://www.nature.com/articles/s41587-022-01377-0 [cited 2022 Sep 16] Neelapu SS, Locke FL, Bartlett NL, Lekakis LJ, Miklos DB, Jacobson CA et al (2017) Axicabtagene Ciloleucel CAR T-cell therapy in refractory large B-cell lymphoma. N Engl J Med [Internet] 377(26):2531–2544. Available from: https://pubmed.ncbi.nlm.nih.gov/2922 6797/ [cited 2022 Sep 16] Nguyen DN, Roth TL, Li PJ, Chen PA, Apathy R, Mamedov MR et al (2020) Polymer-stabilized Cas9 nanoparticles and modified repair templates increase genome editing efficiency. Nat Biotechnol [Internet] 38(1):44–49. Available from: https://pubmed.ncbi.nlm.nih.gov/318192 58/ [cited 2022 Sep 16] Odé Z, Condori J, Peterson N, Zhou S, Krenciute G (2020) CRISPR-mediated non-viral sitespecific gene integration and expression in T cells: protocol and application for T-cell therapy. Cancers (Basel) [Internet] 12(6):1–16. Available from: https://pubmed.ncbi.nlm.nih.gov/32604 839/ [cited 2022 Sep 16] Olenchock BA, Guo R, Carpenter JH, Jordan M, Topham MK, Koretzky GA et al (2006) Disruption of diacylglycerol metabolism impairs the induction of T cell anergy. Nat Immunol [Internet] 7(11):1174–1181. Available from: https://www.nature.com/articles/ni1400 [cited 2022 Sep 17] Osborn MJ, Webber BR, Knipping F, Lonetree CL, Tennis N, DeFeo AP et al (2016) Evaluation of TCR gene editing achieved by TALENs, CRISPR/Cas9, and megaTAL nucleases. Mol Ther [Internet] 24(3):570–581. Available from: https://pubmed.ncbi.nlm.nih.gov/26502778/ [cited 2022 Sep 16] Park JH, Romero FA, Taur Y, Sadelain M, Brentjens RJ, Hohl TM et al (2018) Cytokine release syndrome grade as a predictive marker for infections in patients with relapsed or refractory B-cell acute lymphoblastic Leukemia treated with chimeric antigen receptor T cells. Clin Infect Dis [Internet] 67(4):533–540. Available from: https://pubmed.ncbi.nlm.nih.gov/29481659/ [cited 2022 Sep 16] Peng PD, Cohen CJ, Yang S, Hsu C, Jones S, Zhao Y et al (2009) Efficient nonviral sleeping beauty transposon-based TCR gene transfer to peripheral blood lymphocytes confers antigen-specific antitumor reactivity. Gene Ther [Internet] 16(8):1042–1049. Available from: https://www. nature.com/articles/gt200954 [cited 2022 Sep 16] Poirot L, Philip B, Schiffer-Mannioui C, Le Clerre D, Chion-Sotinel I, Derniame S, Potrel P et al (2015) Multiplex genome-edited T-cell manufacturing platform for ‘off-the-shelf’ adoptive T-cell immunotherapies. Cancer Res 75(18):3853–3864. https://doi.org/10.1158/0008-5472. CAN-14-3321 Png YT, Vinanica N, Kamiya T, Shimasaki N, Coustan-Smith E, Campana D (2017) Blockade of CD7 expression in T cells for effective chimeric antigen receptor targeting of T-cell malignancies. Blood Adv [Internet] 1(25):2348–2360. Available from: https://pubmed.ncbi.nlm.nih. gov/29296885/ [cited 2022 Sep 16] Porteus MH, Carroll D (2005) Gene targeting using zinc finger nucleases. Nat Biotechnol [Internet] 23(8):967–973. Available from: https://www.nature.com/articles/nbt1125 [cited 2022 Sep 16] Qasim W, Zhan H, Samarasinghe S, Adams S, Amrolia P, Stafford S et al (2017) Molecular remission of infant B-ALL after infusion of universal TALEN gene-edited CAR T cells. Sci Transl Med [Internet] 9(374):eaaj2013. Available from: https://pubmed.ncbi.nlm.nih.gov/2 8123068/ [cited 2022 Sep 16] Quann EJ, Liu X, Altan-Bonnet G, Huse M (2011) A cascade of protein kinase C isozymes promotes cytoskeletal polarization in T cells. Nat Immunol [Internet] 12(7):647–654. Available from: https://pubmed.ncbi.nlm.nih.gov/21602810/ [cited 2022 Sep 16] Quigley M, Pereyra F, Nilsson B, Porichis F, Fonseca C, Eichbaum Q et al (2010) Transcriptional analysis of HIV-specific CD8+ T cells shows that PD-1 inhibits T cell function by upregulating BATF. Nat Med [Internet] 16(10):1147–1151. Available from: https://pubmed.ncbi.nlm.nih. gov/20890291/ [cited 2022 Sep 16]

6

Genome Editing for Engineering the Next Generation of Advanced Immune. . .

107

Ren J, Zhang X, Liu X, Fang C, Jiang S, June CH et al (2017a) A versatile system for rapid multiplex genome-edited CAR T cell generation. Oncotarget [Internet] 8(10):17002–17011. Available from: https://www.oncotarget.com/article/15218/text/ [cited 2022 Sep 16] Ren J, Liu X, Fang C, Jiang S, June CH, Zhao Y (2017b) Multiplex genome editing to generate universal CAR T cells resistant to PD1 inhibition. Clin Cancer Res [Internet] 23(9):2255–2266. Available from: https://pubmed.ncbi.nlm.nih.gov/27815355/ [cited 2022 Sep 16] Robbins PF, Li YF, El-Gamil M, Zhao Y, Wargo JA, Zheng Z et al (2008) Single and dual amino acid substitutions in TCR CDRs can enhance antigen-specific T cell functions. J Immunol [Internet] 180(9):6116–6131. Available from: https://pubmed.ncbi.nlm.nih.gov/18424733/ [cited 2022 Sep 16] Roth TL, Puig-Saus C, Yu R, Shifrut E, Carnevale J, Li PJ et al (2018) Reprogramming human T cell function and specificity with non-viral genome targeting. Nature [Internet] 559(7714): 405–409. Available from: https://pubmed.ncbi.nlm.nih.gov/29995861/ [cited 2022 Sep 16] Ruella M, Xu J, Barrett DM, Fraietta JA, Reich TJ, Ambrose DE et al (2018) Induction of resistance to chimeric antigen receptor T cell therapy by transduction of a single leukemic B cell. Nat Med [Internet] 24(10):1499–1503. Available from: https://pubmed.ncbi.nlm.nih.gov/30275568/ [cited 2022 Sep 16] Rupp LJ, Schumann K, Roybal KT, Gate RE, Ye CJ, Lim WA et al (2017) CRISPR/Cas9-mediated PD-1 disruption enhances anti-tumor efficacy of human chimeric antigen receptor T cells. Sci Rep [Internet] 7(1):1–10. Available from: https://www.nature.com/articles/s41598-017-004 62-8 [cited 2022 Sep 16] Sadelain M, Brentjens R, Rivière I (2013) The basic principles of chimeric antigen receptor design. Cancer Discov [Internet] 3(4):388–398. Available from: https://pubmed.ncbi.nlm.nih.gov/23 550147/ [cited 2022 Sep 16] Scherer LD, Brenner MK, Mamonkin M (2019) Chimeric antigen receptors for T-cell malignancies. Front Oncol [Internet] 9. Available from: https://pubmed.ncbi.nlm.nih.gov/30891427 [cited 2022 Sep 16] Schober K, Müller TR, Gökmen F, Grassmann S, Effenberger M, Poltorak M et al (2019) Orthotopic replacement of T-cell receptor α- and β-chains with preservation of nearphysiological T-cell function. Nat Biomed Eng [Internet] 3(12):974–984. Available from: https://pubmed.ncbi.nlm.nih.gov/31182835/ [cited 2022 Sep 16] Scholler J, Brady TL, Binder-Scholl G, Hwang WT, Plesa G, Hege KM et al (2012) Decade-long safety and function of retroviral-modified chimeric antigen receptor T cells. Sci Transl Med [Internet] 4(132). Available from: https://pubmed.ncbi.nlm.nih.gov/22553251/ [cited 2022 Sep 16]; Schuster SJ, Bishop MR, Tam CS, Waller EK, Borchmann P, McGuirk JP et al (2019) Tisagenlecleucel in adult relapsed or refractory diffuse large B-cell lymphoma. N Engl J Med [Internet] 380(1):45–56. Available from: https://pubmed.ncbi.nlm.nih.gov/30501490/ [cited 2022 Sep 16] Seo H, Chen J, González-Avalos E, Samaniego-Castruita D, Das A, Wang YH et al (2019) TOX and TOX2 transcription factors cooperate with NR4A transcription factors to impose CD8 + T cell exhaustion. Proc Natl Acad Sci U S A [Internet] 116(25):12410–12415. Available from: https://pubmed.ncbi.nlm.nih.gov/31152140/ [cited 2022 Sep 16] Shao H, Zhang W, Hu Q, Wu F, Shen H, Huang S (2010) TCR mispairing in genetically modified T cells was detected by fluorescence resonance energy transfer. Mol Biol Rep [Internet] 37(8): 3951–3956. Available from: https://link.springer.com/article/10.1007/s11033-010-0053-y [cited 2022 Sep 16] Shifrut E, Carnevale J, Tobin V, Roth TL, Woo JM, Bui CT et al (2018) Genome-wide CRISPR screens in primary human T cells reveal key regulators of immune function. Cell [Internet] 175(7):1958–1971.e15. Available from: https://pubmed.ncbi.nlm.nih.gov/30449619/ [cited 2022 Sep 16]

108

S. C. G. de Lima et al.

Shui JW, Boomer JS, Han J, Xu J, Dement GA, Zhou G et al (2007) Hematopoietic progenitor kinase 1 negatively regulates T cell receptor signaling and T cell-mediated immune responses. Nat Immunol [Internet] 8(1):84–91. Available from: https://pubmed.ncbi.nlm.nih.gov/171150 60/ [cited 2022 Sep 16] Si J, Shi X, Sun S, Zou B, Li Y, An D et al (2020) Hematopoietic progenitor Kinase1 (HPK1) mediates T cell dysfunction and is a Druggable target for T cell-based immunotherapies. Cancer Cell [Internet] 38(4):551–566.e11. Available from: https://pubmed.ncbi.nlm.nih.gov/32860 752/ [cited 2022 Sep 16] Singh N, Perazzelli J, Grupp SA, Barrett DM (2016) Early memory phenotypes drive T cell proliferation in patients with pediatric malignancies. Sci Transl Med [Internet] 8(320). Available from: https://pubmed.ncbi.nlm.nih.gov/26738796/ [cited 2022 Sep 16] Song S, Han M, Zhang H, Wang Y, Jiang H (2013) Full screening and accurate subtyping of HLA-A*02 alleles through group-specific amplification and mono-allelic sequencing. Cell Mol Immunol [Internet] 10(6):490–496. Available from: https://www.nature.com/articles/ cmi201333 [cited 2022 Sep 16] Stadtmauer EA, Fraietta JA, Davis MM, Cohen AD, Weber KL, Lancaster E et al (2020) CRISPRengineered T cells in patients with refractory cancer. Science (1979) [Internet] 367(6481). Available from: https://www.science.org/doi/10.1126/science.aba7365 [cited 2022 Sep 16] Sterner RM, Sakemura R, Cox MJ, Yang N, Khadka RH, Forsman CL et al (2019) GM-CSF inhibition reduces cytokine release syndrome and neuroinflammation but enhances CAR-T cell function in xenografts. Blood [Internet] 133(7):697–709. Available from: https://pubmed.ncbi. nlm.nih.gov/30463995/ [cited 2022 Sep 16] Su S, Hu B, Shao J, Shen B, Du J, Du Y et al (2016) CRISPR-Cas9 mediated efficient PD-1 disruption on human primary T cells from cancer patients. Sci Rep [Internet]:6. Available from: https://pubmed.ncbi.nlm.nih.gov/26818188/ [cited 2022 Sep 16] Tang N, Cheng C, Zhang X, Qiao M, Li N, Mu W et al (2020) TGF-β inhibition via CRISPR promotes the long-term efficacy of CAR T cells against solid tumors. JCI Insight [Internet] 5(4). Available from: https://pubmed.ncbi.nlm.nih.gov/31999649/ [cited 2022 Sep 16] Teachey DT, Lacey SF, Shaw PA, Melenhorst JJ, Maude SL, Frey N et al (2016) Identification of predictive biomarkers for cytokine release syndrome after chimeric antigen receptor T-cell therapy for acute lymphoblastic Leukemia. Cancer Discov [Internet] 6(6):664–679. Available from: https://pubmed.ncbi.nlm.nih.gov/27076371/ [cited 2022 Sep 16] Torikai H, Reik A, Liu PQ, Zhou Y, Zhang L, Maiti S et al (2012) A foundation for universal T-cell based immunotherapy: T cells engineered to express a CD19-specific chimeric-antigen-receptor and eliminate expression of endogenous TCR. Blood [Internet] 119(24):5697–5705. Available from: https://pubmed.ncbi.nlm.nih.gov/22535661/ [cited 2022 Sep 16] Urnov FD, Rebar EJ, Holmes MC, Zhang HS, Gregory PD (2010) Genome editing with engineered zinc finger nucleases. Nat Rev Genet [Internet] 11(9):636–646. Available from: https://www. nature.com/articles/nrg2842 [cited 2022 Sep 16] Vong Q, Nye C, Hause R, Clouser C, Jones J, Burleigh S, Borges CM, Chin MSY, Marco E, Barrera L (2017) Inhibiting TGFβ signaling in CAR T-cells may significantly enhance efficacy of tumor immunotherapy. Blood 130:1791 Waldman AD, Fritz JM, Lenardo MJ (2020) A guide to cancer immunotherapy: from T cell basic science to clinical practice. Nat Rev Immunol 20(11):651–668. Available from: https://www. nature.com/articles/s41577-020-0306-5 [cited 2022 Sep 16] Wang X, Li JP, Chiu LL, Lan JL, Chen DY, Boomer J et al (2012) Attenuation of T cell receptor signaling by serine phosphorylation-mediated lysine 30 ubiquitination of SLP-76 protein. J Biol Chem [Internet] 287(41):34091–34100. Available from: https://pubmed.ncbi.nlm.nih.gov/22 902619/ [cited 2022 Sep 16] Wang J, DeClercq JJ, Hayward SB, Li PWL, Shivak DA, Gregory PD et al (2016) Highly efficient homology-driven genome editing in human T cells by combining zinc-finger nuclease mRNA and AAV6 donor delivery. Nucleic Acids Res [Internet] 44(3):e30–e30. Available from: https:// academic.oup.com/nar/article/44/3/e30/2502646 [cited 2022 Sep 16]

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Genome Editing for Engineering the Next Generation of Advanced Immune. . .

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Wang M, Munoz J, Goy A, Locke FL, Jacobson CA, Hill BT et al (2020) KTE-X19 CAR T-cell therapy in relapsed or refractory mantle-cell lymphoma. N Engl J Med [Internet] 382(14): 1331–1342. Available from: https://pubmed.ncbi.nlm.nih.gov/32242358/ [cited 2022 Sep 16] Wang D, Prager BC, Gimple RC, Aguilar B, Alizadeh D, Tang H et al (2021) CRISPR screening of CAR T cells and cancer stem cells reveals critical dependencies for cell-based therapies. Cancer Discov [Internet] 11(5):1192–1211. Available from: https://pubmed.ncbi.nlm.nih.gov/3332821 5/ [cited 2022 Sep 16] Wei J, Long L, Zheng W, Dhungana Y, Lim SA, Guy C et al (2019) Targeting REGNASE-1 programs long-lived effector T cells for cancer therapy. Nature [Internet] 576(7787):471–476. Available from: https://pubmed.ncbi.nlm.nih.gov/31827283/ [cited 2022 Sep 16] Westin JR, Kersten MJ, Salles G, Abramson JS, Schuster SJ, Locke FL et al (2021) Efficacy and safety of CD19-directed CAR-T cell therapies in patients with relapsed/refractory aggressive B-cell lymphomas: observations from the JULIET, ZUMA-1, and TRANSCEND trials. Am J Hematol [Internet] 96(10):1295–1312. Available from: https://pubmed.ncbi.nlm.nih.gov/34310 745/ [cited 2022 Sep 16] Leisegang M, Wilde S, Spranger S, Milosevic S, Frankenberger B, Uckert W et al (2010) MHC-restricted fratricide of human lymphocytes expressing survivin-specific transgenic T cell receptors. J Clin Invest [Internet] 120(11):3869–3877. Available from: https://pubmed. ncbi.nlm.nih.gov/20978348/ [cited 2022 Sep 16] Zetsche B, Gootenberg JS, Abudayyeh OO, Slaymaker IM, Makarova KS, Essletzbichler P et al (2015) Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Cas system. Cell [Internet] 163(3):759–771. Available from: https://pubmed.ncbi.nlm.nih.gov/26422227/ [cited 2022 Sep 16] Zetsche B, Heidenreich M, Mohanraju P, Fedorova I, Kneppers J, Degennaro EM et al (2017) Multiplex gene editing by CRISPR-Cpf1 using a single crRNA array. Nat Biotechnol [Internet] 35(1):31–34. Available from: https://pubmed.ncbi.nlm.nih.gov/27918548/ [cited 2022 Sep 16] Zhang Y, Zhang Z (2020) The history and advances in cancer immunotherapy: understanding the characteristics of tumor-infiltrating immune cells and their therapeutic implications. Cell Mol Immunol [Internet] 17(8):807–821. Available from: https://www.nature.com/articles/s41423020-0488-6 [cited 2022 Sep 16] Zhang L, Yu Z, Muranski P, Palmer DC, Restifo NP, Rosenberg SA et al (2012) Inhibition of TGF-β signaling in genetically engineered tumor antigen-reactive T cells significantly enhances tumor treatment efficacy. Gene Ther [Internet] 20(5):575–580. Available from: https://www. nature.com/articles/gt201275 [cited 2022 Sep 16] Zhang C, Liu J, Zhong JF, Zhang X (2017a) Engineering CAR-T cells. Biomark Res [Internet] 5(1): 1–6. Available from: https://biomarkerres.biomedcentral.com/articles/10.1186/s40364-0170102-y [cited 2022 Sep 16] Zhang Y, Zhang X, Cheng C, Mu W, Liu X, Li N et al (2017b) CRISPR-Cas9 mediated LAG-3 disruption in CAR-T cells. Front Med [Internet] 11(4):554–562. Available from: https:// pubmed.ncbi.nlm.nih.gov/28625015/ [cited 2022 Sep 16] Zhang J, Hu Y, Yang J, Li W, Zhang M, Wang Q et al (2022a) Non-viral, specifically targeted CAR-T cells achieve high safety and efficacy in B-NHL. Nature [Internet] 609(7926). Available from: https://pubmed.ncbi.nlm.nih.gov/36045296/ [cited 2022 Sep 16] Zhang N, Si J, Li G, Wang Y, Long F, Wang T et al (2022b) P1464: decreasing HPK1 expression in CD19 CAR-T cells: a novel strategy to overcome challenges of cell therapy for adult (R/R) B-ALL. Hemasphere [Internet] 6(Suppl):1346–1347. Available from: /pmc/articles/ PMC9431273/ [cited 2022 Sep 16] Zhao J, Lin Q, Song Y, Liu D (2018) Universal CARs, universal T cells, and universal CAR T cells. J Hematol Oncol [Internet] 11(1):1–9. Available from: https://jhoonline.biomedcentral.com/ articles/10.1186/s13045-018-0677-2 [cited 2022 Sep 16] Zheng W, Wei J, Zebley CC, Jones LL, Dhungana Y, Wang YD et al (2021) Regnase-1 suppresses TCF-1+ precursor exhausted T-cell formation to limit CAR-T-cell responses against ALL.

110

S. C. G. de Lima et al.

Blood [Internet] 138(2):122–135. Available from: https://pubmed.ncbi.nlm.nih.gov/33690816/ [cited 2022 Sep 16] Zhong XP, Hainey EA, Olenchock BA, Jordan MS, Maltzman JS, Nichols KE et al (2003) Enhanced T cell responses due to diacylglycerol kinase zeta deficiency. Nat Immunol [Internet] 4(9):882–890. Available from: https://pubmed.ncbi.nlm.nih.gov/12883552/ [cited 2022 Sep 16]

Chapter 7

CRISPR Genome Editing and the Study of Chagas Disease Gabriela de Assis Burle-Caldas, Viviane Grazielle-Silva, Lídia Paula Faustino, and Santuza Maria Ribeiro Teixeira

Chagas disease (CD), described by Carlos Chagas more than 110 years ago, remains a serious health problem affecting about 6–8 million people worldwide and causing approximately 50,000 deaths per year. Endemic in Latin America, CD has spread to other continents due to migrations with current estimations indicating that approximately 75 million people worldwide are at risk of contracting this disease (PerezMolina and Molina 2018). The lack of a deep understanding of the distinct biology of its causative agent, the protozoan parasite Trypanosoma cruzi with its highly heterogeneous population and complex life cycle (Brener 1973; El-Sayed et al. 2005) may be one of the main reasons why we still do not have a more effective treatment or better strategies to control parasite transmission. Although significant progress toward the understanding of molecular mechanisms behind the highly efficient strategies of immune evasion that allow the parasite to persist for decades in its hosts have been made in the last few years, most of these studies require efficient genetic manipulation methodologies, which are only now becoming available. The publication of the complete genome sequence of T. cruzi in 2005, with an estimated haploid size of 55 Mb and about 12,000 genes, was a major breakthrough that allowed studies involving not only unique parasite genes but also genome-wide analyses of virulence factors responsible for the pathogenesis of Chagas disease (Unnikrishnan and Burleigh 2004). Having complete genome sequences for different parasite strains also allowed comparative genome studies as well as in silico,

G. de Assis Burle-Caldas · V. Grazielle-Silva · S. M. R. Teixeira (✉) Departamento de Bioquímica e Imunologia, Universidade Federal de Minas Gerais, Belo Horizonte, Brazil e-mail: [email protected] L. P. Faustino Centro de Pesquisa Rene Rachou, Fundação Osvaldo Cruz, Minas Gerais, Brazil © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_7

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large-scale screening analyses of molecular targets for the development of new diagnostic methods and therapeutic interventions (El-Sayed et al. 2005). Well before its complete genome was available, the first T. cruzi gene knockout, described in 1993, showed a striking morphological phenotype after disruption of the two alleles of the gene encoding the insect stage-specific glycoprotein GP72 (Cooper et al. 1993). Using the classical homologous recombination (HR) protocol for gene deletion that is based on sequential transfection of T. cruzi epimastigotes with two plasmids, the authors revealed a role of the GP72 surface glycoprotein as a mediator of the attachment of the single flagellum to the parasite membrane. The two plasmids were constructed by inserting in each one a different drug-resistant marker, the neomycin phosphotransferase and hygromycin phosphotransferase genes, flanked by GP72 sequences. Due to low transfection rates as well as low recombination efficiency, all studies that followed the work of Cooper et al. (1993) showed that selection of knockout parasite cell lines resistant to the two drugs usually takes 2–3 months. Because of that, almost 20 years later, only a limited number of studies have demonstrated the function of T. cruzi genes using this strategy to generate null mutant parasites. It is interesting to notice, however, that attempts to knockout both alleles of an essential T. cruzi gene by HR sometimes yield double-resistant parasites bearing the two drug resistance markers inserted in the target locus but also bearing extra copies of the target gene that have arisen by unknown mechanisms of gene amplification and translocation (Cardoso et al. 2013). Also, when targeting T. cruzi essential genes, other studies have also shown that disruption of a single allele results in a 50% reduction in the levels of the target mRNA and protein, thus providing a way to assess the phenotype of cells with reduced expression of the target gene (Regis-da-Silva et al. 2006). Together with gene deletion methods to study gene function, gene manipulation strategies have been used to generate parasites overexpressing a specific protein or parasites in which reporter or tagged genes are created to investigate infection rates, protein cellular localization, gene regulation, etc. (Perez-Diaz et al. 2012; KangussuMarcolino et al. 2013; Araujo et al. 2011). Similar to studies based on gene deletion, these studies have also significantly contributed to the understanding of molecular mechanisms involved in host parasite interaction and the establishment of CD (Taylor et al. 2011). The advances observed with T. cruzi studies were however much more limited compared to the studies with another member of the trypanosomatid family, Trypanosoma brucei, a parasite that can cause human and cattle diseases in African countries. One main reason is the fact that, in contrast to T. brucei, in which RNA interference (RNAi) gene knockdown can be used as a more versatile and rapid technique to investigate gene function, the T. cruzi genome does not contain the genes required to assemble an RNAi machinery (El-Sayed et al. 2005; DaRocha et al. 2004a). Also different from T. cruzi, a combination of the T7 RNA polymerase/TetR inducible system that can regulate the RNAi knockdown process, gene silencing has allowed a number of large-scale functional genomics studies addressing different aspects of the T. brucei biology and interaction with its host (Schumann Burkard et al. 2011). Importantly, inducible RNAi knockdown studies targeting essential genes have also been used to provide a first glimpse in

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the phenotype of a mutant that could not be generated using conventional HR gene disruption protocols (Currier et al. 2019; Zikova et al. 2009; Nittolo et al. 2018). Initially, conventional electroporation of epimastigotes, the insect form of the parasite that can be easily cultivated in the laboratory, was used in all experiments requiring introduction of exogenous DNA in T. cruzi (DaRocha et al. 2004b). More recently, DNA transfection efficiency was largely improved with a more efficient methodology named nucleofection. Using the Amaxa Nucleofector system, we have shown that 10% of transiently transfected T. cruzi epimastigotes can be achieved with 90% less plasmid DNA, compared to conventional electroporation (BurleCaldas et al. 2015). However, despite the advent of new transfection technologies, gene knockout based on HR remained a poorly efficient method used in the majority of studies involving genome manipulation not only in trypanosomes but also in most eukaryotic cells. HR is actually a conserved pathway that can repair double-stranded DNA break (DSB) by using an undamaged sister chromosome as a template (Li and Heyer 2008). Knowing that the generation of gene knockout is highly depended on the HR pathway, the idea of inducing a site-specific DSB to enhance integration rates of exogeneous sequences seemed highly plausible, since in the “classical” HR knockout experiments, success rate is strictly dependent on the generation of DSB at the right chromosomal locus. Zinc finger nucleases (ZFNs) are engineered proteins composed of a DNA-binding domain with a zinc finger motif and a cleavage domain of the FokI restriction endonuclease capable of binding and generating a DSB at a target DNA sequence (Durai et al. 2005). Since ZFNs were successfully used in mammalian cells, Burle-Caldas et al. (2017) expressed, in epimastigotes, a pair of ZFNs designed to recognize a sequence in the TcGP72 gene. It was shown that the DSB caused by the nuclease is readily repaired by HR if homologous sequences that are able to pair with the broken DNA are present (Burle-Caldas et al. 2017). In contrast to parasites expressing the ZFNs and transfected with a donor template DNA, no TcGP72 null mutant phenotype was observed in parasites that have not been transfected with a donor DNA fragment. Besides showing the importance of the HR repair pathway, the experiments with the ZFNs clearly showed that the introduction of a site-specific DSB results in improved knockout efficiency: less than 2 weeks after transfecting epimastigotes expressing ZFNs with a DNA fragment containing the neomycin resistance gene flanked by TcGP72 sequences, a population of G418-resistant parasites, in which over 90% of the cells have the TcGP72 null mutant phenotype, was obtained (Burle-Caldas et al. 2017). The use of ZFNs was, however, limited because, for each gene to be disrupted, it was necessary to design and express in the parasite a protein with a zinc finger domain that binds the specific nucleotide sequence in the target gene with high affinity. Although significant progress toward the development of genetic tools for manipulating the T. cruzi genome was achieved, it was not until 2014 that a major methodological breakthrough began to accelerate the studies requiring gene knockout in this parasite, particularly when multigene families are the genes of interest. With the advent of the first CRISPR-Cas9 gene editing protocols published soon after Jennifer Doudna’s seminal article describing the potential of this new technology, a number of studies using different CRISPR-Cas9 strategies have been

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disseminated in different laboratories that study T. cruzi and CD. Genome editing by clustered regularly interspaced short palindromic repeats (CRISPR) requires the expression in the target cell of the nuclease Cas9 (CRISPR-associated gene 9) together with an RNA molecule known as single guide RNA (sgRNA) (Jinek et al. 2012; Cong et al. 2013). Guided to a specific sequence within the target genome after base-pairing with the sgRNA, Cas9 nuclease generates a precise cleavage in the DNA that is subsequently repaired through two main cellular DNA repair pathways, the nonhomologous end-joining (NHEJ) and the homologous recombination (HR) pathways. Whereas NHEJ repair results in the imprecise disruption of the target sequence, HR repair allows the introduction of specific modifications of the target locus (Doudna et al. 2014). To promote HR repair, a DNA repair template is required, i.e., a DNA fragment with sequence homology to the sequence in which the DSB have occurred. Although genome studies have revealed that in trypanosomatids the components of the NHEJ, the second repair mechanism that is highly efficient in mammalian cells in repairing DSB, is absent, an alternative, error-prone repair mechanism has been described (Passos-Silva et al. 2010). In the absence of a donor sequence to allow HR, it has been proposed that DSB damage can be also repaired by the error-prone microhomology-mediated end-joining (MMEJ), a mechanism that also results in small deletions in the target gene (Glover et al. 2011). Although this mechanism has not yet been clearly demonstrated in T. cruzi, doublestrand breaks (DSB) caused by Cas9 can be repaired by MMEJ. However, as shown by the studies with the ZFNs, if a repair template that shares homology with the target gene is available, the DSB is not only more efficiently repaired by the homologous HR repair pathway, compared to MMEJ pathway, but also can result in the insertion of specific mutations or sequence tags in the gene of interest (BurleCaldas et al. 2017; Glover et al. 2011). As summarized in Fig. 7.1, four main strategies to perform CRISPR/Cas9 editing in T. cruzi have been used. In the first strategy, described by Rick Tarleton’ group in 2014, parasites constitutively expressing Cas9 from Streptococcus pyogenes (SpCas9) were generated by transfecting T. cruzi with the pTrex-b-SpCas9 plasmid (Peng et al. 2014). The plasmid was generated by subcloning the coding sequence of codon-optimized SpCas9 in the T. cruzi expression vector pTrex (Vazquez and Levin 1999), and this parasite cell line was also modified to express the enhanced green fluorescent protein (eGFP) gene reporter. After transfecting epimastigotes constitutively expressing SpCas9 and eGFP with in vitro transcribed sgRNAs, containing sequences of the eGFP gene, 60% of the transfected parasite population had the eGFP gene reporter disrupted only 5 days after transfection. Using the same strategy, the authors also showed the disruption of β-galactofuranosyl glycotransferase (β-GalGT), a family of 65 annotated genes, that share 93.1% nucleotide sequence identity. Whole-genome sequencing of eGFP and β-GalGT mutant cell lines showed that disruption of these genes occurred as a result of DSB repair by MMEJ, which is consistent with the fact that no DNA repair template to allow DSB repair by HR was provided. In the second description of a protocol for CRISPR editing in T. cruzi, Lander and coworkers (2015) used plasmids to constitutively express both the Cas9 nuclease

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Fig. 7.1 Four main strategies used to perform gene editing in T. cruzi with CRISPR/Cas9 technology. (1) In the first step of strategy 1, WT parasites are transfected with a plasmid containing SpCas9 sequence to generate stable cell lines expressing SpCas9. Next, parasites expressing SpCas9 are transfected with in vitro transcribed sgRNA that targets the gene of interested (GOI). No drug selection is required considering that no repair template is used and the DSB generated by Cas9 is repaired by MMEJ. Three to five days after transfection, mutant cells lines are generated. (2) In strategy 2, only one step is required to generate mutant cell lines. Wild-type parasites are co-transfected with a plasmid containing SpCas9 sequence and the sequence of the sgRNA that targets the GOI and with a repair template, i.e., a DNA fragment containing a gene that confers resistance to blasticidin (BSD) flanked by sequences of the GOI. After drug selection, mutant cell lines are generated. (3) In strategy 3, the first step is the generation of parasites stably expressing SpCas9 and T7 RNA polymerase by transfecting epimastigotes with the plasmid pLEW13-Cas9 containing both sequences. Next, parasites stably expressing SpCas9 and T7 RNA polymerase are transfected with PCR products containing sequence of the sgRNAs that targets the GOI downstream from the T7 RNA polymerase promoter and with a repair template with BSD resistance gene. After drug selection, mutant cell lines are generated. (4) In strategy 4, generation of mutant cell lines take only one step, but different from strategies 1 to 3, wild-type parasites are co-transfected with recombinant SaCas9 protein, produced in E. coli, and complexed with in vitro transcribed sgRNA that targets the GOI. An oligonucleotide containing sequences of the GOI with stop codons in all reading frames is used as a repair template. No drug selection is needed, since the repair template does not contain a resistance marker. Few days after transfection, mutant cell lines are generated

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and the sgRNA. Using endogenous T. cruzi genes as targets, these authors tested two different strategies. Because T. cruzi null mutants of the TcPFR1, TcPFR2, and TcGP72 genes have been previously described and their phenotypes could be easily determined due to morphological changes, these genes were selected as targets. In the first protocol, the SpCas9 with a SV40 nuclear localization signal and a GFP tag was expressed from one plasmid, whereas the sgRNAs containing sequences of each target gene were expressed from a second plasmid. In the second plasmid, a pTREX vector that also contains the tdTomato reporter gene, the sgRNA sequence were inserted downstream from the T. cruzi ribosomal promoter (sgRNA/tdTomato/ pTREX). As an alternative strategy, parasite transfection was performed with only one plasmid, the pTREX vector containing both sequences, i.e., the SpCas9 and the sgRNA. Both strategies were first tested in the absence of a repair template, with the DSB expected to be repaired by MMEJ. After transfection, using plasmids carrying sgRNA sequences targeting TcPFR1, TcPFR2, or TcGP72, the authors observed a mix parasite population, with some cells displaying morphological alterations consistent with the disruption of these genes, i.e., parasites with their flagellum detached from the cell membrane. However, sequence analysis of a clonal mutant population to TcPFR2 failed to find mutations in the genome sequence of TcPFR2 consistent with MMEJ repair. For that reason, the authors decided to co-transfect the parasites with a DNA repair template composed of the gene that confers resistance to blasticidin (BSD), flanked by 100 bp of homology arms, together with the plasmid containing SpCas9 and sgRNA complementary to the TcPFR2 gene. After drug selection, Northern blot and PCR analysis of the transfected population confirmed the efficient disruption, by HR, of TcPFR2 gene with the insertion of the BSD resistance marker. A third strategy, described by Costa et al. (2018), requires the generation of parasites constitutively expressing SpCas9 and the T7 RNA polymerase (Costa et al. 2018). Besides expression of the nuclease, using this strategy, constitutive expression of the T7 RNA polymerase is also required to transcribe, within the parasite, the sgRNA-containing sequences of the target gene. A transfected PCR product containing the target sequence downstream from the T7 RNA polymerase promoter was used as template to transcribe the sgRNA. To express both enzymes, the SpCas9 and T7 RNA polymerase, the authors used the pLEW13-Cas9 vector which integrates in the β-tubulin locus. Using again the TcGP72 gene as proof of concept, they generated a PCR product containing sequences of TcGP72 and the T7 RNA polymerase promoter, which was delivered to parasite nucleus by nucleofection. In contrast to the results from Peng et al. (2015), which showed a negative effect of the expression of the SpCas9 on the parasite doubling, Costa et al. (2018) did not observe differences in the growth curve of epimastigotes stably expressing SpCas9 and T7 RNA polymerase compared to WT parasites. Using this strategy, the authors demonstrated efficient disruption of TcGP72 gene after transfection of parasites stably expressing Cas9 and T7 RNA polymerase with two repair templates (one containing the blasticidin resistance and another with puromycin resistance genes, both with 30-bp homology arms). The authors have also showed the successful replacement of the reporter gene mNeonGreen previously integrated in the genome

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of the parasite, by the reporter gene mScarlet and the addition of a C-terminal tag with the mNeonGreen in the gene of DNA topoisomerase 1A. Finally, a fourth strategy, first described in mammalian cells (Kim et al. 2014), also resulted in highly efficient editing of T. cruzi genes using recombinant Cas9 protein transfected into epimastigotes. However, because the Cas9 nuclease from Staphylococcus aureus (SaCas9) is significantly smaller than the S. pyogenes enzyme, purified recombinant SaCas9 expressed in E. coli was used in this protocol (Soares Medeiros et al. 2017; Burle-Caldas et al. 2018). Before transfection, the recombinant nuclease was incubated with in vitro transcribed RNA, and the ribonucleoprotein (RNP) complex was transfected into parasites using the Amaxa Nucleofector device. As shown by Soares-Medeiros et al. (2017), a highly efficient disruption (72–90%) of two reporter genes, tdTomato and eGFP, was observed in different T. cruzi strains. It is noteworthy that the authors also described that Cas9RNP size impacts knockout efficiency since they failed to generate knockout cell lines using SpCas9 which has a molecular weight of 164 kDa, while SaCas9 has 124 kDa. Although less efficient compared to the transfection of epimastigotes, transfection of trypomastigotes with the SaCas9/sgRNA RNP complex also resulted in gene knockouts. Besides using the RNP transfection strategy, the authors also showed disruption of endogenous genes by co-delivering with the RNP a repair oligonucleotide template. This oligonucleotide is composed of a single-stranded sequence with three stop codons in all reading frames, flanked by short homology arms (30 nucleotides) complementary to the target genes. SaCas9/sgRNA RNP complex targeting two alleles encoding a conserved hypothetical protein or the T. cruzi nitroreductase (NTR) gene co-transfected with the repair oligonucleotide resulted in mutant alleles in the population 15 days after electroporation (Soares Medeiros et al. 2017). Using the oligonucleotide repair template containing sequences homologous to the TcGP72 gene with two in-frame influenza hemagglutinin (HA) epitopes, the authors also showed that the SaCas9/ sgRNA RNP complex is highly efficient in generating endogenous gene tagging (Soares Medeiros et al. 2017). These results showed that the SaCas9/sgRNArepair-templated protocol is as efficient in creating a tagged protein (gene knock in) and in generating knockout mutants. Following the publication of different strategies describing efficient CRISPR/ Cas9 editing including transfection with RNP complex, Burle-Caldas et al. (2018) compared these strategies by determining the efficiencies of a protocol that required the generation of parasites constitutively expressing SpCas9 and a protocol based on transfection of WT parasites with the recombinant SaCas9 RNP complex. For both tests, the TcGP72 gene was used as target gene, and in vitro transcribed sgRNAs containing TcGP72 sequences and oligonucleotide repair templates were generated. Different from the work described by other groups, Burle-Caldas et al. (2018) used the pROCKSpCasNeo, a plasmid that integrates in the β-tubulin locus in the T. cruzi genome, to generate a stable transfected population of parasites expressing SpCas9 fused to GFP (DaRocha et al. 2004b). After selecting the SpCas9 expressing parasites, they were co-transfected with the in vitro transcribed sgRNA targeting TcGP72 and a repair template composed of oligonucleotide sequences containing

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three in frame stop codons, flanked by 40 nucleotides of TcGP72 homology arms. Five days after transfection, 50% of the parasites showed disruption of both TcGP72 alleles. Re-transfecting parasites with sgRNA and the repair template, 3 days after the first transfection, increased the knockout efficiency to 99.9%. The authors have also showed that, in the absence of a repair template, transfection of parasites stably expressing SpCas9 with in vitro transcribed sgRNAs targeting TcGP72 resulted in an abnormal morphology that does not correspond to the typical TcGP72 knockout phenotype. The efficiency of disruption of both alleles of the TcGP72 gene using the strategy of parasites stably expressing SpCas9 was significantly superior when compared with the strategy of transfection with SaCas9/sgRNA RNP complex (Soares Medeiros et al. 2017). As shown by Burle-Caldas et al. (2018), an efficiency of 10% of parasites with disrupted TcGP72 was obtained with wild-type epimastigotes co-transfected with the SaCas9/sgRNA RNP complex and an oligonucleotide repair template. Re-transfection of the same parasite population resulted in increased knockout efficiency up to 34%. Despite its lower efficiency, the strategy using SaCas9/sgRNA RPN complex has the advantages of not requiring the development of a parasite cell line with constitutive expression of the Cas9 nuclease. Most importantly, considering that constitutively expression of Cas9 may lead to offtarget effects (Kim et al. 2014), transient expression with the nuclease greatly reduces this risk. Soon after the first protocols describing CRISPR/Cas9 gene knockouts were published, various groups have shown that CRISPR technology can be used to generate parasites overexpressing a protein or expressing a protein with sequence tags inserted in the endogenous gene. Besides the determination of protein localization, protein fusion with epitope tags is widely used in studies of protein-protein interactions, and the ability to insert sequence tags in the endogenous gene location allows the expression of the tagged protein within their natural regulatory context. Lander et al. (2016), used the CRISPR/Ca9 strategy to create endogenous tags, with a 3X HA or 3X c-myc, in two T. cruzi genes. Using a c-myc tag, the authors showed that the T. cruzi mitochondrial calcium uniporter (TcMCU), the orthologue of T. brucei MCU, responsible for mitochondrial Ca2+ uptake, co-localizes with the mitochondrial voltage-dependent channel (VDAC), as expected. After HA-tagging the gene encoding the inositol 1,4,5-triphosphate receptor (TcIP3), these authors showed that the protein localizes to the acidocalcisomes. Importantly, the protein localization observed after endogenously tagging the TcIP3 gene was different from the results obtained when the TcIP3 gene that was overexpressed with an HA tag from an ectopic locus. Besides protein tagging using CRISPR/Cas9, a recent adaptation of the editing protocol described by Picchi-Constante et al. 2022, allowed the investigation of the role of essential T. cruzi genes (Picchi-Constante et al. 2022). With this protocol, knocking out the target gene is combined with its replacement with a mutated copy carrying any kind of desired alterations, including changes in specific amino acid residues, motifs, and protein domains. The DNA repair template used to replace the target sequence also carries modifications that result in resistance to Cas9 cleavage. Using the gene encoding the histone deacetylase gene tcDAC2 as a proof of concept

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and different DNA repair templates sequences to allow homology-driven repair, the authors were able to generate parasites expressing different variants of the gene to be used in functional studies. The use of this strategy in T. cruzi is particularly welcome since gene silencing using inducible RNA interference (RNAi), which have been largely employed in functional studies of essential genes in T. brucei, cannot be done in T. cruzi. Indeed, the enormous potential of CRISPR/Cas9 editing in T. cruzi have already resulted in an exponential increase in the number of studies that investigate, at the molecular level, different aspects of the parasite biology and its interactions with the mammalian host. After more than 100 years of the study of CD pathology, it is now well established that the damage caused by CD is not only due to the presence of parasite in infected tissues, which can often last for decades, but also to a continuous inflammatory reaction in the heart and other human tissues. Although major progress toward the identification of parasite factors that cause the CD pathology and contribute to the severity of the disease, a better understanding of the complex interactions between T. cruzi and the human host still requires the molecular characterization of parasite genes responsible for (1) parasite proliferation, differentiation, and survival within different host environments; (2) parasite escape from host immune response and modulation of host immune pathways; and (3) genetic elements that control gene expression of these parasite virulence factors. We next describe a group of experiments that have been published in the past 5 years, which showed clearly how much improvement we are experiencing toward the molecular characterization of T. cruzi and how these studies have been largely benefitted from the highly efficient CRISPR protocols. Similar to other members of the Trypanosomatidae family, control of gene expression in T. cruzi relies on post-transcriptional mechanisms. These regulatory mechanisms are largely dependent on RNA-binding proteins (RBPs), which act as key modulators of mRNA stability and translation efficiency (Araujo and Teixeira 2011). Global gene expression analyses have identified several T. cruzi RBPs, some of them showing differential expression when epimastigotes, trypomastigotes, and amastigotes are compared. One of these RBPs, a zinc finger protein, presenting a tenfold higher expression in epimastigotes compared to the mammalian stages, has been studied using CRISPR/Cas9 to generate knockout cell lines. After performing comparative RNA-seq analyses of wild-type and knockout cell lines for this zinc finger protein, Tavares-Silva and coworkers (2020) provided evidence for a role of this RBP as a positive regulator of the expression of epimastigote-specific genes that controls epimastigote proliferation as well as a negative regulator of genes that trigger metacyclogenesis (Tavares et al. 2021). Different groups have used CRISPR/Cas9 to study T. cruzi DNA metabolism. Pavani et al. (2020), generate parasite cell lines with mutations inserted in one subunit of the heterotrimeric replication protein (RPA) complex composed by RPA-1, RPA-2, and RPA-3, which is implicated in DNA replication and damage response. Metacyclogenesis assays with parasites with mutations in a leucine-rich exportation signal of the RPA-2 subunit showed that these mutations impaired the differentiation process (Pavani et al. 2020). Studying another group of proteins

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involved in DNA metabolism and gene expression, Pezza et al. (2022), tried to use CRISPR/Cas9 to generate knockout cell lines of TcBDF2, an ortholog of a bromodomains (BDs) factor recently identified in Leishmania (Pezza et al. 2022). This family of modifier reader proteins recognizes the acetylation marks on histones and acts as a scaffold for the assembly of macromolecular complexes that interact with the chromatin. Despite several attempts, these authors were only able to generate single-allele knockout parasites of TcBDF2 and showed that one copy of TcBDF2 gene is enough for normal growth of epimastigotes. Protein phosphorylation, involved in several important biological processes in the T. cruzi life cycle, has been investigated by various groups using CRISPR/Cas9 gene editing. Malvezzi and collaborators (2020) generated knockout parasites of the T. cruzi kinase 1 (Tck1), a kinase implicated in the phosphorylation of the initiation factor 2 alpha subunit (eIF2α) in response to changes in nutrients levels (Malvezzi et al. 2020). Tck1 knockouts are less infective but have increase replication capacity inside mammalian cells. Mutant cell lines in which a version of eIF2α with the phosphorylated residue threonine 169 was replaced to alanine was described by Castro Machado et al. (2020). Compared to wild-type cells, the eIF2αT169A mutant parasites produced fewer trypomastigotes with lower infection capacity (Castro Machado et al. 2020). Despite several attempts, Chiurillo et al. (2021) were only able to generate parasites single knockout to the T. cruzi AGC essential kinase 1 (TcAEK1) (Chiurillo et al. 2021). The TcAEK1-/+ mutant parasites displayed impaired capacity to invade and replicate in host cells. Significant advances in the study of T. cruzi mitochondrial metabolism have occurred thanks to results obtained from gene manipulation using CRISPR/Cas9 technology targeting sequences encoding mitochondrial proteins. Lander et al. (2018) generated parasite knockout of the pyruvate dehydrogenase phosphatase (TcPDP) gene and showed that the TcPDP mutants have impaired metacyclogenesis and impaired capacity to infect cells in vitro (Lander et al. 2018). Disruption of two gene encoding subunits of the mitochondrial calcium uniporter complex, TcMCU1 and TcMCU2 described by Bertolini et al. (2019) and Chiurillo et al. (2019), also resulted in impaired capacity to infect cells in vitro, as well as impaired capacity of amastigotes to replicate inside cells (Bertolini et al. 2019; (Chiurillo et al. 2019). Negreiros et al. (2021) showed that the disruption of the mitochondrial pyruvate carriers 1 and 2 (TcMPC1 and TcMPC2) resulted in a threefold decrease of the in vitro infectivity of trypomastigotes, as compared to infectivity of wild-type parasites (Negreiros et al. 2021). In addition, replication of intracellular amastigotes was also impaired in the TcMPC1 and TcMPC2 mutants. The T. cruzi intracellular ammonium transporter (TcAMT) that localizes to acidic compartments and the mechanosensitive channel (TcMscS) belonging to the superfamily of small-conductance mechanosensitive channels (MscS) were also characterized in T. cruzi using knockout cell lines. Disruption of the TcAMT gene resulted in reduced growth of the epimastigote forms in LIT culture, and a significant reduction in the proliferation of the amastigote forms within the cell (Cruz-Bustos et al. 2018). Although no impact in the infectivity of TcMscS knockout parasites at early time points post-infection was observed, a significant decrease in the number

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of intracellular amastigotes 48 hours post-infection as well as the number of trypomastigotes collected in the tissue culture supernatant of cells occurred, indicating that knockout of TcMscS impacts replication of amastigotes (Dave et al. 2021). The ability to infect and to replicate inside cells in vitro were also affected in knockout cells lines encoding the Golgi-localized Mn2+-Ca2+/H+ exchanger (TcGDT1) (Ramakrishnan et al. 2021) and the inositol 1,4,5-trisphosphate (IP3) receptor (Chiurillo et al. 2020). It was also observed that cells infected with TcGDT1 knockout released less trypomastigotes in the culture supernatant compared to wild-type parasites. The control of cell cycle and parasite differentiation have been also investigated in recent studies such as the one published by Teixeira et al. (2022). Seeking to investigate the role of the P21, an immunomodulatory T. cruzi protein, the authors observed that P21 knockout parasites have reduced invasion capacity in vitro, compared to WT parasites (Teixeira et al. 2022). Besides presenting reduced rates of epimastigote growth, the P21 knockout epimastigote mutants have delayed cell cycle progression, with mutant parasites accumulating in the G1 phase. However, P21 knockout mutants were able to differentiate into metacyclic trypomastigotes. SaenzGarcia et al. (2021) generated T. cruzi cell lines knockout of TcTrypanin, a gene first described in T. brucei, that encodes a protein that is part of the dynein regulatory complex (Saenz-Garcia et al. 2021). TcTrypanin knockout epimastigotes showed reduced metacyclogenesis rates as well as reduced rates of in vitro infection compared to wild-type and addback trypomastigotes. In another study Alves et al. (2021) characterized the MyoF protein, which is part of the cytostome-cytopharynx complex (SPC) and promotes endocytosis in epimastigotes. Using CRISPR/Ca9 to generate cell lines with the MyoF gene fused to the mNeonGreen, these authors showed that MyoF signal disappeared during G2 phase and reappeared at early cytokinesis (Alves et al. 2022). This study also revealed a role during metacyclogenesis since a progressive disappearance of the SPC was observed and the MyoF signal became absent in metacyclic trypomastigotes. As the first study that directly addresses the role of one of the most important virulence determinants in the T. cruzi infection, Burle-Caldas et al. (2022) used CRISPR/Cas9 to generate knockout cell lines in which several copies of transsialidases (TS) genes were disrupted (Burle-Caldas et al. 2022). Encoded by more than 1000 genes, only a sub-set of members of this large gene family of surface proteins has enzymatic activity. Active TS catalyzes the transfer of sialic acid from host glycoconjugates to mucins at the surface of parasite (da Costa et al. 2021; Schenkman et al. 1994). After identifying all gene copies encoding active transsialidases and designing sgRNAs targeting these sequences, mutant cell lines with no TS activity (TSKO7) were tested using in vitro and in vivo infection assays. Although no differences in cell invasion capacity, these mutants showed a strong reduction in the number of trypomastigotes that are released in culture supernatant compared to control parasites. It was also demonstrated that active TS knockout parasites are largely attenuated and completely unable to infect even immunodeficient mice (IFN-γ knockout mice). The attenuated TS knockout parasites were used to immunize BALB/C mice, which were fully protected against a challenge with a

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virulent T. cruzi strain. Because it is a multicopy gene family, knocking out active trans-sialidases was only possible after the advent of CRIRS/Cas9 technology. Besides challenging earlier studies which have suggested that TS is a key element for cell invasion and parasite escape from the parasitophorous vacuole (Rubin-deCelis et al. 2006; Schenkman et al. 1991), this study not only showed that absence of TS activity causes a severe impairment of the parasite egress from the host cell but also paved the way for the development of new vaccine strategies for Chagas disease based on attenuated mutant parasites.

References Alves AA, Alcantara CL, Dantas-Jr MVA, Sunter JD, De Souza W, Cunha ESNL (2022) Dynamics of the orphan myosin MyoF over Trypanosoma cruzi life cycle and along the endocytic pathway. Parasitol Int. 86:102444. https://doi.org/10.1016/j.parint.2021.102444 Araujo PR, Teixeira SM (2011) Regulatory elements involved in the post-transcriptional control of stage-specific gene expression in Trypanosoma cruzi: a review. Mem Inst Oswaldo Cruz. 106(3):257–266. https://doi.org/10.1590/s0074-02762011000300002 Araujo PR, Burle-Caldas GA, Silva-Pereira RA, Bartholomeu DC, Darocha WD, Teixeira SM (2011) Development of a dual reporter system to identify regulatory cis-acting elements in untranslated regions of Trypanosoma cruzi mRNAs. Parasitol Int. 60(2):161–169. https://doi. org/10.1016/j.parint.2011.01.006 Bertolini MS, Chiurillo MA, Lander N, Vercesi AE, Docampo R (2019) MICU1 and MICU2 play an essential role in mitochondrial Ca(2+) uptake, growth, and infectivity of the human pathogen Trypanosoma cruzi. mBio. 10(3). https://doi.org/10.1128/mBio.00348-19 Brener Z (1973) Biology of Trypanosoma cruzi. Annu Rev Microbiol. 27:347–382. https://doi.org/ 10.1146/annurev.mi.27.100173.002023 Burle-Caldas Gde A, Grazielle-Silva V, Laibida LA, DaRocha WD, Teixeira SM (2015) Expanding the tool box for genetic manipulation of Trypanosoma cruzi. Mol Biochem Parasitol. 203(1-2): 25–33. https://doi.org/10.1016/j.molbiopara.2015.10.004 Burle-Caldas GA, Grazielle-Silva V, Soares-Simoes M, Schumann Burkard G, Roditi I, DaRocha WD et al (2017) Editing the Trypanosoma cruzi genome with zinc finger nucleases. Mol Biochem Parasitol. 212:28–32. https://doi.org/10.1016/j.molbiopara.2017.01.002 Burle-Caldas GA, Soares-Simoes M, Lemos-Pechnicki L, DaRocha WD, Teixeira SMR (2018) Assessment of two CRISPR-Cas9 genome editing protocols for rapid generation of Trypanosoma cruzi gene knockout mutants. Int J Parasitol. 48(8):591–596. https://doi.org/10. 1016/j.ijpara.2018.02.002 Burle-Caldas GA, Dos Santos NSA, de Castro JT, Mugge FLB, Grazielle-Silva V, Oliveira AER et al (2022) Disruption of active trans-sialidase genes impairs egress from mammalian host cells and generates highly attenuated Trypanosoma cruzi parasites. mBio:e0347821. https://doi.org/ 10.1128/mbio.03478-21 Cardoso MS, Junqueira C, Trigueiro RC, Shams-Eldin H, Macedo CS, Araujo PR et al (2013) Identification and functional analysis of Trypanosoma cruzi genes that encode proteins of the glycosylphosphatidylinositol biosynthetic pathway. PLoS Negl Trop Dis 7(8):e2369. https:// doi.org/10.1371/journal.pntd.0002369 Castro Machado F, Bittencourt-Cunha P, Malvezzi AM, Arico M, Radio S, Smircich P et al (2020) EIF2alpha phosphorylation is regulated in intracellular amastigotes for the generation of infective Trypanosoma cruzi trypomastigote forms. Cell Microbiol. 22(11):e13243. https:// doi.org/10.1111/cmi.13243

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Chiurillo MA, Lander N, Bertolini MS, Vercesi AE, Docampo R (2019) Functional analysis and importance for host cell infection of the Ca(2+)-conducting subunits of the mitochondrial calcium uniporter of Trypanosoma cruzi. Mol Biol Cell. 30(14):1676–1690. https://doi.org/ 10.1091/mbc.E19-03-0152 Chiurillo MA, Lander N, Vercesi AE, Docampo R (2020) IP3 receptor-mediated Ca(2+) release from acidocalcisomes regulates mitochondrial bioenergetics and prevents autophagy in Trypanosoma cruzi. Cell Calcium. 92:102284. https://doi.org/10.1016/j.ceca.2020.102284 Chiurillo MA, Jensen BC, Docampo R (2021) Drug target validation of the protein kinase AEK1, essential for proliferation, host cell invasion, and intracellular replication of the human pathogen Trypanosoma cruzi. Microbiol Spectr. 9(2):e0073821. https://doi.org/10.1128/Spectrum. 00738-21 Cong L, Ran FA, Cox D, Lin S, Barretto R, Habib N et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science. 339(6121):819–823. https://doi.org/10.1126/science. 1231143 Cooper R, de Jesus AR, Cross GA (1993) Deletion of an immunodominant Trypanosoma cruzi surface glycoprotein disrupts flagellum-cell adhesion. J Cell Biol. 122(1):149–156. https://doi. org/10.1083/jcb.122.1.149 Costa FC, Francisco AF, Jayawardhana S, Calderano SG, Lewis MD, Olmo F et al (2018) Expanding the toolbox for Trypanosoma cruzi: a parasite line incorporating a bioluminescence-fluorescence dual reporter and streamlined CRISPR/Cas9 functionality for rapid in vivo localisation and phenotyping. PLoS Negl Trop Dis. 12(4):e0006388. https://doi. org/10.1371/journal.pntd.0006388 Cruz-Bustos T, Potapenko E, Storey M, Docampo R (2018) An intracellular ammonium transporter is necessary for replication, differentiation, and resistance to starvation and osmotic stress in Trypanosoma cruzi. mSphere. 3(1). https://doi.org/10.1128/mSphere.00377-17 Currier RB, Ulrich K, Leroux AE, Dirdjaja N, Deambrosi M, Bonilla M et al (2019) An essential thioredoxin-type protein of Trypanosoma brucei acts as redox-regulated mitochondrial chaperone. PLoS Pathog. 15(9):e1008065. https://doi.org/10.1371/journal.ppat.1008065 da Costa KM, Marques da Fonseca L, Dos Reis JS, Santos M, Previato JO, Mendonca-Previato L et al (2021) Trypanosoma cruzi trans-sialidase as a potential vaccine target against chagas disease. Front Cell Infect Microbiol. 11:768450. https://doi.org/10.3389/fcimb.2021.768450 DaRocha WD, Otsu K, Teixeira SM, Donelson JE (2004a) Tests of cytoplasmic RNA interference (RNAi) and construction of a tetracycline-inducible T7 promoter system in Trypanosoma cruzi. Mol Biochem Parasitol. 133(2):175–186. https://doi.org/10.1016/j.molbiopara.2003.10.005 DaRocha WD, Silva RA, Bartholomeu DC, Pires SF, Freitas JM, Macedo AM et al (2004b) Expression of exogenous genes in Trypanosoma cruzi: improving vectors and electroporation protocols. Parasitol Res. 92(2):113–120. https://doi.org/10.1007/s00436-003-1004-5 Dave N, Cetiner U, Arroyo D, Fonbuena J, Tiwari M, Barrera P et al (2021) A novel mechanosensitive channel controls osmoregulation, differentiation, and infectivity in Trypanosoma cruzi. Elife 10. https://doi.org/10.7554/eLife.67449 Doudna JA, Charpentier E, Genome editing. (2014) The new frontier of genome engineering with CRISPR-Cas9. Science. 346(6213):1258096. https://doi.org/10.1126/science.1258096 Durai S, Mani M, Kandavelou K, Wu J, Porteus MH, Chandrasegaran S (2005) Zinc finger nucleases: custom-designed molecular scissors for genome engineering of plant and mammalian cells. Nucleic Acids Res. 33(18):5978–5990. https://doi.org/10.1093/nar/gki912 El-Sayed NM, Myler PJ, Bartholomeu DC, Nilsson D, Aggarwal G, Tran AN et al (2005) The genome sequence of Trypanosoma cruzi, etiologic agent of Chagas disease. Science. 309(5733): 409–415. https://doi.org/10.1126/science.1112631 Glover L, Jun J, Horn D (2011) Microhomology-mediated deletion and gene conversion in African trypanosomes. Nucleic Acids Res. 39(4):1372–1380. https://doi.org/10.1093/nar/gkq981 Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science. 337(6096): 816–821. https://doi.org/10.1126/science.1225829

124

G. de Assis Burle-Caldas et al.

Kangussu-Marcolino MM, de Paiva RM, Araujo PR, de Mendonca-Neto RP, Lemos L, Bartholomeu DC et al (2013) Distinct genomic organization, mRNA expression and cellular localization of members of two amastin sub-families present in Trypanosoma cruzi. BMC Microbiol. 13:10. https://doi.org/10.1186/1471-2180-13-10 Kim S, Kim D, Cho SW, Kim J, Kim JS (2014) Highly efficient RNA-guided genome editing in human cells via delivery of purified Cas9 ribonucleoproteins. Genome Res. 24(6):1012–1019. https://doi.org/10.1101/gr.171322.113 Lander N, Li ZH, Niyogi S, Docampo R (2015) CRISPR/Cas9-induced disruption of paraflagellar rod protein 1 and 2 genes in Trypanosoma cruzi reveals their role in flagellar attachment. mBio. 6(4):e01012. https://doi.org/10.1128/mBio.01012-15 Lander N, Chiurillo MA, Storey M, Vercesi AE, Docampo R (2016) CRISPR/Cas9-mediated endogenous C-terminal tagging of Trypanosoma cruzi genes reveals the acidocalcisome localization of the inositol 1,4,5-trisphosphate receptor. J Biol Chem. 291(49):25505–25515. https:// doi.org/10.1074/jbc.M116.749655 Lander N, Chiurillo MA, Bertolini MS, Storey M, Vercesi AE, Docampo R (2018) Calciumsensitive pyruvate dehydrogenase phosphatase is required for energy metabolism, growth, differentiation, and infectivity of Trypanosoma cruzi. J Biol Chem. 293(45):17402–17417. https://doi.org/10.1074/jbc.RA118.004498 Li X, Heyer WD (2008) Homologous recombination in DNA repair and DNA damage tolerance. Cell Res. 18(1):99–113. https://doi.org/10.1038/cr.2008.1 Malvezzi AM, Arico M, Souza-Melo N, Dos Santos GP, Bittencourt-Cunha P, Holetz FB et al (2020) GCN2-like kinase modulates stress granule formation during nutritional stress in Trypanosoma cruzi. Front Cell Infect Microbiol. 10:149. https://doi.org/10.3389/fcimb.2020. 00149 Negreiros RS, Lander N, Chiurillo MA, Vercesi AE, Docampo R (2021) Mitochondrial pyruvate carrier subunits are essential for pyruvate-driven respiration, infectivity, and intracellular replication of Trypanosoma cruzi. mBio 12(2). https://doi.org/10.1128/mBio.00540-21 Nittolo AG, Banuelos CP, Saborit JI, Tekiel V, Sanchez DO, Levy GV (2018) TbRRM1 knockdown produces abnormal cell morphology and apoptotic-like death in the bloodstream form of T. brucei. Mol Biochem Parasitol. 224:1–5. https://doi.org/10.1016/j.molbiopara.2018.07.006 Passos-Silva DG, Rajao MA, Nascimento de Aguiar PH, Vieira-da-Rocha JP, Machado CR, Furtado C (2010) Overview of DNA Repair in Trypanosoma cruzi, Trypanosoma brucei, and Leishmania major. J Nucleic Acids 2010:840768. https://doi.org/10.4061/2010/840768 Pavani RS, de Lima LP, Lima AA, Fernandes CAH, Fragoso SP, Calderano SG et al (2020) Nuclear export of replication protein A in the nonreplicative infective forms of Trypanosoma cruzi. FEBS Lett. 594(10):1596–1607. https://doi.org/10.1002/1873-3468.13755 Peng D, Kurup SP, Yao PY, Minning TA, Tarleton RL (2014) CRISPR-Cas9-mediated single-gene and gene family disruption in Trypanosoma cruzi. mBio. 6(1):e02097–e02014. https://doi.org/ 10.1128/mBio.02097-14 Perez-Diaz L, Correa A, Moretao MP, Goldenberg S, Dallagiovanna B, Garat B (2012) The overexpression of the trypanosomatid-exclusive TcRBP19 RNA-binding protein affects cellular infection by Trypanosoma cruzi. Mem Inst Oswaldo Cruz. 107(8):1076–1079. https://doi.org/ 10.1590/s0074-02762012000800021 Perez-Molina JA, Molina I (2018) Chagas disease. Lancet. 391(10115):82–94. https://doi.org/10. 1016/S0140-6736(17)31612-4 Pezza A, Tavernelli LE, Alonso VL, Perdomo V, Gabarro R, Prinjha R et al (2022) Essential bromodomain TcBDF2 as a drug target against chagas disease. ACS Infect Dis. 8(5): 1062–1074. https://doi.org/10.1021/acsinfecdis.2c00057 Picchi-Constante GFA, Hiraiwa PM, Marek M, Rogerio VZ, Guerra-Slompo EP, Romier C et al (2022) Efficient CRISPR-Cas9-mediated genome editing for characterization of essential genes in Trypanosoma cruzi. STAR Protoc. 3(2):101324. https://doi.org/10.1016/j.xpro.2022.101324 Ramakrishnan S, Unger LM, Baptista RP, Cruz-Bustos T, Docampo R (2021) Deletion of a Golgi protein in Trypanosoma cruzi reveals a critical role for Mn2+ in protein glycosylation needed

7

CRISPR Genome Editing and the Study of Chagas Disease

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for host cell invasion and intracellular replication. PLoS Pathog. 17(3):e1009399. https://doi. org/10.1371/journal.ppat.1009399 Regis-da-Silva CG, Freitas JM, Passos-Silva DG, Furtado C, Augusto-Pinto L, Pereira MT et al (2006) Characterization of the Trypanosoma cruzi Rad51 gene and its role in recombination events associated with the parasite resistance to ionizing radiation. Mol Biochem Parasitol. 149(2):191–200. https://doi.org/10.1016/j.molbiopara.2006.05.012 Rubin-de-Celis SS, Uemura H, Yoshida N, Schenkman S (2006) Expression of trypomastigote trans-sialidase in metacyclic forms of Trypanosoma cruzi increases parasite escape from its parasitophorous vacuole. Cell Microbiol. 8(12):1888–1898. https://doi.org/10.1111/j. 1462-5822.2006.00755.x Saenz-Garcia JL, Borges BS, Souza-Melo N, Machado LV, Miranda JS, Pacheco-Lugo LA et al (2021) Trypanin disruption affects the motility and infectivity of the protozoan Trypanosoma cruzi. Front Cell Infect Microbiol. 11:807236. https://doi.org/10.3389/fcimb.2021.807236 Schenkman S, Jiang MS, Hart GW, Nussenzweig V (1991) A novel cell surface trans-sialidase of Trypanosoma cruzi generates a stage-specific epitope required for invasion of mammalian cells. Cell. 65(7):1117–1125. https://doi.org/10.1016/0092-8674(91)90008-m Schenkman S, Eichinger D, Pereira ME, Nussenzweig V (1994) Structural and functional properties of Trypanosoma trans-sialidase. Annu Rev Microbiol. 48:499–523. https://doi.org/10.1146/ annurev.mi.48.100194.002435 Schumann Burkard G, Jutzi P, Roditi I (2011) Genome-wide RNAi screens in bloodstream form trypanosomes identify drug transporters. Mol Biochem Parasitol. 175(1):91–94. https://doi.org/ 10.1016/j.molbiopara.2010.09.002 Soares Medeiros LC, South L, Peng D, Bustamante JM, Wang W, Bunkofske M et al (2017) Rapid, selection-free, high-efficiency genome editing in protozoan parasites using CRISPR-Cas9 ribonucleoproteins. mBio. 8(6). https://doi.org/10.1128/mBio.01788-17 Tavares TS, Mugge FLB, Grazielle-Silva V, Valente BM, Goes WM, Oliveira AER et al (2021) A Trypanosoma cruzi zinc finger protein that is implicated in the control of epimastigote-specific gene expression and metacyclogenesis. Parasitology. 148(10):1171–1185. https://doi.org/10. 1017/S0031182020002176 Taylor MC, Huang H, Kelly JM (2011) Genetic techniques in Trypanosoma cruzi. Adv Parasitol. 75:231–250. https://doi.org/10.1016/B978-0-12-385863-4.00011-3 Teixeira TL, Chiurillo MA, Lander N, Rodrigues CC, Onofre TS, Ferreira ER et al (2022) Ablation of the P21 gene of Trypanosoma cruzi provides evidence of P21 as a mediator in the control of epimastigote and intracellular amastigote replication. Front Cell Infect Microbiol. 12:799668. https://doi.org/10.3389/fcimb.2022.799668 Unnikrishnan M, Burleigh BA (2004) Inhibition of host connective tissue growth factor expression: a novel Trypanosoma cruzi-mediated response. FASEB J. 18(14):1625–1635. https://doi.org/ 10.1096/fj.04-1554com Vazquez MP, Levin MJ (1999) Functional analysis of the intergenic regions of TcP2beta gene loci allowed the construction of an improved Trypanosoma cruzi expression vector. Gene. 239(2): 217–225. https://doi.org/10.1016/s0378-1119(99)00386-8 Zikova A, Schnaufer A, Dalley RA, Panigrahi AK, Stuart KD (2009) The F(0)F(1)-ATP synthase complex contains novel subunits and is essential for procyclic Trypanosoma brucei. PLoS Pathog. 5(5):e1000436. https://doi.org/10.1371/journal.ppat.1000436

Chapter 8

Genome Editing Tools for Lysosomal Storage Disorders Esteban Alberto Gonzalez, Helena Nader, Marina Siebert, Diego A. Suarez, Carlos J. Alméciga-Díaz, and Guilherme Baldo

1 Lysosomes: A Brief Overview In cell biology, lysosomes play a major role in the breakdown and recycling of extracellular particles and intracellular components. These intracellular membranebound organelles are dedicated to catabolism function through the activity of more than 60 hydrolytic enzymes inside their lumen (Futerman and Van Meer 2004; Fernández-Pereira et al. 2021). Those soluble lysosomal hydrolases are capable of degrading, under acidic conditions, a variety of macromolecules, including proteins,

E. A. Gonzalez · G. Baldo (✉) Cell, Tissue and Gene Laboratory, Experimental Research Center, Hospital de Clínicas de Porto Alegre, Porto Alegre, RS, Brazil Postgraduate Program in Genetics and Molecular Biology, Universidade Federal do Rio Grande do Sul, Porto Alegre, RS, Brazil e-mail: [email protected] H. Nader Departamento de Bioquímica, Instituto de Farmacologia e Biologia Molecular, Escola Paulista de Medicina, Universidade Federal de São Paulo, São Paulo, São Paulo, Brazil M. Siebert Postgraduate Program in Sciences of Gastroenterology and Hepatology, Universidade Federal do Rio Grande do Sul, Porto Alegre, RS, Brazil Basic Research and Advanced Investigations in Neurosciences Laboratory, Experimental Research Center, Hospital de Clínicas de Porto Alegre, Porto Alegre, RS, Brazil Unit of Laboratorial Research, Experimental Research Center, Hospital de Clínicas de Porto Alegre, Porto Alegre, RS, Brazil D. A. Suarez · C. J. Alméciga-Díaz Institute for the Study of Inborn Errors of Metabolism, Faculty of Science, Pontificia Universidad Javeriana, Bogotá, Colombia © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_8

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carbohydrates, lipids, and nucleic acids. Once metabolized, the digested products (e.g., monosaccharides and amino acids, free fatty acids) are transported out of the lysosome for energy homeostasis or reutilization in biosynthetic pathways (Xu and Ren 2015). All lysosomal enzymes are produced in the cytoplasm and migrate along the biosynthetic route to the lysosomes managed by a specialized system of sorting signals (Braulke and Bonifacino 2009). Most hydrolytic enzymes are targeted to lysosomes through the addition of mannose-6-phosphate (M6P) residues onto the oligosaccharide moieties. This modification takes place in the late Golgi compartment, where they are also recognized and packaged in transport vesicles. However, some specific enzymes do not depend on the M6P signal for lysosomal delivery such as β-glucocerebrosidase, which is transported by lysosomal integral membrane protein 2 (LIMP2) (Parenti et al. 2015). Besides the interaction of hydrolases and substrates, the lysosomal function requires acidification machinery, membrane proteins (e.g., transporters, membrane-bound enzymes, pump protons), and enzyme activators. Thus, several lysosomal and non-lysosomal proteins contribute to maintaining lysosomal function. Defects in degradation, catabolite export, or trafficking result in lysosomal dysfunction and, as expected, to drastic consequences for cells and tissues homeostasis, and ultimately multisystemic disease.

2 Lysosomal Storage Disorders Lysosomal storage disorders (LSDs) comprise a heterogeneous group of more than 50 monogenic disorders caused by defects in some components of the lysosomal system. They are caused by pathogenic alterations in genes encoding lysosomal hydrolases, integral membrane proteins, enzyme modifiers, and activator or transporters proteins (Futerman and Van Meer 2004; Platt et al. 2018). Deficiencies of these proteins affect the catabolic process, and specific macromolecules or monomeric compounds accumulate inside lysosomes. Over time these anomalies trigger a pathogenic cascade causing cell dysfunction and death and tissue damage (Platt et al. 2012). According to the type of storage, LSDs are grouped into broad categories and includes the sphingolipidoses, oligosaccharidoses, mucolipidoses, mucopolysaccharidoses, and neuronal ceroid lipofuscinoses (Poswar et al. 2019; Giugliani et al. 2017). For example, an impairment in the catabolism of sphingolipids causes sphingolipidoses such as Fabry, Krabbe, and Gaucher diseases or metachromatic leukodystrophy. Otherwise, defect in some steps of glycosaminoglycans degradation results in mucopolysaccharidoses I–IX (MPS I–IX). Most of these disorders are inherited in an autosomal recessive manner with the exception of Fabry disease and mucopolysaccharidosis type II (also called Hunter syndrome) which are both X-linked, and Danon disease which is inherited in an Xlinked-dominant form (Solomon and Muro 2017). Individually, LSDs are rare disorders; some of them are even considered ultra-rare disorders due to the low number of cases described. As an example, MPS VII is an ultra-rare LSD with an

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estimated worldwide birth prevalence of less than 1:1,000,000 (Giugliani et al. 2021; Federhen et al. 2020). Nevertheless, collectively these are common disorders with a combined prevalence estimated at 1:8000 live births. This number, however, may be underestimated since in many cases patients are not diagnosed because the disease presentation can be nonspecific and signs and symptoms are often mistakenly attributed to other disorders (Fuller et al. 2006). Moreover, some particular populations have higher prevalence of certain LSD due to diverse types of barriers (e.g., geographic, ethnicities) that prevent gene flow between them or even by high consanguineous marriage rates among individuals. This is observed, for example, in the Ashkenazi Jews population where the prevalence of Gaucher disease goes up to 1:855 births (Fuller et al. 2006). Similarly, mucopolysaccharidosis VI is found in the general population at a prevalence of 1 in 500,000 births (Baehner et al. 2005), but an isolated community in northeast Brazil has over 1 in 5000 live births (CostaMotta et al. 2014). Most LSDs are caused by deficient activity of a specific soluble lysosomal enzyme. Some pathogenic variants in the genes that encode these enzymes render them inactive. Other pathogenic variants generate active enzymes that may fail to fold correctly and, consequently, are degraded (Solomon and Muro 2017). However, some variants may cause a reduction in the enzymatic activity. As a norm, the residual enzyme activity is closely related to severity of phenotype (Vellodi 2005). For example, in the mucopolysaccharidoses I (MPS I), the complete deficiency in the lysosomal enzyme α-L-iduronidase (IDUA) results in the severe form of the MPS I (Hurler syndrome), while a partial deficiency leads to a continuum spectrum of a less aggressive disease (Scheie syndrome or Hurler-Scheie syndrome). Lysosomal storage may also be caused by alterations in nonenzymatic lysosomal proteins or even non-lysosomal proteins (enzymatic or not) but involved with lysosomal function. In this context, mucolipidosis type II is caused by the deficiency or absence of the enzyme, 6-phospho-N-acetyl-glucosamine transferase, which is involved in the mechanism of lysosome sorting through the M6P marker. As a result, multiple acid hydrolases cannot be transported to the lysosome, but instead are secreted from the cells, and many different substrates are not degraded (Solomon and Muro 2017; Vellodi 2005; Alroy and Lyons 2014). Defects in some integral membrane proteins of lysosomes lead to disruption in metabolites transport out of lysosomes. This is observed in Niemann-Pick type C (NPC), whose defects in membrane transporter of lysosomes lead to failures in cholesterol transport and, consequently, lysosomal storage. Defects in the integral lysosomal membrane protein, LAMP2A, can result in Danon disease (Futerman and Van Meer 2004). Some other LSDs are caused by defects in nonenzymatic cofactors, which assist acid hydrolases in the substrate degradation process. Lysosomal degradation of sphingolipids by hydrolases, for example, requires the presence of activator proteins (saposins) for catabolic function (Vellodi 2005; Alroy and Lyons 2014). Likewise, an impairment in these indispensable factors affect the catabolism of sphingolipids, leading to atypical forms of Krabbe disease (Sap A), metachromatic leukodystrophy (Sap B), or Gaucher’s disease (Sap C) (Vellodi 2005; Alroy and Lyons 2014; Shaimardanova et al. 2020; Kuchař et al. 2009).

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Progressive accumulation of undigested materials leads to enlargement and dysfunctional lysosomes. When very high levels of substrates accumulate, they may compromise the activity of other proteins not related to genetic defects resulting in secondary substrate accumulation (Platt et al. 2012). For example, in NiemannPick disease type C1 (caused by defects in NPC1 gene), sphingosine storage causes altered calcium homeostasis in lysosomes leading to secondary storage of cholesterol and sphingolipid (Lloyd-Evans et al. 2008). The accumulation of primary and secondary substrates triggers a cascade of events that impacts on cell function and eventually lead to cell death and tissue damage (Platt et al. 2012; Parenti et al. 2013). In general, the common pathways dysregulated in LSDs include defects in cellular transport and degradation process, calcium homeostasis, oxidative stress, and inflammatory response (Platt et al. 2018). The permeability and integrity of the lysosomal membrane may also be compromised in LSDs, leading to leakage of lysosomal content to the cytosol (Futerman and Van Meer 2004; Pereira et al. 2010). This lysosomal perturbation may cause release of proteases and other lysosomal hydrolases, which in turn results in degradation of several cellular components (Boya and Kroemer 2008). In MPS I, the glycosaminoglycans storage causes alterations in Ca+2 and H+ homeostasis and lysosomal leakage of cysteine proteases such as cathepsins (Pereira et al. 2010; Gonzalez et al. 2018). Consequently, the increase of cathepsin B activity may lead to the cleavage of extracellular matrix components and contributes to the physiopathology of MPS I (Gonzalez et al. 2018; Baldo et al. 2017). Many factors appear to play a key role in the pathophysiology of LSDs through complex mechanisms that are still not fully understood. LSDs are clinically heterogeneous disorders, but they generally have a progressive multisystemic course, including peripheral organs and neurological manifestations (Futerman and Van Meer 2004; Platt et al. 2018; Parenti et al. 2013). Peripheral symptoms include spleen and liver enlargement (hepatosplenomegaly), heart and kidney damage, skeletal alterations, muscle atrophy, and ocular manifestations (Futerman and Van Meer 2004). The broad spectrum of clinical manifestations depends on several parameters, such as the levels of residual enzyme activity, pathway affected, substrate accumulated, and site of storage (Platt et al. 2012; Solomon and Muro 2017; Sun 2018). For instance, in Krabbe disease and metachromatic leukodystrophy, the impairment in the degradation of certain lipids results in the destruction of the myelin sheath (demyelination) of the nervous system (Shaimardanova et al. 2020; Feltri et al. 2021). The age of clinical onset also varies among different LSDs (Platt et al. 2012). In general, most LSDs affect children (early-onset), but adult forms (late-onset) also occur (Platt et al. 2012; Solomon and Muro 2017; Wraith 2002). The lower the residual enzyme activity, the earlier the age of onset, and the more severe the disease. Almost all LSDs have neurological involvement, which may manifest in all patients within a particular enzyme deficiency or be limited to the most severe form of the disease. This last is seen in the severe form of MPS I from the milder MPS I. This is also the case for Gaucher disease (β-glucocerebrosidase deficiency) types II and III that have CNS impairment, while type I Gaucher disease is a non-neuropathic disorder (Vellodi 2005). Alternatively, in Niemann-Pick type C disease, all patients develop neurological or

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psychiatric symptoms (Sévin et al. 2007). Even though several LSDs are characterized by prominent neurological involvement and minimal peripheral impairment (e.g., Sanfilippo disease), other conditions have peripheral dysfunction with rare brain involvement (e.g., Fabry disease) (Futerman and Van Meer 2004). Otherwise, less severe LSDs could manifest in childhood, progressing slowly into adulthood, causing mainly peripheral disease, such as liver, spleen, heart, striated muscle, kidneys, lungs, bone, connective tissue, and immune system damage (Solomon and Muro 2017). The past 30 years have been marked by continued efforts to understand the pathologic molecular mechanism around LSDs and by the development of therapies. The first well-established treatment for some LSDs involved the intravenous administration of a purified enzyme or, subsequently, the transplantation of hematopoietic stem cells capable of producing and secreting normal enzymes. This normal protein, tagged with the M6P signal, may be taken up by neighbor cells and targeted to lysosomes. Basically, these therapeutic approaches aim to increase the activity of the defective enzyme or protein. Despite its effectiveness and advantages, both treatments are available for some LSDs and show certain limitations. In recent years, the development of new tools for genetic manipulations has allowed the creation of biological models for the understanding of the pathophysiology of these disorders, as well as new therapeutic options based on gene editing have emerged, which we will discuss in this chapter.

3 Genome Editing Technologies Genome editing technologies comprise a variety of useful tools to accurately modify a DNA sequence through the deletion of disease-causing genes as a whole or part of them, correction of disease-causing mutations due to insertions, or even base pair conversion at a specific location of the reference genome. This approach allows the manipulation of any gene in an extended range of cells, tissues, and even organisms (Gaj et al. 2013). Nowadays, many different approaches are available for genome editing, such as transcription activator-like effector nucleases (TALENs), zinc finger nucleases (ZFNs), clustered regularly interspaced short palindromic repeat (CRISPR)-associated (Cas) systems, CRISPR-Cas9-base editors, and prime editing. All of them are based on the same principle that involves the use of a nuclease enzyme to create double-strand breaks (DSBs) in a determined location of the target DNA to trigger endogenous DNA repair mechanisms. The effect on the gene of interest will depend on the type of recombination being stimulated. In cases in which nonhomologous end joining are involved, it usually leads to a gene disruption due to an insertion or deletion of nucleotides in a random manner, and it is commonly known as error-prone. On the other hand, the homologous-directed repair mechanism can result in a correction of the target gene driven by a template, and it is considered error-free (Bétermier et al. 2014). The main difference among the genome editing platforms is the way each one uses to recognize the target DNA

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sequence. Both ZFNs and TALENs are chimeric nucleases constituted by two parts, which are the nonspecific DNA cleavage domain (formed by the FokI restriction enzyme) and the sequence-specific DNA-binding domain. The domain that binds to the DNA may be customized to target the desired genome sequence (Gaj et al. 2013). The genome editing techniques that integrate an endonuclease (typically Cas9) and a guide-RNA (gRNA) are known as CRISPR-associated (Cas) systems. The primary function of the gRNA is to orientate the cleavage of DNA by Cas9. The classic CRISPR-Cas9 methodology was based on DSBs in a specific DNA location which is mostly repaired by nonhomologous end joining or to a lesser predominant homologous-directed repair. Nevertheless, recent advances in the CRISPR techniques have allowed the occurrence of genome editing using a pathway independent of DSBs. Base editing and prime editors are both results of this evolution, being base editors used to substitute a target base or a base pair to another by altering Cas9 (inactive Cas9 or Cas9 Nickase) in a combination with base-modifying enzymes (cytosine deaminase or adenosine deaminase). On the other hand, the prime editor method employs a reverse transcriptase enzyme coupled to a Cas9 Nickase along with a prime editing guide RNA (pegRNA) (Anzalone et al. 2020). The genome editing systems encounter difficulties in crossing cell membranes due to their molecular size. To solve that, several specialized carriers to overcome membrane barriers and facilitate gene or protein transfer into the cells have been developed. These methods include viral or nonviral vectors and even chemical and physical techniques. But not only is it important to make the target cells accessible to be reached by genome editing systems, the delivery method (ex vivo or in vivo) must also be defined in advance. An in vivo approach involves the administration of the genome editing tool using a straightforward injection in the body. In this way, genome editing takes place within the subject. Contrary, an ex vivo approach comprises the retrieval of the target cells from host organisms, followed by an in vitro modification before the reinjection of the edited cells into the body (Poletto et al. 2020).

3.1

Modeling LSDs Using Gene Editing Tools

Genome editing has been used to create disease-cell models, animal models, and turning treatment more specific and viable for rare diseases, such as LSDs. The majority of research on LSDs was classically developed using fibroblasts, although in many of these disorders, it was not the ideal cell model to be used. The simple explanation for that is the easiness to obtain these primary cells from patients and cultivating them. However, some biomedical research needs to be carried out in certain cell-type (e.g., myocytes, neurons) because it can mimic aspects of a biological process or disease found in humans. Nowadays, genome editing tools have come up with an improvement to make studies easier, cheaper, and possible. Induced pluripotent stem cells (iPSCs) and commercially available cell lines have been used in genome editing studies for those purposes.

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Patient-derived fibroblasts or blood cells can be reprogrammed in vitro before being separated into the target cell type to create iPSC models. Genome editing in iPSCs can be used to generate a specific disease cell model in two ways: genome edit patient’s cells to produce wild-type cells before differentiation protocol or genome edit wild-type cells (healthy) to produce mutant cells with a specific variant in the gene of interest or even multiple distinct genotypes. In both situations, isogenic cells with the same genetic background will be produced. Although there are extensive advantages in using iPSCs to create cellular models of disease, their practical application in research laboratories worldwide is limited due to their high cost of implementation and execution and maintenance. In contrast, cell lines are cheaper, have a high degree of repeatability, and are simple to cultivate. Since they are often easier to transfect, editing them is made simpler, and a wide variety of cell types are simply accessible (Pimentel-Vera et al. 2021). Thus, genome editing tools could be used to create and mimic a disease model (cell or animal), making a wide variety of studies possible in a much easier and feasible way, such as understanding the pathophysiology of a specific disease as well as in the field of drug discovery. Genome editing technology has been applied to develop both cell and animal models for LSDs, and some examples of them will be presented during this section to illustrate the broad utility of these tools for basic research (Table 8.1).

3.1.1

Sphingolipidoses

Fabry disease (FD) is an X-linked disorder caused by enzymatic deficiency of alphagalactosidase A (α-GLA) leading to the buildup of glycosphingolipid, mainly globotriaosylceramide (Gb3) (Politei et al. 2019; Villalobos et al. 2013). As a consequence, multisystem damage is installed, being characterized by cardiac, renal, and cerebrovascular symptoms. The first study using genome editing for modeling a cell model for FD was conducted by Son and colleagues in HEK-293 T cells (immortalized human embryonic kidney cells). They generated a GLA knockout cell line (GLA-KO HEK-293 T) using the CRISPR-Cas9 genome editing tool to allow high-throughput screening of drug candidates that prolong the activity of the recombinant enzyme used as enzyme replacement therapy (ERT) for this disease (Song et al. 2016). Although ERT is available for FD, it is not completely effective in alleviating the suffering of patients (Schiffmann 2006). Neuropathic pain is one of the most debilitating symptoms related to FD, and its pathophysiology is not completely understood (Schuller et al. 2016). There are many cell lines developed using CRISPR-mediated genome editing tools for simulating FD. Kaneski and colleagues developed a Fabry cell model using an immortalized dorsal root ganglion neuron with nociceptive characteristics derived from rat (known as 50B11 cell line) through genome editing. As a result, GLA was knocked out (KO), and a cell model was developed mimicking the phenotypic characteristics of FD. Some of the greatest advantages were the possibility to grow edited cells in standard culture conditions,

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Table 8.1 Summary of cell lines and animal modelling LSDs created using gene editing technologies Disease Fabry disease (FD)

Edited gene GLA

Cell line/ animal model HEK-293 T

50B11

Patient-derived iPSCs

Gaucher disease (GD)

GBA1

THP-1

U87

Main findings A KO cell line was obtained The efficacy of a proteostasis modulator – MG132 on the recombinant enzyme GLA activity MG132 enhanced intracellular half-life of recombinant GLA MG132 potentiated the Gb3 clearance A KO cell line was obtained Severe reduction of lysosomal AGA activity Impaired degradation of Gb3 Stable phenotype over longterm culture Increased levels of TRPV1 pain receptor, suggesting a possible mechanism for increased pain sensitization in Fabry patients Genetic correction reverses the accumulation of LIMP-2, secretion of cathepsin F and HSPA2/HSP70–2 Restoration of α-GLA expression Novel cellular and secreted protein biomarkers with suggested roles in heart disease in FD were identified An isogenic GD model of monocytes was obtained Low levels of acid β-glucosidase expression Massive accumulation of substrate An isogenic GD line of glial cells was obtained Low levels of acid β-glucosidase expression Massive accumulation of substrate Mutant enzyme was retained in the endoplasmic reticulum and subjected to proteasomal degradation, triggering unfolded protein response Increased production of

Reference Song et al. (2016)

Kaneski et al. (2022)

Birket et al. (2019)

Pavan et al. (2020)

Pavan et al. (2020)

(continued)

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Table 8.1 (continued) Disease

Edited gene

Cell line/ animal model

HEK293, A549

gba1; gba2

Zebrafish

gba1; miR-155

Zebrafish

CCR5

Human hematopoietic stem and progenitor cells

Main findings interleukin-1β, both with and without inflammasome activation, alpha-synuclein accumulation, and a higher rate of cell death KO cell lines were obtained Diminished influenza virus infection in KO cell lines Influenza virus trafficking and others cargos were impaired in KO cells GCase is critically important for endocytic trafficking of viruses as well as many cellular cargos, including growth factor receptors Three KO animal models were generated Gba1 (gba1-/-), Gb2 (gba2-/-), and a double KO (gba1-/- gba2-/-). Enzyme deficiency Difference in substrate accumulation Distinctive characteristics of zebrafish Gba2 deficiency were identified Zebrafish larvae offers an attractive model to study glucosidase actions in glycosphingolipid metabolism in vivo A mutant miR-155-/- and a double mutant gba1-/- miR155-/- zebrafish lines were obtained miR-155 is upregulated in several animal models of GF Genetic ablation of miR-155 does not ameliorate neuroinflammation and GD progression Efficient integration of GCase cassettes into the safe harbor locus Targeted cells generate glucocerebroside-expressing macrophages Capacity for long-term engraftment and multi-lineage differentiation

Reference

Drews et al. (2019)

Lelieveld et al. (2019)

Watson et al. (2019)

Scharenberg et al. (2020)

(continued)

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Table 8.1 (continued) Disease Cystinosis

Edited gene Ctns

Cell line/ animal model Rat

Krabbe disease

GALC

Mouse

Niemann-pick type C

NPC1

HAP1

NPC1

HeLa

ncp2

Zebrafish

Metachromatic leukodystrophy

ARSA

Human Schwann cells

GM2 gangliosidosis (Sandhoff disease)

Alb (albumin locus)

Mouse

Main findings A KO animal model was obtained Progressive cystine accumulation Defect in lysosomal activity and autophagy Early-onset, kidney failure, and multisystemic manifestation, as observed in patients Two new animal models carrying point mutations frequently found in infantile and adult form were obtained Decrease in GALC activity Psychosine accumulation (lower to high) Mutations affect life span Neurological phenotype Generation of haploid cell models of NPC1 variants Reduction of NPC1 protein Defect in intracellular cholesterol transport Protocol for producing KO cellular models KO animal was obtained Nile blue staining allow the robust and reliable detection of npc2-/- larvae Downregulation in genes related to calcium homeostasis and myelination Pathological changes in the nervous system, kidney, liver, and pancreas correlated with inflammatory responses A cell model was obtained Sulfatide accumulation affects mitochondrial bioenergetics Metformin treatment reduced the reactive oxygen species generation and improved mitochondrial bioenergetic performance Integration of HEXM into the safe harbor locus Increase in hex enzyme activities (plasma and tissues,

Reference Krohn et al. (2022)

Rebiai et al. (2022)

Erwood et al. (2019)

Du et al. (2017) Wiweger et al. (2021)

SanchezÁlvarez et al. (2022)

Ou et al. (2020a)

(continued)

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Table 8.1 (continued) Disease

Edited gene

Cell line/ animal model

GM1 gangliosidosis

Gbl1

Mouse

Pompe disease

Gaa

Mouse

Sol8

Mucolipidoses

GNPTAB

HeLa

Main findings including the brain) Reduction in GM2 ganglioside levels in multiple tissues (except the brain) Improvements in coordination, motor function, and motor memory Reduced cellular vacuolation in the brain and liver A mouse line carrying the pathogenic variant p. (Gly455Arg) was established Negligible b-galactosidase enzyme activity GM1 ganglioside accumulation Motor dysfunction, activation of microglia, and disturbed autophagy A mouse carrying the pathogenic variant p.(Tyr609*) was established Reduced GAA expression and enzymatic activity Glycogen storage Impaired autophagy Early onset and severe hypertrophic cardiomyopathy Skeletal muscle weakness Isogenic murine GAA-KO cell lines resembling severe mutation found in Pompe patient were obtained Lacked GAA activity Increased autophagy markers and glycogen content Downregulation of mannose 6-phosphate receptors A KO cell line was obtained Swollen lysosomes and lysosomal dysfunction Alteration in others lysosomal enzymes Cathepsin B not properly processed and not detected in lysates of cells Specific residue substitution on based the structure of Drosophila melanogaster GNPTAB homolog was generated, and the

Reference

Liu et al. (2021)

Huang et al. (2020)

AguilarGonzález et al. (2022)

Du et al. (2022)

(continued)

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Table 8.1 (continued) Disease

Edited gene

Cell line/ animal model

MPS II

IDS

SHSY-5Y

MPS III (A, B, C)

Sgsh

Zebrafish

NAGLU

Human iPSC

HGSNAT

Human iPSC

Main findings abilities of these mutants to rescue CatB maturation were examined The importance of certain residues in human GNPTAB implicated as donor substrate binding and dimerization was validated A KO cell line was obtained Reduced IDS activity GAG storage Leakage of cathepsin B from the lysosome Activation of the inflammasome pathway A KO animal model was obtained Complete absence of Sgsh enzymatic activity Progressive accumulation of heparan sulfate products with age CNS manifestations, microglial activation, and neuroinflammation Pharmacological inhibition of Caspase-1 reduces neuroinflammation and partially rescues behavioral abnormalities Two homozygous cell lines were obtained Normal stem cell-like morphology and karyotype, expression of pluripotency-specific markers and maintain capability to differentiate into all three germ layers in vitro Reduced NAGLU enzyme activity Two compound heterozygous cell lines were obtained Normal stem cell-like morphology and karyotype, expression of pluripotency-specific markers, and capable to

Reference

Azambuja et al. (2020)

Douek et al. (2021)

Benetó et al. (2020)

Benetó et al. (2019)

(continued)

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Table 8.1 (continued) Disease

Edited gene

Cell line/ animal model

Main findings

Reference

differentiate into all three germ layers in vitro Reduced HGSNAT enzyme activity

being able to transform them into neurites using forskolin and maintaining edited cell features after long-term culture. This cell model was conveniently used in highthroughput screening studies focused on treatment, or to understand the basic molecular mechanisms involved in neuropathic pain that affects FD patients (Kaneski et al. 2022). Another cell model developed for FD was based on patientderived iPSCs engineered using CRISPR-Cas9 tools. The cells were differentiated into cardiomyocytes before submitted to genome editing, and the result allowed the discovery of several novel cellular and secreted protein biomarkers related to heart dysfunction in this disorder (Birket et al. 2019). Gaucher disease (GD) is the second most common LSD, caused by pathogenic variants in GBA1 gene-encoding lysosomal enzyme glucocerebrosidase (GCase) that degrades glucosylceramide and glucosylsphingosine substrates, or the deficiency in saposin C protein, which acts as an activator of GCase. As a result, the deficiencies of GCase or saposin C produce the accumulation of glucocerebroside within lysosomes of macrophages and monocytes. Due to the presence of mutations in the GBA1 gene, patients present engorged macrophages known as Gaucher cells that impair the normal function of several tissues and organs due to their infiltration and inflammation. There is a complex heterogeneity of clinical symptoms even among patients sharing the same GBA1 genotype. Classically, GD is subdivided into three types according to the absence (type I) or appearance and severity of neurological signs (types II and III). The mechanisms involved in neuronal cell death in GD types II and III are not well understood. Also, the presence of mutations in the GBA1 is the most common genetic risk factor for Parkinson’s disease, although the mechanism that links both diseases remains full of uncertainties (Stirnemann et al. 2017; Sidransky et al. 2009). Pavan and colleagues used a human monocytic THP-1 cell line as an input cell to develop macrophage-like cells following a differentiation step. Additionally, a microglia cell model for GD from the glioblastoma U87 cell line was generated through a large deletion in GBA1 using genome editing techniques. The U87 mutant cells demonstrated accumulation of alpha-synuclein, who was retained in the endoplasmic reticulum (ER) and triggered an unfolded protein response. Both cellular models of GD were able to reproduce the main hallmarks of the disease, being suitable for high-throughput screening to discover new therapeutic approaches or even to contribute to the comprehension of basic mechanisms involved in neurodegeneration (Pavan et al. 2020).

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Genome editing is also used to study the indirect effects of silencing a specific lysosomal enzyme that is crucial to endosomal cargo trafficking, β-glucocerebrosidase (GCase), and its contribution to viral infections such as influenza virus A. This virus is an RNA molecule encapsulated by sphingolipids derived from the host membrane. Drews and colleagues edited HEK293 and A549 cell lines through CRISPR-Cas9, resulting in two KO cell models. The absence of GCase led to an impairment of virus migration to the late endosomes and its subsequent fusion, entry, and infection of the host cells. The results demonstrated that GCase KO cells were less infected by the virus compared to wild-type cells and also highlighted the relevance of GCase for endocytic trafficking in general, which includes growth factors and its receptors (Drews et al. 2019). Not only genome-engineered cellular models were developed for GD but also animal models such as zebrafish. To date, three KO animal models were generated through targeting gba1 and gba2 (cytosolic) genes using CRISPR-Cas9 technology, resulting in gba1-/-, gba2-/-, and a double KO gba1-/-; gba2-/-. The main focus was to evaluate the contribution of the cytosolic version of GCase (GBA2) to the glycosphingolipid metabolism in cases where a deficiency of GBA1 is present due to the misunderstanding of its role is damaging, neutral, or beneficial to the body’s cells (Lelieveld et al. 2019). Moreover, achieving these studies in nonmammalian models such as zebrafish may be an alternative to be considered in the face of their comparability to human anatomy and its genetic traits (Zhang and Peterson 2020). It is already known that microRNAs (miRNAs) regulate the expression of several genes including GBA1. One miRNA, miR-155, has been considered a master regulator of inflammation and has been linked to many neurodegenerative diseases, such as Parkinson’s disease and Alzheimer’s disease, leading to its upregulation with markedly neuroinflammation (Watson et al. 2019). A double-mutant gba1-/-; miR155-/- zebrafish was created using genome editing tools to evaluate the effects that ablation of miR-155 would produce in a gba1-deficient model. The results showed that both neuroinflammation and GD progression seem not to be dependent on miR-155 levels, suggesting that miR-155 did not demonstrate a neuroprotective action and would not be a valuable target for developing a new therapy for GD (Watson et al. 2019). In 2020, Scharenberg and colleagues edited human hematopoietic stem cells using a CRISPR/Cas9 approach to generate monocyte/macrophage cells expressing GCase. For this purpose, the GBA1 was targeted to the CCR5 locus, a safe-harbor site (SHS), where genes or other genetic elements can be safely inserted and expressed without disrupting adjacent genes. This methodology demonstrated the capability to modify cells with multi-lineage differentiation characteristics and longterm repopulation, leading to the establishment of lineage-specific protein expression by targeting at a SHS that could be used as a strategy for restoring levels of several lysosomal enzymes to find new opportunities to treat diseases (Scharenberg et al. 2020). Cystinosis is an LSD caused by pathogenic variants in the CTNS gene, which results in the abnormal accumulation of the amino acid cystine in all tissues and organs of the body, but especially in the proximal tubule segment of kidneys. As a

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consequence, a deficiency in the cystinosin transporter that removes cystine from the lysosomes is established (Town et al. 1998). CRISPR-Cas9 technology was used to produce Ctns KO rat model. This new engineered model was able to reproduce the buildup of cystinosin throughout the body, leading to kidney failure and multisystem manifestations similar to those presented by patients, and it will open up an in vivo cystinosis model for studying new therapies (Krohn et al. 2022). Krabbe disease (KD) is an LSD caused by the deficiency of acid hydrolase β-galactosylceramidase (GALC). As a result, galactosylceramide (GalCer) and galactosylsphingosine (psychosine) accumulate and cause different manifestations depending on the cell type that has been affected. This disorder mainly affects children, but it is also present in adult forms. KD lacks a deeper understanding of the damage caused by psychosine accumulation in nervous tissues, especially in the adult form. The CRISPR-Cas9 approach was used as an attempt to create two distinct knock-in mouse models to mimic the severe early infantile and adult-onset forms of KD due to p.(Thr513Met) and p.(Gly41Ser) in the GALC gene, respectively. Both models demonstrated reliability to recapitulate the clinical signs verified in early- and late-onset of KD patients. The advantages of this genome editing technique focused on KD are not limited to the consequences generated by homozygous GALC pathogenic variants, though instead, it highlights the possibility to introduce different already known mutations in cis or even in trans in combination with single-nucleotide polymorphisms located in the same gene (Rebiai et al. 2022). Niemann-Pick type C (NPC) is caused by pathogenic variants in the NPC1 (in 95% of cases) or NPC2 genes, leading to the accumulation of unesterified cholesterol and glycolipids inside lysosomes and late endosomes in the spleen, liver, and brain (Erwood et al. 2019; Wiweger et al. 2021). NPC patients present a fatally progressive neurodegeneration, and frequently they are compound heterozygotes for a private mutation, which complicates the characterization of these variants. In 2019, the first human near-haploid cell line (HAP1) model for NPC was created using CRISPR-Cas9 genome editing. This model was useful for studying patients’ private mutations since it allowed to understand its related biochemical behavior and molecular phenotypes seen in patients’ fibroblasts. The study proposed that HAP1 engineered cell models could be a suitable strategy to improve genotypephenotype correlation studies not exclusively in NPC but also in several other genetic disorders (Erwood et al. 2019). Despite improvements in our understanding of the molecular processes behind NPC and the existence of several NPC models, there are still many unanswered questions regarding this condition. The majority of studies are focused on NPC1 (e.g., NPC1 KO in HeLa cells edited by CRISPR/Cas9) (Du et al. 2017); a zebrafish model was developed using CRISPR/Cas9 technology to generate a NPC2 KO, once human NPC1 and NPC2 have homologs in these vertebrates. This NPC2 model may be useful for high-throughput screening for drug development to treat the early stages of NPC (Wiweger et al. 2021). Metachromatic leukodystrophy (MLD) is characterized by the deficiency of arylsulfatase A due to pathogenic variants in the ARSA coding gene, which affects the breakdown of cerebroside 3-sulfate, a sulfatide mainly found in the central and peripheral nervous systems. MLD patients usually present severe motor and

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cognitive deficits caused by progressive demyelination. To date, there is no effective treatment for MLD that slows down the advancement of symptoms and avoids the patient’s death. Nevertheless, it has been proposed the usage of drugs with known mechanisms of action to alleviate clinical signs, such as metformin that has been applied to delay aging in patients with neurodegenerative disease and cancer. Human Schwann cells (HSCs) isolated from the human spinal nerve were used for the generation of a MLD cell model using CRISPR-Cas9 genome editing through a mutation in exon 2 of the ARSA that caused a frameshift alteration. This model was used to evaluate the contribution of metformin in the amelioration and decrease of impairment caused by sulfatide accumulation at the mitochondrial level. The study’s results demonstrated an improvement in mitochondrial metabolism and a reduction of reactive oxygen species mitochondrial production (Sanchez-Álvarez et al. 2022). Tay-Sachs (TSD) and Sandhoff (SD) are diseases classified as GM2-gangliosidoses due to deficiency of β-hexosaminidases (Hex) and subsequent accumulation of its substrate (GM2 ganglioside) or deficiency in the GM2 activator protein (Cachon-Gonzalez et al. 2018). Hex are a group of isozymes constituted by dimers of α and β subunits, encoded by HEXA and HEXB genes, respectively. Pathogenic variants in the HEXA lead to TSD, while pathogenic variants in the HEXB result in SD. Both diseases are characterized by neurological impairment owing to GM2 ganglioside buildup in neurons and its cytotoxic effects. To date, there is no therapeutic approach for TSD and SD due to the incapacity of drug molecules of crossing the blood-brain barrier (Leal et al. 2020a). Previous studies demonstrated that a modified Hex μ subunit (HEXM) is able to degrade GM2 gangliosides as a result of the formation of homodimers of both α and β subunits. For this reason, it is being considered an alternative for treating both TSD and SD. Ou and colleagues used genome editing CRISPR-Cas9 tools to the HEXM subunit in a gene editing system to treat neonatal Sandhoff mice as an example of in vivo gene editing focused on GM2-gangliosidoses (Ou et al. 2020a).

3.1.2

Oligosaccharidoses

Pathogenic variants in the GLB1 gene cause defect in β-galactosidase enzyme and result in GM1 gangliosidosis. This disorder is characterized by a progressive accumulation of GM1 ganglioside, which mainly affects the CNS and viscera, associated also with important skeletal abnormalities. In 2021, the CRISPR-Cas9 approach was applied to create a Gbl1 knock-in mouse model carrying the mutation p. (Gly455Arg). This animal model showed progressive neuronal damage, decline in performance when submitted to behavioral tests, an activation of microglia, and autophagy, which seems to be a reliable model to mimic chronic effects that result from the accumulation of GM1 ganglioside and may be used during the development of new therapies (Liu et al. 2021). Another example of LSD that used a murine knock-in model developed by application of CRISPR-Cas9 techniques is Pompe disease (PD). This model had the p.(Tyr609*) mutation in the Gaa gene through the use of a dual-single guide

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RNA approach flanking the intended Gaa insertion site. The engineered animals presented early onset of severe hypertrophic cardiomyopathy as expected, in addition to the skeletal muscle weakness seen in PD patients (Huang et al. 2020). PD is caused by the deficiency of acid alpha-glucosidase (GAA) that converges to the accumulation of glycogen and its consequences to the organs adequately functioning. A distinct study focused on the generation of KO isogenic cell lines derived from mice carrying severe mutations in the Gaa using genome editing. These new cell lines were capable of reproducing PD features such as the buildup of glycogen and autophagic debris, which contribute to the disease progression and muscle weakness (Aguilar-González et al. 2022).

3.1.3

Mucolipidoses

The N-acetylglucosamine-1-phosphotransferase (GlcNAc-1-phosphotransferase) is an enzyme complex formed by six different units (α2/β2/γ2), which is responsible for the phosphorylation of N-linked glycans to produce M6P tags in proteins that should be transferred from the Golgi to the lysosomes. Both catalytic subunits (α and β) are encoded by the GNPTAB gene, while the γ one is produced by the GNPTG gene. Patients with GlcNAc-1-phosphotransferase defects are known to have mucolipidoses (Velho et al. 2019). In 2022, Du and colleagues published the first study that generated a GNPTAB KO HeLa cell line using genome editing tools to confirm the relevance of human GNPTAB residues implicated as donor substrate binding and dimerization. Also, they evaluated the consequences of specific missense mutations. Their hypothesis was based on the cryo-electron microscopy structure of the Drosophila melanogaster GNPTAB homolog in which they were able to pinpoint critical residues for the human GNPTAB (Du et al. 2022).

3.1.4

Mucopolysaccharidoses

MPS type II (also known as Hunter syndrome) is an X-linked LSD caused by the deficiency of iduronate 2-sulfatase enzyme (IDS), necessary for the catalysis of the sulfate group of the iduronic acid present in HS and DS (Peña et al. 2005). Consequently, both GAGs are accumulated inside the lysosomes. Patients with severe form present neurological impairment, but the mechanisms related to neuronal dysfunction are unknown. Azambuja and colleagues employed the CRISPRCas9 genome editing tool to create an IDS KO neuroblastoma (SHSY-5Y) cell line to evaluate the location and role of cathepsin B (CtsB) in disease pathology. They hypothesized that cathepsin is released to the cytoplasm acting as a stimulator to the secretion of cytokines such as interleukin-1-beta (IL 1-β) and caspase-1 (Casp-1). Their findings demonstrated that glycosaminoglycan storage due to the absence of IDS activity leads to alterations in lysosome permeability, and, consequently, CtsB leaks out of lysosomes (Azambuja et al. 2020).

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MPS III or Sanfilippo syndrome comprises a group of four different conditions (A, B, C, and D) characterized by the accumulation of HS. MPS type IIIA (MPS IIIA, Sanfilippo syndrome type A) is the most common form, caused by a loss-of-function mutation in the SGSH gene, which encodes the enzyme N-sulfoglucosamine sulfohydrolase (sulfamidase). MPS IIIA patients tend to be more severely affected. MPS type IIIB (Sanfilippo syndrome type B) is caused by pathogenic variants in the NAGLU gene, which encodes for N-acetyl-alphaglucosaminidase enzyme, while MPS type IIIC (Sanfilippo syndrome type C) is a disease characterized by the deficiency of heparan-α-glucosaminide N-acetyltransferase due to pathogenic variants in the HGSNAT gene (Wagner and Northrup 2019). Benetó and colleagues developed cell line models for MPS IIIB and IIIC from human iPSCs using CRISPR-Cas9 technology through disruption of NAGLU and HGSNAT genes by nonhomologous end joining, respectively (Benetó et al. 2019; Benetó et al. 2020). Since there are no treatments available, these cell models may open up future studies to improve the knowledge of mechanisms behind those diseases and propose new therapeutic approaches. Otherwise, a zebrafish model was generated using CRISPR-Cas9 to produce a KO model of MPS IIIA. This engineered fish was able to recapitulate CNS features seen in patients due to the HS accumulation, in addition to the neuroinflammation and behavioral phenotypes. The availability of the MPS IIIA model may contribute to the understanding of the pathogenesis of the disease, which is still full of gaps; this subtype of MPS III is the most common, and it frequently displays more severe clinical symptoms and advances faster (Douek et al. 2021).

4 Genome Editing as Treatment for Lysosomal Storage Diseases Genome editing holds great potential to treat monogenetic disorders affecting key process in cell metabolism. In the case of LSD, genome editing-based treatments have been evaluated both in vitro and in vivo using ZFNs and the more recently described CRISPR-Cas9 system (Poletto et al. 2020; Leal et al. 2020b). In this section, we will describe the use of these tools for some LSD (Fig. 8.1). Gaucher disease: Sharma et al. used zinc finger nucleases (ZFN) and a donor template, delivered via AAV8 vector, to insert the Gba1 cDNA in the murine albumin locus. In treated wild-type mice, a threefold increase in the plasma GCase activity was observed during the 8 weeks demonstrating the stability of both GBA integration, expression, and secretion (Sharma et al. 2015). In the design of an ex vivo genome editing strategy, Scharenberg et al. (2020) evaluated a CRISPR/ Cas9-based approach to target the GBA cDNA to the CCR5 locus in human hematopoietic stem and progenitor cells (HSPCs). The sgRNA/Cas9 ribonucleoprotein (RNP) complex was delivered through electroporation, and the donor template was transduced via AAV6 vector. When differentiated, these cells were able not only

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Fig. 8.1 In vivo and ex vivo gene editing approaches in lysosomal storage disorders. In vivo gene editing involves the systemic or targeted tissue (e.g., eye, brain, or heart) injection of vectors carrying the components necessary for gene editing. For ex vivo gene editing, cells are collected from a patient (e.g., HSCP, iPSC), edited, and reintroduced into the patient body. TVI tail vein injection, IV intravenous injection, ICV intracerebroventricular injection, ROI retro-orbital injection, HSCPs hematopoietic stem and progenitor cells, iPSCs induced pluripotent stem cell, hNSCs, human neural stem cells

to produce GBA-expressing macrophages but also retain the long-term repopulation and multi-lineage differentiation potential (Scharenberg et al. 2020). These results show the potential of this strategy not only for the design of an ex vivo genome editing therapy for Gaucher disease but also for other LSDs. Fabry disease: By using CRISPR/Cas9, it was possible to correct Fabry disease patient iPSCs carrying the mutation IVS4+919G>A, which has been associated with the cardiac phenotype on these patients (Song et al. 2021). Decrease on multilayered lysosome bodies, intracellular Gb3 clearance, and recovery of autophagy flux was observed on the edited Fabry disease iPSCs (Song et al. 2021). GM2 gangliosidoses: A recent study using CRISPR/Cas9 and the albumin locus as a safe harbor evaluated the therapeutic effect of a chimeric protein termed HexM, which contains the α-subunit active site and the stable β-subunit interface to degrade the GM2 ganglioside (Ou et al. 2020a). Using a hydrodynamic injection to deliver this cDNA into a Sandhoff disease animal model, the treated group showed a ~ sevenfold higher enzymatic activity than the untreated control group in several tissues such as the liver, heart, and spleen. Nonetheless, neither a reduction in GM2 ganglioside levels was observed in the brain nor an improvement in the pole and fear test, which indicates this genome editing tool requires further improvement (Ou et al. 2020a). Importantly, this study adds evidence that supports the use of albumin locus as a safe harbor for gene integration to correct the enzyme deficiency in several tissues. Cerebral organoids, which model the first trimester of neurodevelopment, were generated using human Sandhoff disease iPSCs (Allende et al. 2018). CRISPR/Cas9 genome editing iPSCs showed β-hexosaminidase activity ~50% of wild-type iPSC levels. Cerebral organoids generated with CRISPR/Cas9 genome editing iPSCs showed reduced GM2 and GD3 ganglioside accumulation and increased size and

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cellular proliferation compared to those organoids generated with untreated iPSCs. Whole-transcriptome analysis demonstrated that neurodevelopment was impaired in the Sandhoff disease organoids, which was restored when genome editing iPSCs were used for organoid generation (Allende et al. 2018). Finally, CRISPR/Cas9 prime editing was used to correct most frequent pathogenic variant in Tay-Sachs disease among the Ashkenazi Jewish population (HEXA 1278+TATC) by using Tay-Sachs disease-induced 293T cell lines (Anzalone et al. 2019). From the 43 pegRNAs used in that study, 19 resulted in >20% successful editing, and the best pegRNA resulted in 33% efficiency with 0.32% Indels (Anzalone et al. 2019). Krabbe disease: As a proof-of-concept, Dever et al. (2019) genome edited human neural stem cells (hNSCs) and generated hNSC overexpressing GALC. Edited cells were able to differentiate into astrocytes, neurons, and myelin-producing oligodendrocytes when transplanted into oligodendrocyte mutant shiverer-immunodeficient mice. These cells secreted the GALC enzyme, which was capable to cross-correct Krabbe fibroblasts in vitro (Dever et al. 2019). Niemann-Pick type C: Cytosine base editors (CBEs) which can convert CG base pairs to TA, and adenine base editors (ABEs) which convert AT to CG, have been tested as possible gene editing tool for NPC (Levy et al. 2020). After injecting AAV coupled with split-intein CBEs and split-intein ABEs on NPC mice, a 9.2% increase on animals’ life span and ~ 50% correction of the mutated gene copies on cortical neurons were observed (Levy et al. 2020). Mucopolysaccharidosis I: This deficiency leads to the accumulation of glycosaminoglycans (GAGs) mainly in the liver, spleen, bone, connective tissues, and the CNS (Simonaro et al. 2016). ZFN delivered via adeno-associated viral (AAV9) vectors have been tested as a therapeutic approach for MPS I using a murine model of this disease (Ou et al. 2019). By using ZFNs, a functional copy of the human IDUA gene was delivered into the hepatic albumin intron 1, which is commonly known as a safe harbor region for insertion of exogenous DNA (Ou et al. 2019). In this study, 10- to 16-fold increase of IDUA activity was observed in treated mice after 1 and 4 months with subsequent reduction of GAGs on the liver, spleen, lungs, and heart, indicating sustained effect of the therapy after a single intravenous injection (Ou et al. 2019). Nevertheless, heparan sulphate was not reduced in the brain of MPS I mouse. These findings allowed starting a phase 1/2 multicenter clinical trials for MPS I (ClinicalTrials.gov Identifier: NCT02702115). In this trial, three adult patients were enrolled to evaluate the safety and tolerability the ZFN therapy. Noteworthy, no adverse effects related to ZFN therapy were reported. Nevertheless, although IDUA activity increased in leukocytes 22 weeks posttreatment, no improvement was observed neither in the plasma IDUA activity levels nor in the urinary GAGs (Harmatz 2019). Based on these results using ZFN, Ou L et al. (2020b) designed a genome editing strategy based on CRISPR/Cas9 to insert the IDUA cDNA into the albumin locus, using a AAV8 vector for the delivery of the system. The results showed a supraphysiological increase of IDUA activity in the plasma, liver, hearth, spleen, and brain, with a normalization of GAGs on the same tissues and an improvement in the memory and learning ability of treated mice

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(Ou et al. 2020b). Thus, these results confirm the potential of albumin locus to insert therapeutic transgenes and encourage further clinical evaluation of this strategy. Additional studies have shown CRISPR/Cas9 as a promising platform for the development of a gene therapy for MPS I. In 2018, Carvalho et al. used CRISPR/ Cas9 to correct a mutation p.(Trp402*) on human MPS I patient-derived fibroblasts (de Carvalho et al. 2018). The results showed that 30 days posttreatment, there was an increase in the IDUA activity reaching 5% of normal levels and the reduction on the lysosomal mass (de Carvalho et al. 2018). In vivo evaluation was performed in neonatal MPS I mice through the intravenous administration of a liposomal CRISPR/Cas9 complex, to mediate the integration of the IDUA gene into the Rosa26 locus of an expression cassette carrying the murine IDUA cDNA (Schuh et al. 2018). The results showed a significant increase of IDUA activity in serum leading to 6% of wild-type levels, as well as an increase in enzyme activity levels in the heart (12%), lung (9%), liver (3%), kidney (2%), and spleen (0.8%), with no increase observed in the brain. This increase in enzyme activity led to a GAGs reduction in the lung, heart, liver, kidney, and spleen, although the levels were still higher than those observed in wild-type mice (Schuh et al. 2018). In addition, 6 months posttreatment, treated animals showed an improvement in cardiac function. Further characterization of this liposome CRISPR/Cas9 complex showed the reduction of HS and DS levels in the serum, kidney, lung, and spleen, while HS was reduced in the heart and urine and DS in the liver (Schuh et al. 2020). Nevertheless, HS and DS levels in treated animals were higher than those observed in wild-type mice. Noteworthy, there were significant improvements (but not normalization) in facial morphology, body weight (lower weight than untreated), and bone pathology and a normalization of respiratory disease (Schuh et al. 2020). As expected, since no improvements were observed in the brain IDUA activity and GAGs levels, CRISPR/ Cas9 failed to show any effect in the behavioral alterations observed in the MPS I mice, as well in the neuroinflammatory biomarkers (Schuh et al. 2020). Overall, these results showed that even low but constant enzyme activity levels may have a significant impact on several disease biomarkers and alterations, although an improvement in the delivery strategy may be needed to correct the CNS. In this sense, a recent report used frequent nasal administration of the liposome CRISPR/ Cas9 complex in MPS I mice (Vera et al. 2022). After 6 months, IDUA activity levels were 3.6% of wild-type levels, which allowed a significant reduction of HS and DS both in urine and serum, although the levels were higher than those observed in the wild-type animals. Enzyme activity was modestly increased (between 0.5% and 0.7% of wild-type levels) in the heart, lung, and in some brain areas (the olfactory bulb, frontal cortex, and total cortex). A significant reduction in HS and DS levels was observed in the heart, liver, kidney, spleen, and brain cortex. Noteworthy, enzyme activity increases, and GAGs reduction in brain had a positive impact in behavioral alterations observed in MPS I mice (Vera et al. 2022). In this sense, nasal administration represents a potential alternative to treat both the somatic and CNS alterations observed in MPS I. An ex vivo genome editing strategy for MPS I used human CD34+ hematopoietic stem progenitor cells (HSPCs) and a CRISPR/Cas9 system to target IDUA cDNA to

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the CCR5 locus (Gomez-Ospina et al. 2019). In this study, the sgRNA/Cas9 ribonucleoprotein (RNP) complex was delivered through electroporation to the cells, while the donor template was transduced via AAV6 vector. Modified cells secreted supraphysiological IDUA levels, a characteristic that was conserved after differentiation into macrophages. MPS I mice, capable of receiving human cells, treated with modified HSPCs increased the IDUA activity to a 1.3, 50.1, 167.5, and 6.8% of wild-type mice in the plasma, liver, spleen, and brain, respectively (GomezOspina et al. 2019). This increase in enzyme activity led to a 65% reduction of urinary GAGs, as well as normalization of GAGs in the liver and spleen but not in the brain. Treated animals showed an improvement in hepatomegaly and the normalization of bone parameters. At the behavioral level, the authors observed that transplantation of modified bulk cells had a toxic effect associated with a high human cell chimerism, while the use of modified sorted cells (i.e., cells positive for genome editing) did not show any toxicity effect and allowed to see a therapeutic improvement in the behavioral tests performed (Gomez-Ospina et al. 2019). Additional studies have shown the possibility to use CRISPR/Cas9 to correct compound heterozygous mutations in MPS I (Wang et al. 2018). In this strategy, it was proposed that it would be possible to repair the genetic defect by using the correct information present in the heterozygous mutant alleles without the need to deliver an exogenous template. Thus, Cas9 was used to create double-stranded DNA breaks in homologous chromosomes, inducing allelic exchange of the mutant alleles and leading to a wild-type allele and a mutant allele carrying the two mutations initially present in a heterozygous state (Wang et al. 2018). After an AAV9-mediated delivery of Cas9 and sgRNA, allelic exchange was observed in the heart, leading to an increase in IDUA activity (0.5% of wild-type levels) and a reduction of GAGs in this tissue. Although this strategy may have some advantages over other CRISPR/ Cas9 strategies, such as the lack of a template or the possibility to correct several mutations with only one sgRNA, it would be necessary to improve the rate of allelic exchange and design a system that allows a positive selection of the repaired cells (Wang et al. 2018). Bose et al. (Bose et al. 2021) evaluated the use of an in utero base editing strategy to correct the p.(Trp392*) mutation present in a MPS I mouse. Fetal injection of an AAV9 vector carrying an adenine base editor showed editing in the heart, liver, and brain. At 6 months of age, base editing showed a significant increase in IDUA activity in the serum, heart, and liver (~50% of wild-type levels) and a reduction of GAGs in the urine, heart, and liver. Treated mice also showed an improvement in the cardiac and skeletal pathology and an increased survival, but a limited impact in the neurocognitive defects was also observed, which was associated with the low editing of the brain (Bose et al. 2021). Overall, it has been demonstrated the potential of different genome editing strategies in the design of a therapeutic alternative for MPS I, showing that even low activity levels may have a significant impact in the disease phenotype, at least in preclinical studies. Nevertheless, it would be necessary not only to increase the efficacy of these strategies but also the brain delivery of the system or the enzyme.

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Mucopolysaccharidosis II: Following the same strategy described for MPS I, ZFN was used to integrate the IDS cDNA into the albumin locus of MPS II mice via AAV delivery. In treated MPS II mice, the IDS activity increased up to 207-fold compared to untreated mice, while GAGs levels normalized in the liver, spleen, kidney, lungs, heart, and muscle. However, no changes were observed in brain GAGs (Laoharawee et al. 2018). Moreover, high doses of ZNF-donor formulation induced higher enzyme activity in the brain, ~1.5% of wild-type, and improved cognitive capacity. Based on these results, a phase 1/2 multicenter clinical trial for MPS II (ClinicalTrials.gov Identifier: NCT03041324) was performed using a single administration of a ZFN delivered via AAV vectors. The clinical trial was conducted in eight patients to evaluate the safety, tolerability, and effect on leukocyte and plasma IDS activity of ascending doses of this ZFN therapy. The results indicated that the therapy was well-tolerated with no reports of serious adverse effects. Nevertheless, similar to what was observed in the MPS I clinical trial, plasma enzyme activity was only increased in one patient, which returned to baseline levels after the development of mild transaminitis, and reduction of urinary GAGs was not observed after 24 weeks posttreatment (Muenzer 2019). Overall, these results show that genome editing by ZFN is safe, but the therapeutic efficacy needs to be improved. Mucopolysaccharidosis IV A- MPS IVA (or Morquio A syndrome) is a LSD caused by pathogenic variants in the gene encoding for the enzyme N-acetylgalactosamine-6-sulfate sulfatase (GALNS) (Sawamoto et al. 2018). GALNS deficiency leads to the lysosomal accumulation of the KS and chondroitin-6-sulfate (C6S) (Sawamoto et al. 2018; Khan et al. 2017). The first evaluation of the therapeutic potential of CRISPR/Cas9 for MPS IVA was recently reported (Leal and Alméciga-Díaz 2022). In this study, a CRISPR/Cas9 nickase was used to integrate a functional version of the GALNS cDNA into the AAVS1 locus of MPS IVA patient’s fibroblasts via a liposome delivery of the CRISPR/nCas9 and donor plasmids (Leal and Alméciga-Díaz 2022). The results showed an on-target mutagenic efficiency of 37% with no detected changes in the top ten of predicted offtarget sequences. GALNS activity increases up to 40% of wild-type levels 30 days posttreatment, leading to normalization of lysosomal mass, total GAGs, and oxidative stress, which are some of the major findings regarding the pathophysiological events in MPS IVA. Mucopolysaccharidosis VII: MPS VII (or Sly syndrome) is a LSD caused by pathogenic variants in the gene encoding for the enzyme β-glucuronidase (GUSB), leading to the accumulation of HS, CS, and DS. Kao et al. developed a strategy to treat the corneal disease observed in MPS VII by using CRISPR/Cas9 genome editing (Kao et al. 2017). CRISPR/Cas9 and a donor template were delivered, through an AAV vector, either in separate or single vectors. AAV vectors were administered via tail vein or retro-orbital intravenous infusion (ROIV). This report showed that the use of a single vector and ROIV led to better results in eye pathology and survival compared to tail vein administration and the use of separate vectors.

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5 Final Remarks Gene editing technologies have multiple applications for LSDs. Modelling specific cell types and creating animal models with particular genetic variants are already possible and may serve for different purposes: from confirming pathogenicity of genetic alterations to allow screening of potential new drugs. Furthermore, the possibility of targeting the causative variant and correcting the altered sequence allows the development of gene therapy. Controlling the insertion site of the therapeutic sequence is one of the advantages of these technologies, reducing the risk of insertional oncogenesis and limiting the number of copies of the therapeutic gene in the cell’s genome. Improvements and variations in these techniques are already being described to improve safety and efficacy, and in the near future, the first gene editing products for LSD should be reaching advanced clinical trials phases. Acknowledgments DAS received doctoral a scholarship from Pontificia Universidad Javeriana. CJAD is supported by Ministerio de Ciencia, Tecnología e Innovación, Colombia (Contract 120380763212, ID 8352); Pontificia Universidad Javeriana (ID 20289); the National MPS Society (ID 9509); and the Institute for the Study of Inborn Errors of Metabolism (Activity 120289301011ZZ). This work was supported by FIPE-HCPA, CNPq, CAPES, FAPESP (2019/ 15369-6), and FAPERGS (22/2551-0000385-0).

References Aguilar-González A, González-Correa JE, Barriocanal-Casado E et al (2022) Isogenic GAA-KO murine muscle cell lines mimicking severe Pompe mutations as preclinical models for the screening of potential gene therapy strategies. Int J Mol Sci 23:6298. https://doi.org/10.3390/ ijms23116298 Allende ML, Cook EK, Larman BC, Nugent A, Brady JM, Golebiowski D, Sena-Esteves M, Tifft CJ, Proia RL (2018) Cerebral organoids derived from Sandhoff disease-induced pluripotent stem cells exhibit impaired neurodifferentiation. J Lipid Res 59:550. https://doi.org/10.1194/jlr. M081323 Alroy J, Lyons JA (2014) Lysosomal storage diseases. J Inborn Errors Metab Screen 2: 2326409813517663. https://doi.org/10.1177/2326409813517663 Anzalone AV, Randolph PB, Davis JR et al (2019) Search-and-replace genome editing without double-strand breaks or donor DNA. Nature 576:149. https://doi.org/10.1038/s41586-0191711-4 Anzalone AV, Koblan LW, Liu DR (2020) Genome editing with CRISPR-Cas nucleases, base editors, transposases and prime editors. Nat Biotechnol 38:824–844 Azambuja AS, Pimentel-Vera LN, Gonzalez EA, Poletto E, Pinheiro CV, Matte U, Giugliani R, Baldo G (2020) Evidence for inflammasome activation in the brain of mucopolysaccharidosis type II mice. Metab Brain Dis 35:1231–1236 Baehner F, Schmiedeskamp C, Krummenauer F, Miebach E, Bajbouj M, Whybra C, Kohlschütter A, Kampmann C, Beck M (2005) Cumulative incidence rates of the mucopolysaccharidoses in Germany. J Inherit Metab Dis 28:1011–1017

8

Genome Editing Tools for Lysosomal Storage Disorders

151

Baldo G, Tavares AMV, Gonzalez E, Poletto E, Mayer FQ, Matte U d S, Giugliani R (2017) Progressive heart disease in mucopolysaccharidosis type I mice may be mediated by increased cathepsin B activity. Cardiovasc Pathol 27:45–50 Benetó N, Cozar M, García-Morant M, Creus-Bachiller E, Vilageliu L, Grinberg D, Canals I (2019) Generation of two compound heterozygous HGSNAT-mutated lines from healthy induced pluripotent stem cells using CRISPR/Cas9 to model Sanfilippo C syndrome. Stem Cell Res 41:101616 Benetó N, Cozar M, Gort L, Pacheco L, Vilageliu L, Grinberg D, Canals I (2020) Generation of two NAGLU-mutated homozygous cell lines from healthy induced pluripotent stem cells using CRISPR/Cas9 to model Sanfilippo B syndrome. Stem Cell Res 42:101668 Bétermier M, Bertrand P, Lopez BS (2014) Is non-homologous end-joining really an inherently error-prone process? PLoS Genet 10:e1004086 Birket MJ, Raibaud S, Lettieri M et al (2019) A human stem cell model of Fabry disease implicates LIMP-2 accumulation in cardiomyocyte pathology. Stem cell reports 13:380–393 Bose SK, White BM, Kashyap MV et al (2021) In utero adenine base editing corrects multi-organ pathology in a lethal lysosomal storage disease. Nat Commun 12:4291 Boya P, Kroemer G (2008) Lysosomal membrane permeabilization in cell death. Oncogene 27: 6434–6451 Braulke T, Bonifacino JS (2009) Sorting of lysosomal proteins. Biochim Biophys Acta, Mol Cell Res 1793:605–614 Cachon-Gonzalez MB, Zaccariotto E, Cox TM (2018) Genetics and therapies for GM2 Gangliosidosis. Curr Gene Ther 18:68–89 Costa-Motta FM, Bender F, Acosta A et al (2014) A community-based study of Mucopolysaccharidosis type VI in Brazil: the influence of founder effect, endogamy and consanguinity. Hum Hered 77:189–196 de Carvalho TG, Schuh R, Pasqualim G, Pellenz FM, Filippi-Chiela EC, Giugliani R, Baldo G, Matte U (2018) CRISPR-Cas9-mediated gene editing in human MPS I fibroblasts. Gene 678: 33–37 Dever DP, Scharenberg SG, Camarena J et al (2019) CRISPR/Cas9 genome engineering in engraftable human brain-derived neural stem cells. Iscience 15:524–535. https://doi.org/10. 1016/j.isci.2019.04.036 Douek AM, Amiri Khabooshan M, Henry J, Stamatis S-A, Kreuder F, Ramm G, Änkö M-L, Wlodkowic D, Kaslin J (2021) An engineered sgsh mutant zebrafish recapitulates molecular and Behavioural pathobiology of Sanfilippo syndrome a/MPS IIIA. Int J Mol Sci 22:5948 Drews K, Calgi MP, Harrison WC et al (2019) Glucosylceramidase maintains influenza virus infection by regulating endocytosis. J Virol 93:e00017–e00019. https://doi.org/10.1128/jvi. 00017-19 Du X, Lukmantara I, Yang H (2017) CRISPR/Cas9-mediated generation of Niemann-pick C1 knockout cell line. Methods Mol Biol 1583:73–83 Du S, Wang G, Zhang Z, Ma C, Gao N, Xiao J (2022) Structural insights into how GlcNAc-1phosphotransferase directs lysosomal protein transport. J Biol Chem 298:101702 Erwood S, Brewer RA, Bily TMI, Maino E, Zhou L, Cohn RD, Ivakine EA (2019) Modeling Niemann-pick disease type C in a human haploid cell line allows for patient variant characterization and clinical interpretation. Genome Res 29:2010. https://doi.org/10.1101/gr.250720.119 Federhen A, Pasqualim G, de Freitas TF, Gonzalez EA, Trapp F, Matte U, Giugliani R (2020) Estimated birth prevalence of mucopolysaccharidoses in Brazil. Am J Med Genet Part A 182: 469–483 Feltri ML, Weinstock NI, Favret J, Dhimal N, Wrabetz L, Shin D (2021) Mechanisms of demyelination and neurodegeneration in globoid cell leukodystrophy. Glia 69:2309–2331 Fernández-Pereira C, Millán-Tejado BS, Gallardo-Gómez M, Pérez-Márquez T, Alves-Villar M, Melcón-Crespo C, Fernández-Martín J, Ortolano S (2021) Therapeutic approaches in lysosomal storage diseases. Biomol Ther 11:1–19

152

E. A. Gonzalez et al.

Fuller M, Meikle PJ, Hopwood JJ (2006) Epidemiology of lysosomal storage diseases: an overview. In: Fabry Disease: Perspectives from 5 Years of FOS. Oxford PharmaGenesis, Oxford Futerman AH, Van Meer G (2004) The cell biology of lysosomal storage disorders. Nat Rev Mol Cell Biol 5:554–565 Gaj T, Gersbach CA, Barbas CF (2013) ZFN, TALEN, and CRISPR/Cas-based methods for genome engineering. Trends Biotechnol 31:397–405 Giugliani R, Vairo F, Beck M, Wraith E, Cowan T, Grabowski G (2017) Lysosomal disorders. In: Pediatric endocrinology and inborn errors of metabolism, 2nd edn. McGraw-Hill Education, New York, pp 983–1021 Giugliani R, Barth AL, Dumas MRC et al (2021) Mucopolysaccharidosis VII in Brazil: natural history and clinical findings. Orphanet J Rare Dis 16:1–9 Gomez-Ospina N, Scharenberg SG, Mostrel N et al (2019) Human genome-edited hematopoietic stem cells phenotypically correct Mucopolysaccharidosis type I. Nat Commun 10:1–14 Gonzalez EA, Martins GR, Tavares AMV, Viegas M, Poletto E, Giugliani R, Matte U, Baldo G (2018) Cathepsin B inhibition attenuates cardiovascular pathology in mucopolysaccharidosis I mice. Life Sci 196:102–109 Harmatz P (2019) EMPOWERS: A phase 1/2 clinical trial of SB-318 ZFN-mediated in vivo human genome editing for treatment of MPSI (Hurler Syndrome). Mol Genet Metab 126(2):S68 Huang JY, Kan S-H, Sandfeld EK, Dalton ND, Rangel AD, Chan Y, Davis-Turak J, Neumann J, Wang RY (2020) CRISPR-Cas9 generated Pompe knock-in murine model exhibits early-onset hypertrophic cardiomyopathy and skeletal muscle weakness. Sci Rep 10:10321 Kaneski CR, Hanover JA, Schueler Hoffman UH (2022) Generation of an in vitro model for peripheral neuropathy in Fabry disease using CRISPR-Cas9 in the nociceptive dorsal root ganglion cell line 50B11. Mol Genet Metab reports 31:100871 Kao W, Ferreira T, Yuan Y, Dong F, Hu Y-C, Call M, Coulson-Thomas VJ, Zhang J, Rice T (2017) Gene therapy of Mucopolysaccharidosis type VII (MPS VII) with CRISPR/Cas9 genome editing. Invest Ophthalmol Vis Sci 58:3374 Khan S, Alméciga-Díaz CJ, Sawamoto K, Mackenzie WG, Theroux MC, Pizarro C, Mason RW, Orii T, Tomatsu S (2017) Mucopolysaccharidosis IVA and glycosaminoglycans. Mol Genet Metab 120:78–95 Krohn P, Rega LR, Harvent M et al (2022) Multisystem involvement, defective lysosomes and impaired autophagy in a novel rat model of nephropathic cystinosis. Hum Mol Genet 31:2262– 2278 Kuchař L, Ledvinová J, Hřebíček M et al (2009) Prosaposin deficiency and saposin B deficiency (activator-deficient metachromatic leukodystrophy): report on two patients detected by analysis of urinary sphingolipids and carrying novel PSAP gene mutations. Am J Med Genet Part A 149A:613–621 Laoharawee K, DeKelver RC, Podetz-Pedersen KM et al (2018) Dose-dependent prevention of metabolic and neurologic disease in murine MPS II by ZFN-mediated in vivo genome editing. Mol Ther 26:1127. https://doi.org/10.1016/j.ymthe.2018.03.002 Leal AF, Alméciga-Díaz CJ (2022) Efficient CRISPR/Cas9 nickase-mediated genome editing in an in vitro model of mucopolysaccharidosis IVA. Gene Ther 30:107. https://doi.org/10.1038/ s41434-022-00344-3 Leal AF, Benincore-Flórez E, Solano-Galarza D, Garzón Jaramillo RG, Echeverri-Peña OY, Suarez DA, Alméciga-Díaz CJ, Espejo-Mojica AJ (2020a) GM2 Gangliosidoses: clinical features, pathophysiological aspects, and current therapies. Int J Mol Sci 21:6213 Leal AF, Espejo-Mojica AJ, Sánchez OF, Ramírez CM, Reyes LH, Cruz JC, Alméciga-Díaz CJ (2020b) Lysosomal storage diseases: current therapies and future alternatives. J Mol Med 98: 931–946 Lelieveld LT, Mirzaian M, Kuo C-L et al (2019) Role of β-glucosidase 2 in aberrant glycosphingolipid metabolism: model of glucocerebrosidase deficiency in zebrafish. J Lipid Res 60:1851–1867

8

Genome Editing Tools for Lysosomal Storage Disorders

153

Levy JM, Yeh WH, Pendse N, Davis JR, Hennessey E, Butcher R, Koblan LW, Comander J, Liu Q, Liu DR (2020) Cytosine and adenine base editing of the brain, liver, retina, heart and skeletal muscle of mice via adeno-associated viruses. Nat Biomed Eng 4:97. https://doi.org/10.1038/ s41551-019-0501-5 Liu S, Feng Y, Huang Y, Jiang X, Tang C, Tang F, Zeng C, Liu L (2021) A GM1 gangliosidosis mutant mouse model exhibits activated microglia and disturbed autophagy. Exp Biol Med (Maywood) 246:1330–1341 Lloyd-Evans E, Morgan AJ, He X, Smith DA, Elliot-Smith E, Sillence DJ, Churchill GC, Schuchman EH, Galione A, Platt FM (2008) Niemann-pick disease type C1 is a sphingosine storage disease that causes deregulation of lysosomal calcium. Nat Med 14:1247–1255 Muenzer J (2019) CHAMPIONS: A phase 1/2 clinical trial with dose escalation of SB-913 ZFN-mediated in vivo human genome editing for treatment of MPS II (Hunter Syndrome). Mol Genet Metab 126:S104 Ou L, DeKelver RC, Rohde M et al (2019) ZFN-mediated in vivo genome editing corrects murine hurler syndrome. Mol Ther 27:178. https://doi.org/10.1016/j.ymthe.2018.10.018 Ou L, Przybilla MJ, Tăbăran A-F, Overn P, O’Sullivan MG, Jiang X, Sidhu R, Kell PJ, Ory DS, Whitley CB (2020a) A novel gene editing system to treat both Tay–Sachs and Sandhoff diseases. Gene Ther 27(5):226–236 Ou L, Przybilla MJ, Ahlat O, Kim S, Overn P, Jarnes J, O’Sullivan MG, Whitley CB (2020b) A highly efficacious PS gene editing system corrects metabolic and neurological complications of Mucopolysaccharidosis type I. Mol Ther 28:1442–1454 Parenti G, Pignata C, Vajro P, Salerno M (2013) New strategies for the treatment of lysosomal storage diseases (review). Int J Mol Med 31:11–20 Parenti G, Andria G, Ballabio A (2015) Lysosomal storage diseases: from pathophysiology to therapy. Annu Rev Med 66:471–486 Pavan E, Ormazabal M, Peruzzo P, Vaena E, Rozenfeld P, Dardis A (2020) CRISPR/Cas9 editing for Gaucher disease modelling. Int J Mol Sci 21:3268. https://doi.org/10.3390/ijms21093268 Peña O, Sosa Á, Echeverri O, Saenz H, Barrera LA (2005) Producción de anticuerpos policlonales IgG contra la proteína iduronato-2-sulfato sulfatasa y desarrollo de un sistema de detección para IDS humana recombinante. Biomedica 25:181–188 Pereira VG, Gazarini ML, Rodrigues LC, Da Silva FH, Han SW, Martins AM, Tersariol ILS, D’Almeida V (2010) Evidence of lysosomal membrane permeabilization in mucopolysaccharidosis type I: rupture of calcium and proton homeostasis. J Cell Physiol 223: 335–342 Pimentel-Vera LN, Poletto E, Gonzalez EA, de Oliveira PF, Giugliani R, Baldo G (2021) Genome editing in lysosomal disorders. Prog Mol Biol Transl Sci 182:289–325 Platt FM, Boland B, van der Spoel AC (2012) Lysosomal storage disorders: the cellular impact of lysosomal dysfunction. J Cell Biol 199:723–734 Platt FM, D’Azzo A, Davidson BL, Neufeld EF, Tifft CJ (2018) Lysosomal storage diseases. Nat Rev Dis Prim 4:27 Poletto E, Baldo G, Gomez-Ospina N (2020) Genome editing for Mucopolysaccharidoses. Int J Mol Sci 21:500 Politei J, Cabello J, Jacobo V, Guillermo V, Antonio L, Ballesteros A, Ana Maria M (2019) Enfermedad de Fabry: Nuevos conceptos en su historia natural, evolución y tratamiento, en relación a los hallazgos del Registro Fabry. Revista de nefrología, diálisis y transplante 29(4): 145–152 Poswar F, Vairo F, Burin M, Michelin-Tirelli K, Brusius-Facchin A, Kubaski F, Desouza C, Baldo G, Giugliani R (2019) Lysosomal diseases: overview on current diagnosis and treatment. Genet Mol Biol 42:165–177 Rebiai R, Rue E, Zaldua S et al (2022) CRISPR-Cas9 Knock-in of T513M and G41S mutations in the murine β-Galactosyl-ceramidase gene re-capitulates early-onset and adult-onset forms of Krabbe disease. Front Mol Neurosci 15:896314

154

E. A. Gonzalez et al.

Sanchez-Álvarez NT, Bautista-Niño PK, Trejos-Suárez J, Serrano-Díaz NC (2022) A model of metformin mitochondrial metabolism in metachromatic leukodystrophy: first description of human Schwann cells transfected with CRISPR-Cas9. Open Biol 12:210371. https://doi.org/ 10.1098/rsob.210371 Sawamoto K, Alméciga-Díaz CJ, Mason RW, Orii T, Tomatsu S (2018) Mucopolysaccharidosis type IVA: clinical features, biochemistry, diagnosis, genetics, and treatment. In: Tomatsu S, Lavery C, Giugliani R, Harmatz P, Scarpa M, Węgrzyn G, Orii T (eds) Mucopolysaccharidoses Updat. (2 Vol. set). Nova Science Publishers, Inc., Hauppauge, pp 235–272 Scharenberg SG, Poletto E, Lucot KL, Colella P, Sheikali A, Montine TJ, Porteus MH, GomezOspina N (2020) Engineering monocyte/macrophage-specific glucocerebrosidase expression in human hematopoietic stem cells using genome editing. Nat Commun 11:3327. https://doi. org/10.1038/s41467-020-17148-x Schiffmann R (2006) Neuropathy and Fabry disease: pathogenesis and enzyme replacement therapy. Acta Neurol Belg 106:61–65 Schuh RS, Poletto É, Pasqualim G, Tavares AMV, Meyer FS, Gonzalez EA, Giugliani R, Matte U, Teixeira HF, Baldo G (2018) In vivo genome editing of mucopolysaccharidosis I mice using the CRISPR/Cas9 system. J Control Release 288:23–33 Schuh RS, Gonzalez EA, Tavares AMV et al (2020) Neonatal nonviral gene editing with the CRISPR/Cas9 system improves some cardiovascular, respiratory, and bone disease features of the mucopolysaccharidosis I phenotype in mice. Gene Ther 27:74–84 Schuller Y, Linthorst GE, Hollak CEM, Van Schaik IN, Biegstraaten M (2016) Pain management strategies for neuropathic pain in Fabry disease--a systematic review. BMC Neurol 16:25 Sévin M, Lesca G, Baumann N, Millat G, Lyon-Caen O, Vanier MT, Sedel F (2007) The adult form of Niemann-pick disease type C. Brain 130:120–133 Shaimardanova AA, Chulpanova DS, Solovyeva VV, Mullagulova AI, Kitaeva KV, Allegrucci C, Rizvanov AA (2020) Metachromatic Leukodystrophy: diagnosis, modeling, and treatment approaches. Front Med 7:1–17 Sharma R, Anguela XM, Doyon Y et al (2015) In vivo genome editing of the albumin locus as a platform for protein replacement therapy. Blood 126:1777–1784 Sidransky E, Nalls MA, Aasly JO et al (2009) Multicenter analysis of glucocerebrosidase mutations in Parkinson’s disease. N Engl J Med 361:1651–1661 Simonaro CM, Tomatsu S, Sikora T et al (2016) Pentosan Polysulfate: Oral versus subcutaneous injection in Mucopolysaccharidosis type I dogs. PLoS One 11:e0153136 Solomon M, Muro S (2017) Lysosomal enzyme replacement therapies: historical development, clinical outcomes, and future perspectives. Adv Drug Deliv Rev 118:109–134 Song H-Y, Chiang H-C, Tseng W-L et al (2016) Using CRISPR/Cas9-mediated GLA gene knockout as an in vitro drug screening model for Fabry disease. Int J Mol Sci 17:2089. https://doi.org/10.3390/ijms17122089 Song H-Y, Yang Y-P, Chien Y et al (2021) Reversal of the inflammatory responses in Fabry patient iPSC-derived cardiovascular endothelial cells by CRISPR/Cas9-corrected mutation. Int J Mol Sci 22:2381 Stirnemann J, Belmatoug N, Camou F et al (2017) A review of Gaucher disease pathophysiology, Clinical Presentation and Treatments. Int J Mol Sci 18:441. https://doi.org/10.3390/ ijms18020441 Sun A (2018) Lysosomal storage disease overview. Ann Transl Med 6:476–476 Town M, Jean G, Cherqui S et al (1998) A novel gene encoding an integral membrane protein is mutated in nephropathic cystinosis. Nat Genet 18:319–324 Velho RV, Harms FL, Danyukova T et al (2019) The lysosomal storage disorders mucolipidosis type II, type III alpha/beta, and type III gamma: update on GNPTAB and GNPTG mutations. Hum Mutat humu 40(7):842–864 Vellodi A (2005) Lysosomal storage disorders. Br J Haematol 128:413–431

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Vera LNP, Schuh RS, Fachel FNS et al (2022) Brain and visceral gene editing of mucopolysaccharidosis I mice by nasal delivery of the CRISPR/Cas9 system. J Gene Med 24: e3410. https://doi.org/10.1002/jgm.3410 Villalobos J, Politei JM, Martins AM, Cabrera G, Amartino H, Lemay R, Ospina S, Ordoñez SS, Varas C (2013) Fabry disease in latin america: data from the fabry registry. JIMD Rep 8:91–99 Wagner V, Northrup H (2019) Mucopolysaccharidosis Type III. In: Adam M, Ardinger H, Pagon R et al (eds) GeneReviews® [Internet]. University of Washington, Seattle, Seattle (WA), pp 1993–2021 Wang D, Li J, Song CQ et al (2018) Cas9-mediated allelic exchange repairs compound heterozygous recessive mutations in mice. Nat Biotechnol 36:839. https://doi.org/10.1038/nbt.4219 Watson L, Keatinge M, Gegg M, Bai Q, Sandulescu MC, Vardi A, Futerman AH, Schapira AHV, Burton EA, Bandmann O (2019) Ablation of the pro-inflammatory master regulator miR-155 does not mitigate neuroinflammation or neurodegeneration in a vertebrate model of Gaucher’s disease. Neurobiol Dis 127:563–569 Wiweger M, Majewski L, Adamek-Urbanska D, Wasilewska I, Kuznicki J (2021) npc2-deficient zebrafish reproduce neurological and inflammatory symptoms of Niemann-pick type C disease. Front Cell Neurosci 15:647860. https://doi.org/10.3389/fncel.2021.647860 Wraith JE (2002) Lysosomal disorders. Semin Neonatol 7:75–83 Xu H, Ren D (2015) Lysosomal physiology. Annu Rev Physiol 77:57–80 Zhang T, Peterson RT (2020) Modeling lysosomal storage diseases in the zebrafish. Front Mol Biosci 7:82. https://doi.org/10.3389/fmolb.2020.00082

Chapter 9

CRISPR Libraries and Whole-Genome Screening to Identify Essential Factors for Viral Infections Isadora Marques Paiva, Samara Damasceno, and Thiago Mattar Cunha

1 Introduction About CRISPR Libraries The CRISPR (clustered regularly interspaced short palindromic repeats) system has revolutionized genetics and offers a simple and inexpensive way for the edition of the genome. Researchers developed the CRISPR-Cas9 technique based on the natural defense mechanism of prokaryote against reinvasion by bacteriophages, transposons, and plasmids (Sampson and Weiss 2014). This defense mechanism can be described in three stages: first is the primary contact, in which the prokaryote can cleave the pathogen’s DNA and integrate small fragments called spacers into its CRISPR locus, thus acquiring a memory against new invasions. Second is the uninterrupted transcription of the entire CRISPR locus followed by processing of small guide RNAs (sgRNA), each corresponding to a separate spacer. And finally, after the synthesis of these sgRNAs and the Cas protein endonuclease (components of the CRISPR locus), they form a complex that recognizes an exogenous gene sequence and destroys it (Makarova et al. 2011). The CRISPR technique has as its central element the endonuclease Cas9 which is directed to selectively achieve its target by a guide RNA, generating perturbation that results in gene repression, activation, or editing. Repression: named CRISPRi,

I. M. Paiva · S. Damasceno Center for Research in Inflammatory Diseases (CRID), Ribeirao Preto Medical School, University of Sao Paulo, Ribeirão Preto, SP, Brazil T. M. Cunha (✉) Center for Research in Inflammatory Diseases (CRID), Ribeirao Preto Medical School, University of Sao Paulo, Ribeirão Preto, SP, Brazil Department of Pharmacology, Ribeirão Preto Medical School, University of São Paulo, Ribeirão Preto, SP, Brazil e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_9

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where the transcription of the target gene is blocked without the need to alter the DNA sequence. In order to do that, nuclease-inactivated Cas9 (dCas9), through sgRNA, is taken to the promoter region of the target gene, preventing the association of DNA with its transcription factors. Activation: named CRISPRa, in this case, the objective is to increase the transcription of a target gene without modifying the DNA sequence. For that, nuclease-inactivated Cas9 (dCas9) fused to transcriptional activation domains is taken by sgRNAs to the promoter region of the target gene and then promotes the association of transcription factors. Editing: modification, addition, or deletion of DNA bases. According to this definition, we have CRISPRko, SNPs (single-nucleotide polymorphisms) integration, and other insertions using donor DNA. Considering CRISPRko, the knockout alleles are generated by the Cas9 endonuclease, which is targeted by a sgRNA to a specific genomic region. Active Cas9 promotes a double-strand break (DSB) at the target site, which is then repaired by nonhomologous end-junction (NHEJ), which usually results in a frameshift mutation and the expression of truncated or nonfunctional proteins (Sander and Joung 2014). The advances brought using the CRISPR-Cas9 technique make possible the development of CRISPR libraries which consist of a set of sgRNAs to cause perturbations in several genes in the same cell population. The use of libraries raised the CRISPR-Cas9 technique to a genomic scale and provides a powerful approach for identifying previously unknown molecular mechanisms and pathways involved in a specific phenotype or biological process (Bock et al. 2022). CRISPR libraries have been developed by several authors, and, currently, various are available for different organisms. The libraries can vary in the number of sgRNAs and in the number of target genes, which can be genome-wide or only a subpool of genes. In addition, custom libraries can be built for a specific class of genes or for validation screenings (for more information, access https://www.addgene.org/pooled-library/) (Hart et al. 2015; Wang et al. 2016; Doench et al. 2016). The CRISPRko libraries are widely used in research with viral agents. These libraries are composed of a set of plasmids containing specific sgRNAs to generate mutations in numerous protein-coding genes. In the plasmids, it always includes the sgRNA associated with a selection marker that can be an antibiotic resistance gene or one that encodes a fluorescent protein. The sgRNAs are usually delivered by lentiviral transduction to achieve genome integration; thus, individual cells receive different sgRNAs and are perturbed according to the sgRNA received (Bock et al. 2022). The recovery of transduced cells is based on the selection marker, and edited cells that carry mutations can be challenged by viral infection or some other selective pressure, such as treatment with some drug, or inducers and repressors of cell proliferation. Then, the sgRNAs are high-throughput sequenced based on the pool of cells retained after the challenge. In the resulting data, depletion of specific sgRNAs identifies genes whose depletion sensitizes cells, whereas their enrichment identifies genes whose disruption confers a selective advantage to that challenge (Bock et al. 2022).

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Based on all the possibilities and advantages of these high-throughput screenings at genomic level using CRISPR-Cas9, we aim with this chapter to point out how this tool helps in the understanding virus-host relationships, such as the mechanisms of virus entry into the host cell, the essential factors for its replication, and the cellular pathways involved in the response against the pathogen.

2 Practical Considerations for Whole-Genome Screening in Viral Research Viruses are obligate intracellular pathogens that depend on the host to complete their life cycle. They enter cells through cell receptors and take advantage of cell machinery and metabolism to replicate their genome, assemble new viruses, and spread. In the same way, the host cell can reassign its functions to react to that infection by detecting pathogen-associated molecular patterns (PAMPs) that will subsequently induce the expression of antiviral genes. The viral groups have particularities regarding the molecular factors that are essential for viral entry, infection, and reproduction. In this context, the genome-scale CRISPR screens have helped to characterize virus-host relationships, making it possible to identify factors that are necessary for the cycle of different viruses and so, new targets for antiviral therapies (Chulanov et al. 2021; Puschnik et al. 2017).

2.1

Cell Type Definition

When studying virus-host interactions by CRISPR screening, we need to follow some practical criteria. First, we need to properly choose the cell line and the type of library to be used according to the objective. The genetic screens can reveal genes that promote or restrict viral replication depending on the type of host cell and the type of screening. Viruses can differ in terms of host and tissue tropism, so to define the study cell, it is necessary to verify the permissiveness of the candidate cells in relation to the virus (Puschnik et al. 2017). This test is performed through virus-cell contact at different viral concentrations called MOI (multiplicity of infection) and different time intervals. The evaluation of cellular permissiveness to the virus can be done by different methods, such as viral titration or quantification of viral load by quantitative PCR of cell cultures after contact with the virus, where a higher titer or viral load is observed in the culture whose cells allowed the virus to complete its life cycle. Another method is by cell viability curve, where a higher viability is observed in the culture whose cells are not permissive. When viruses are able to complete their reproductive cycle (entry, replication of genetic material, assembly, and exit from the cell), more viruses will be available to infect and to kill new cells which results in lower cell viability. With these tests, it is also important to establish the proper MOI and virus-cell contact times to obtain a lethal or sublethal infection depending on the purpose of the study.

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After considering these first points, other challenges can be faced in wholegenome CRISPR screening. Being permissive does not indicate that the cell is the best choice for genomic-scale screening. Some cells have low proliferation capacity, as is the case of primary cells, or are very sensitive and do not resist to the genetic modification protocols. In this context, it is possible to genetically modify cell lines of interest in order to make it more or less permissive. With genomic editing tools, including CRISPR technology, we can increase or reduce the expression of molecular factors essential to the viral cycle. For example, the new coronavirus (SARSCoV-2) has tropism for lung tissue, so it is suggested to study the virus-host relationship using lung cells. Among the two cell lines of pulmonary origin, Calu3 and A549, Calu-3 is the most permissive, presenting greater expression of the ACE2 receptor, necessary for coronavirus infection. However, these cells are less resistant to genetic modification and exhibit a slow expansion. Therefore, it is more interesting to modify the most resistant cell with a higher expansion rate to make it permissive to viral infection, which can be done by inserting new copies of the ACE2 gene.

2.2

Genetic Screen Definition and Library

The genetic screens can reveal genes that promote or restrict viral replication depending on the type of host cell (permissive or nonpermissive) and the type of screening (gain of function or loss of function) according to CRISPR screens: CRISPRa, CRISPRi, and CRISPRko screens (Fig. 9.1a). For example, in a lossof-function screen using CRISPRko technology, knocking out a viral receptor in a permissive cell line will make the cell resistant to infection by the virus. However, in a gain-of-function screen using CRISPRa technology, overexpression of the viral receptor in a nonpermissive cell line will allow infection by the virus (Puschnik et al. 2017). The CRISPRko screens are the most used to study virus-host interaction, where the “human Brunello genome-wide library” and “human GeCKO genomewide library” stand out. Both were developed using a puromycin resistance factor as a selection marker and consider, at least, four guides for each target gene. The greater number of sgRNAs per gene increases the probability of significant enrichment of candidate genes. However, libraries with a large set of sgRNAs require a huge number of cells for the appropriate representation of each guide. This fact limits the use of these libraries in primary cells since they have a low survival and low proliferative capacity, which makes their expansion difficult. Thus, when amplification is unfeasible due to cost or number of cells, libraries that contain fewer sgRNAs per gene can be used for initial screening, followed by secondary screening or careful validation (Fig. 9.1b) (Puschnik et al. 2017). In general, CRISPR libraries can be found in all-in-one system, where each plasmid that composes the pool contains the sgRNA and Cas9 cassette expression, allowing direct perturbation into target cells. On the other hand, the delivery of the guide and the endonuclease can be done separately. In this case, the library consists

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Fig. 9.1 Genome-wide screening with CRISPRko library to study virus infection. (a) In permissive cells where viruses can infect and replicate, the loss-of-function strategy can be used to identify the essential factors for viral success. In contrast, the gain-of-function strategy can be used to study these factors in nonpermissive cells. (b) The construction of commercial or customized libraries is initiated in silico by designing specific guides that are later synthesized and cloned into plasmids. (c) The library’s plasmid pool is amplified by transformation into electrocompetent bacteria. After verifying the coverage and diversity of the library, the plasmids are transfected into HEK cells to produce the lentiviruses that will be transduced into target cells. After transduction, the knockout cells are selected and kept for expansion. (d) Selective pressure occurs by exposing the knockout cells to the virus of interest. After this contact, the surviving cells are expanded and collected for DNA extraction. The genetic material is sequenced and the data analyzed for guide counts

of a pool of plasmids containing only the sgRNAs, while target cells must be previously modified to stable express Cas9. The insertion of the Cas9 endonuclease gene into the cell genome can be carried out through delivery by a lentivirus containing the Cas9 expression vector linked to the selection marker, antibiotic resistance factor, or fluorescent protein (GFP), which allows the isolation of the modified cells.

2.3

Library Amplification

Once the library type has been defined, it is necessary to amplify and verify its coverage to ensure sufficient quantity and maintain the complexity of the sgRNA pool (Fig. 9.1c). Library amplification is performed in a large scale by transforming the plasmid pool into commercial electrocompetent bacteria. These bacteria are grown on solid medium for expansion and collected for plasmid extraction. A

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sample of the extracted pool is subjected to next-generation sequencing to verify the coverage by the guides. In genome-wide CRISPRko, the sequence analysis can be done using MAGeCK-VISPR system, a quality control (QC) analysis, and visualization pipeline for CRISPR screens (Yau and Rana 2018; Joung et al. 2017; Li et al. 2015). In this first sequencing, to verify the amplification quality, we observe the QC measures at sequence and read count level: (1) GC content distribution of the sequencing reads, in which we expect a similar distribution for all samples from the same library; (2) base quality distribution of the sequencing reads, in which one expect a single-peak distribution with median base quality at least 25%; (3) percentage of mapped reads to the total number of sequencing reads, in which expect at least 65%; (4) number of sgRNAs with zero read counts, in which expect at most 1% of total sgRNAs; and (5) total number of reads mapped to the sgRNA Library, which expect preferably over 300 reads per sgRNA. In general, successful amplification is considered when coverage is greater than 200–300 reads per sgRNA in at least 99% of guides. However, considering the diversity of CRISPR libraries and the fact that some contain four or more guides per gene, the minimum number of reads per sgRNA can vary according to the composition of the library since there is no loss of gene representation (Yau and Rana 2018; Joung et al. 2017; Li et al. 2015).

2.4

Library Transduction

With the cell lineage established and the library amplified, the next step is the production of delivery agents for cell transduction (Fig. 9.1c). The vast majority of CRISPR libraries use the lentivirus system for introducing the sgRNAs into cells. For the production of lentivirus, HEK cells are transfected with the envelope and viral packaging plasmids and the library plasmids containing the sgRNAs. Given the transfection time, lentiviral particles are produced and collected for quantification and transduction of target cells (Fig. 9.1c). Efficient transduction will depend on the conditions recommended for each type of library. One of the biggest challenges in the genome-wide CRISPR screening is providing adequate coverage of the library using appropriate amounts of cells and maintaining coverage throughout the experiment. The number of cells to be transduced and the MOI of lentivirus are the criteria that need to be considered for successful transduction. For CRISPRko libraries, the recommended MOI is approximately 0.3 to ensure that most cells receive only one genetic modification. The number of cells must be calculated considering coverage of at least 500 cells expressing each sgRNA, guaranteeing that each modification will be sufficiently represented. For example, for a genome-wide CRISPRko library containing 100,000 sgRNA, it is necessary to transduce at least 1.67 × 108 cells with 30% efficiency to maintain 5 × 107 cells at each passage and harvesting time point for each biological replicate. Regarding the selection of modified cells, it is recommended to do for approximately 7 days, due to the time of maximal knockout efficiency that is achieved on the seventh day after sgRNA transduction (Joung et al. 2017).

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Selective Pressure

To proceed with the study of the virus-host interaction, the knockout cells of the previous step must be subjected to selective pressure through contact with the virus (Fig. 9.1d). Selection conditions must be previously defined, such as MOI and viruscell contact times. In addition, 2–4 infection replicates per screen are recommended to account for stochastic noise. For phenotypic selection, strong selection conditions are preferred in which more than 99% of the cells die from infection, considered a lethal infection. This high stringency increases confidence in identified candidate genes, but genes with subtle effects on viral infection may be lost. There are strategies to reduce stringency and help identify these genes, such as using naturally attenuated virus strains or using antiviral compounds during selection to generate sublethal infections (Puschnik et al. 2017). The phenotypic selection must be delineated to each type of virus in order to accomplish the research objectives. If the virus is not efficient at inducing cell death, longer selection time, more rounds of viral challenge, and larger sgRNA libraries may be needed to help increase the signal-to-noise ratio. The fluorescence-activated cell sorting (FACS)-based selection can also be used to study persistent or non-cytolytic viruses. In addition, it is possible to identify host factors that are necessary for a specific stage of the viral cycle. This can be done through pseudotyped viruses, viral replicons, and internal ribosome entry site reporters that can be used in the infection assay to identify the factors essential only for virus entry, genome replication, and translation, respectively (Puschnik et al. 2017).

2.6

Sequencing and Bioinformatics Analysis

After selective pressure by viral infection, the surviving modified cells are expanded and collected for DNA extraction in parallel with the control knockout cells that are not subjected to selective pressure. With the genetic material, the sgRNA integrations are amplified by PCR and submitted to next-generation sequencing for later quantification of the relative abundance of each guide (Fig. 9.1d) (Puschnik et al. 2017). In this sequencing, in addition to the QC measurements at the sequence level and reading count level mentioned above, we must also observe the QC measurements at the sample and gene level: (1) the Gini index of log-scaled read count distributions, in which one expect at most 0.1 for plasmid or initial state samples, and at most 0.2 for negative selection samples; (2) Pearson correlation coefficient between samples, in which one expect at least 0.8 for replicates; (3) hierarchical clustering of samples or first three PCA components, in which samples with similar conditions should cluster together; and (4) the number of mapped reads (preferably above 300 reads per guide) is an indicator of sample quality and influences the statistical power of the downstream analysis. The low percentage of mapped reads could be due to a sequencing error or sample contamination (Joung et al. 2017; Li et al. 2015).

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The Gini index can be used to measure the evenness of read counts, and a lower index means a more even distribution of read counts. In positive selection experiments, it is normal to have higher Gini index since a few cells surviving could dominate the final pool (a few sgRNA with extreme high counts), while most of the other cells die (more sgRNAs with zero-count). From correlation and PCA analyses, replicates or biological samples with similar conditions are expected to have higher correlations and similar read count distributions. Bioinformatics tools available for different types of screenings help to normalize differences in sequencing depth between populations and determine whether a gene is significantly enriched against background. In general, the level of enrichment of sgRNAs is determined by comparing the number of reads mapped in populations of modified cells that are subjected to selective pressure and the control cells (Li et al. 2015). Through the enrichment of sgRNAs, we can identify a set of candidate genes that must be validated to exclude possible false-positive results. In CRISPRko screening, the greater the number of sgRNAs per gene, the greater the probability of gene enrichment, and the improved sequences for cleavage efficiency make current libraries more reliable than previous libraries, thus minimizing false-negative results. The main limitation of the CRISPRko screen is the potential of arbitrary knockout of genes which makes it difficult to identify all the genes that affect the virus life cycle because some of them can be essential for cell growth and viability and consequently will be excluded from the analyses. About 10% of human genes are essential for cellular growth and viability (Puschnik et al. 2017). In the study of virus-host interaction using CRISPRko screening, the cells surviving to virus-cell contact are those whose knockout gene is possibly important for the success of the virus. For example, mentioning the new coronavirus, if the ACE2 gene is knocked out, the virus will not succeed in the process of entering the cell and consequently; that surviving cell will have the sgRNA for ACE2 enriched. Even so, complementary experiments with other methods are necessary to confirm that the effect was due to the knockout. On the other hand, if a ribosomal gene essential for the synthesis of cellular proteins is knocked out, the cell will die, and this sgRNA will be lost even if the cell is not infected by the virus, which makes it impossible to know the importance of this gene for the viral life cycle.

2.7

Candidate Gene Validation

Finally, with the gene enrichment data of CRISPRko screening, candidate genes are defined for validation. Usually, the first validation of the candidate is done separately by transduction on a smaller scale with sgRNAs unique to the target gene. This test aims to confirm that cell survival was exclusively due to gene knockout. Further evaluations should be performed according to the gene, its protein expression, and function in metabolism. Validations can be at the cellular level (in vitro) or at the organism level (in vivo) and must be carried out extremely rigorously, especially regarding the host organism (human), so that safety is guaranteed (Lee 2019).

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3 Research and Antiviral Therapy Diseases caused by viral infections are of great concern worldwide; the impossibility of curing some and the emergence of mutants resistant to antivirals are factors that lead to persistent infections with low treatment efficiency. In the last decades, several tools that allow the genetic modification of viruses have been used as an antiviral strategy. Techniques such as zinc finger nucleases, transcriptional activator-like effector nucleases (TALENS), and, more recently, CRISPR/Cas-associated nucleases have already been used to manipulate the viral genome (Lee 2019; Bayat et al. 2018). The CRISPR-Cas9 system stands out as an antiviral tool due to its specificity, versatility, and ease of application. However, we must consider some limitations regarding its safety at the time of tests and in vivo applications, such as the possibility of off-target effects and the emergence of mutant and resistant escape strains. Another point is the guarantee that all infected cells will receive the CRISPRCas9 complex, because if there are cells that do not receive the treatment, they can serve as a viral reservoir, limiting clinical application. For this last limitation, the use of AAVs as a CRISPR complex delivery tool is currently being evaluated due to their high viral titer and infectivity (Slaymaker et al. 2016; Kleinstiver et al. 2016; Kaminski et al. 2016; Kotterman and Schaffer 2014). Most antiviral therapies have been unable to eliminate the viral genome from all infected cells. This can occur due to no specificity of targets and mainly due to a latent state of the virus. With large-scale studies using CRISPR libraries, it is possible to identify specific molecular targets for the viruses of interest. This screening strategy allows characterizing the host factors that are essential for the viral cycle, entry, replication, assembly, and even the vectors that transport these viruses to their host. In recent years, large-scale screening studies using CRISPR libraries have been published reporting essential factors for the entry and replication of different viruses that can cause human disease, such as dengue (DENV) (Labeau et al. 2020), Zika virus (ZIKV) (Li et al. 2019), chikungunya (CHIKV) (Meertens et al. 2019), Ebola virus (EBOV) (Flint et al. 2019), human immunodeficiency virus (HIV) (Krasnopolsky et al. 2020; Park et al. 2017), herpes simplex virus (Suzuki et al. 2022), chronic hepatitis (Hyrina et al. 2019), and different strains of influenza virus (Song et al. 2021; Li et al. 2020). More recently, with the COVID-19 pandemic, many studies using this screening strategy were carried out with different cell lines to identify targets for more effective antiviral treatments (Antoine et al. 2021; Baggen et al. 2021; Biering et al. 2021; Sun et al. 2021; Zhu et al. 2021, 2022) (Table 9.1). In general, these studies have sought to identify new targets that have a direct effect on the infectivity of these viruses, their permissiveness and resistance, and their ability to generate cellular disorders when infected. In this context, different libraries, whether for gene loss-of-function screening (CRISPRKo) or for gain-offunction (CRISPRa), can be used. Gecko v2 and Brunello, CRISPRko libraries (more efficient and with less off-target effect) are the most common for human

Chronic hepatitis B Chronic hepatitis B DNA (CHB) virus

Hyrina et al. (2019)

Knockout

Knockout

DNA

Herpes

Suzuki et al. (2022)

Herpes simplex virus type 1 (HSV-1)

Knockout

RNA

Ebola

Flint et al. (2019)

Ebola virus (EBOV)

x

Park et al. (2017)

RNA

Knockout

Knockout

Acquired immuno- Human immunodeficiency deficiency virus syndrome (HIV)

RNA

Krasnopolsky et al. (2020)

Zika virus (ZIKV)

Knockout

Knockout

Zika

Shue et al. (2021)

Chikungunya virus RNA (CHIKV)

Knockout

RNA

Li et al. (2019)

Chikungunya

Meertens et al. (2019)

Dengue virus (DENV, Flavivirus)

Knockout

Dengue

Labeau et al. (2020)

Virus

Wang et al. (2020)

Disease

Authorsa

Screening type

RNA or DNA virus

Customized by Potting et al. (2017)

GeCKOv2

GeCKOv2

Customized by Wang et al. (2016)

GeCKOv2

Human activityoptimized CRISPR knockout library

Brunello

GeCKOv2

GeCKOv2

GeCKOv2

Library used

19,050

18,360

x

19,050

18,543

19,050

19,166

19,114

19,050

19,050

19,050

Library coverage (target genes)

122,411

122,411

187,536

122,411

182,134

77,441

122,411

122,411

122,411

Library coverage (number of guides)

Human

Human

Human

Human

Human

Human

Human

Human

Human

Human

0,3 0,3

TS576 (GSCs) 4 × 108 1 × 108

HepG2

HAP-1 Cas9

Huh7.5.1

GXR Cas9

x

0,3 0,3

0,5

2.4 × 108

1.6 × 107 1.2 × 108

1 × 108

Jurkat 1.3 × 108 T-lymphocyte cell line 2D10

hPSC-derived neural progenitors

0,3

0,3

Huh7

3 × 108

0,3

RG7834, TENT4A, TENT4B, and ZCCHC14

PAPSS1

GNPTAB and cathepsin B

CD4, CCR5, TPST2, SLC35B2, and ALCAM

ZNF304

ISG15, ATP6V1F, and ATP6V1C1

Integrin ɑvβ5

RACK1

FHL1

DPM1 and DPM3

Library MOI Targets foundb

0,3

x

Number of cells used in the screening

x

HAP-1

HAP-1

Target Cell used in organism the screening

Table 9.1 Description table of articles published using CRISPR libraries to investigate new essential targets for viral infections

Host factors required for surface antigen (HBsAg) expression

Infection capacity

Infection capacity

Infection capacity entry and cell aggregation

Transcription repression

Virus resistance in neural cells

Cellular internalization

Virus replication

Virus permissiveness and pathogenesis

Infection capacity

Phenotype involved

SARS-Cov2

COVID-19

COVID-19

Wei et al. (2020)

Wei et al. (2021)

SARS-Cov2

Influenza A virus (H1N1)

Sharon et al. (2020)

Influenza A virus (H5N1)

Influenza A virus (H5N1)

Influenza

Han et al. (2018)

Influenza A virus (PR8 virus)

Song et al. (2021)

Influenza

Li et al. (2020)

RNA

RNA

RNA

RNA

Knockout

Knockout

Knockout

Knockout

Knockout

Knockout

20,991 genes Monkey (SARS-CoV-2 and human screen); 500 genes (top-bottom genes screen); and 32 genes (human genes screen)

SARS-CoV-2 Monkey screen, 20,991 and human genes; top-bottom genes screen, 500 genes; and human genes screen, 32 genes

84,963 sgRNAs (SARS-CoV-2 screen); 6,208 sgRNAs (top-bottom genes screen); and 148sgRNAs (human gene screen)

SARS-CoV-2 screen, 84,963 sgRNAs; top-bottom genes screen, 6,208 sgRNAs; and human gene screen, 148sgRNAs

SARS-Cov-2 screen: African green monkey (AGM) genomewide CRISPR knockout library (CP0070); 250 top and bottom genes screen:

Human

Human

Human

Human

SARS-Cov-2 screen: African green monkey (AGM) genomewide CRISPR knockout library (CP0070); 250 top and bottom genes screen: CRISPR knockout subpool library (CP1564); and human validation screen: CRISPR knockout library (CP1560)

19,050 genes

19,050 genes

18,675

19,114 genes

122,411

122,411

74,700

77,441

Brunello

GeCKOv2

GeCKOv2

AVANA-4

Virus internalization

Viral entry

Viral entry

SARS-CoV- SMARCA4 and HMGB1 2 screen, 0.3; top-bottom screen, 0.2; and human screen, 0.2

SARS-CoV-2 screen, 2 × 108 cells; top-bottom screen, 4 × 107 cells; and human screen, 2 × 106 Calu-3 cells Vero E6-Cas9 (two first screens) and Calu-3 (human screen)

SMARCA4 and HMGB1 0.3 MOI (SARSCoV-2 screen); and 0.2 MOI (top-bottom screen); and 0.2 MOI (human screen)

2 × 108 Vero-E6Cas9 (SARSCoV-2 screen); 4 × 107 Vero-E6Cas9-v2 (top-bottom screen); and 2 × 106 Calu-3 cells (human screen)

(continued)

Infection capacity and viral entry

Infection capacity and viral entry

Infection 64 putative capacity restriction factors:, e.g., RAE1, NUPL2, TSC1, and TSC2

IGDCC4

Vero E6-Cas9 (two first screens) and Calu-3 (human screen)

0,3

3.4 × 108

SLC35A1 and CIC

WDR7, CCDC115, and TMEM199

2

0,3

0,4

x

1 × 108

x

HEK-293SF

A549 Cas9

A549 Cas9

A549Cas9

C. Sabeus sgRNA (for Vero E6), Brunello library (for Caco2-ACE2), or the Gattinara library (for Calu-3)

Knockout

Antoine et al. (2021)

CRISPR knockout subpool library (CP1564); and human validation screen: CRISPR knockout library (CP1560)

Library used

Brunello (loss of function, LOF) and Calabrese (gain of function, GOF)

Virus

Screening type

Knockout and activation

Disease

RNA or DNA virus

Biering et al. (2021)

Authorsa

Table 9.1 (continued)

1.2 × 108 cells for 0,3 to 0,5 Vero E6 and Caco2-ACE2 and 4-6 × 107 for Calu-3 Vero E6, Caco-2ACE2, and Calu-3

Monkey C. Sabeus and sgRNA: human 20991, Brunello library: 19,114 genes, and Gattinara library: 19,993 genes

C. Sabeus sgRNA: 84,963 sgRNAs, Brunello library: 77,441 sgRNAs, and Gattinara library: 40,964 sgRNAs)

0,3 for both screenings

AP1G1, AP1B1, ATP8B1, IL6R, and CD44

GOF: CD44, ICAM1, OR8A1, OR2T33, OR51S1, GNG13, GRK3, and others; LOF: CHUK/IKKalpha, RIPK4, AP1, CDH1, ROCK1, and others

Library MOI Targets foundb

Brunello, 2.7 × 108, and Calabrese, 2 × 108

Number of cells used in the screening

Calu-3

Target Cell used in organism the screening

Human Brunello, 19,114 genes, and Calabrese, 18,885 (set A) and 18,843 (set B) genes

Library coverage (target genes)

Brunello, 77,441 sgRNAs, and Calabrese, 56,762 sgRNAs

Library coverage (number of guides)

Infection capacity, viral entry and replication

Infection capacity, viral entry and replication

Phenotype involved

Brunello

Brunello

106,435

77,441

77,441

19,482

19,114

19,114

Human

Human

Human

HEK293T

Huh7

A549-ACE2Cas9

10

0,3

1.5 × 108

x

0,3

1 × 108

LDLRAD3, TMEM30A, and CLEC4G

TMEM41B, PI3K, and TMEM106B

S1/S2 boundary SARS-CoV-2 spike

Infection and viral entry

Infection, viral entry, and replication

Virus entry and transmission

Authors: The articles included in this table were those available for access and that used CRISPR libraries and viral infections (viruses that cause human diseases) as a challenge to identify new therapeutic targets Targets found: All those described in the article may not be listed. It was included those highlighted or used for further validation. (number) means the reference number. X represents information not available or not found

b

a

Knockout

Activation Customized CRISPRa library

RNA

Zhu et al. (2022)

SARS-CoV-2

Knockout

COVID-19

Baggen et al. (2021)

Zhu et al. (2021)

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screenings, both for loss of function testing and targeting approximately the same number of genes in the host genome. Furthermore, the gain- and loss-of-function approach can be performed together as was done by Biering et al. (2021) to trace genes responsible for a bidirectional modulation of SARS-CoV-2 infection. These screens uncovered pro-viral and antiviral host factors across highly interconnected host pathways giving even more scientific rigor to the identified targets (Table 9.1). It is worth mentioning that in addition to the library used, the cell type must be determined according to the effects of the pathology/phenotype to be studied. And prior to highlight a gene as a possible target for treatments, it must be rigorously validated in in vivo and in vitro assays. More details of the approaches and important points in the experimental design of whole-genome assays using CRISPR libraries can be seen in Table 9.1, where there is information from a series of recently published articles with various types of viral infections.

4 Conclusions The use of CRISPR libraries for genome-wide screenings to identify therapeutic targets for viral infections appears as a powerful tool, but subject to a lot of care regarding its application and validation. From the initial experimental procedures, such as choosing the library and screening type, the target cell, the viral strain, the and amplification of the library and guaranteeing its coverage, to the strategies for the gene screening pipeline by bioinformatics must be well established and conducted by trained personnel. And finally, the validation of the screened targets to guarantee the observed phenotype must be carried out with scientific rigor in order to make new targets available for possible antiviral therapies.

References Antoine R et al (2021) Bidirectional genome-wide CRISPR screens reveal host factors regulating SARS-CoV-2, MERS-CoV and seasonal coronaviruses. bioRxiv Prepr:1–60 Baggen J et al (2021) Genome-wide CRISPR screening identifies TMEM106B as a proviral host factor for SARS-CoV-2. Nat Genet 53:435–444 Bayat H, Naderi F, Khan AH, Memarnejadian A, Rahimpour A (2018) The impact of CRISPR-Cas system on antiviral therapy. Adv Pharm Bull 8:591–597 Biering SB et al (2021) Genome-wide, bidirectional CRISPR screens identify mucins as critical host factors modulating SARS-CoV-2 infection. bioRxiv:2021.04.22.440848 Bock C et al (2022) High-content CRISPR screening. Nat Rev Methods Prim 2. https://doi.org/10. 1038/s43586-021-00093-4 Chulanov V et al (2021) CRISPR screening: molecular tools for studying virus-host interactions. Viruses 13:2258 Doench JG et al (2016) Optimized sgRNA design to maximize activity and minimize off-target effects of CRISPR-Cas9 synthesis of an arrayed sgRNA library targeting the human genome. Nat Biotechnol 34:184–191

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CRISPR Libraries and Whole-Genome Screening to Identify Essential. . .

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Flint M et al (2019) A genome-wide CRISPR screen identifies N-acetylglucosamine-1-phosphate transferase as a potential antiviral target for Ebola virus. Nat Commun 10:1–13 Han J et al (2018) Genome-wide CRISPR/Cas9 screen identifies host factors essential for influenza virus replication. Cell Rep 23:596–607 Hart T et al (2015) High-resolution CRISPR screens reveal fitness genes and genotype-specific cancer liabilities. Cell 163:1515–1526 Hyrina A et al (2019) A genome-wide CRISPR screen identifies ZCCHC14 as a host factor required for hepatitis B surface antigen production. Cell Rep 29:2970–2978.e6 Joung J et al (2017) Genome-scale CRISPR-Cas9 knockout and transcriptional activation screening. Nat Protoc 12:4 Kaminski R et al (2016) Elimination of HIV-1 genomes from human T-lymphoid cells by CRISPR/ Cas9 gene editing. Sci Rep 6:1–15 Kleinstiver BP et al (2016) High-fidelity CRISPR-Cas9 nucleases with no detectable genome-wide off-target effects. Nature 529:490–495 Kotterman MA, Schaffer DV (2014) Engineering adeno-associated viruses for clinical gene therapy. Nat Rev Genet 15:445–451 Krasnopolsky S, Kuzmina A, Taube R (2020) Genome-wide crispr knockout screen identifies znf304 as a silencer of HIV transcription that promotes viral latency. PLoS Pathog 16:1–22 Labeau A et al (2020) A genome-wide CRISPR-Cas9 screen identifies the dolichol-phosphate mannose synthase complex as a host dependency factor for dengue virus infection. J Virol 94: e01751-19 Lee C (2019) CRISPR/Cas9-based antiviral strategy: current status and the potential challenge. Molecules 24:1349 Li W et al (2015) Quality control, modeling, and visualization of CRISPR screens with MAGeCKVISPR. Genome Biol 16:281 Li Y et al (2019) Genome-wide CRISPR screen for Zika virus resistance in human neural cells. Proc Natl Acad Sci U S A 116:9527–9532 Li B et al (2020) Genome-wide CRISPR screen identifies host dependency factors for influenza a virus infection. Nat Commun 11:164 Makarova KS et al (2011) Evolution and classification of the CRISPR-Cas systems. Nat Rev Microbiol 9:467–477 Meertens L et al (2019) FHL1 is a major host factor for chikungunya virus infection. Nature 574: 259–263 Park RJ et al (2017) A genome-wide CRISPR screen identifies a restricted set of HIV host dependency factors. Nat Genet 49:193–203 Potting C et al (2017) Genome-wide CRISPR screen for PARKIN regulators reveals transcriptional repression as a determinant of mitophagy. Proc Natl Acad Sci U S A 115:E180–E189 Puschnik AS, Majzoub K, Ooi YS, Carette JE (2017) A CRISPR toolbox to study virus-host interactions. Nat Rev Microbiol 15:351–364 Sampson TR, Weiss DS (2014) CRISPR-Cas systems: new players in gene regulation and bacterial physiology. Front Cell Infect Microbiol 4:1–8 Sander JD, Joung JK (2014) CRISPR-Cas systems for editing, regulating and targeting genomes. Nat Biotechnol 32:4 Sharon DM et al (2020) A pooled genome-wide screening strategy to identify and rank influenza host restriction factors in cell-based vaccine production platforms. Sci Rep 10:12166 Shue B et al (2021) Genome-wide CRISPR screen identifies RACK1 as a critical host factor for Flavivirus replication. J Virol 95:e0059621 Slaymaker IM et al (2016) Rationally engineered Cas9 nucleases with improved specificity. Science (80-) 351:84–88 Song Y et al (2021) A genome-wide CRISPR/Cas9 gene knockout screen identifies immunoglobulin superfamily DCC subclass member 4 as a key host factor that promotes influenza virus endocytosis. PLoS Pathog 17:1–24

172

I. M. Paiva et al.

Sun L et al (2021) Genome-scale CRISPR screen identifies TMEM41B as a multi-function host factor required for coronavirus replication. PLoS Pathog 17:1–30 Suzuki T et al (2022) Genome-wide CRISPR screen for HSV-1 host factors reveals PAPSS1 contributes to heparan sulfate synthesis. Commun Biol:1–11. https://doi.org/10.1038/s42003022-03581-9 Wang T, Lander ES, Sabatini DM (2016) Large-scale single-guide RNA library construction and use for genetic screens. Physiol Behav 176 Wang S et al (2020) Integrin αvβ5 internalizes Zika virus during neural stem cells infection and provides a promising target for antiviral therapy. Cell Rep 30:969–983. https://doi.org/10.1016/ j.celrep.2019.11.020 Wei J et al (2020) Genome-wide CRISPR screen reveals host genes that regulate SARS-CoV-2 infection. bioRxiv Prepr Serv Biol. https://doi.org/10.1101/2020.06.16.155101 Wei J et al (2021) Genome-wide CRISPR screens reveal host factors critical for SARS-CoV-2 infection. Cell 184:76–91.e13 Yau EH, Rana TM (2018) Next-generation sequencing of genome-wide CRISPR screens. Methods Mol Biol 1712:203–216 Zhu Y et al (2021) A genome-wide CRISPR screen identifies host factors that regulate SARS-CoV2 entry. Nat Commun 12:1–11 Zhu S et al (2022) Genome-wide CRISPR activation screen identifies candidate receptors for SARS-CoV-2 entry. Sci China Life Sci 65:701–717

Chapter 10

Gene Editing Technologies Targeting TFAM and Its Relation to Mitochondrial Diseases Vanessa Cristina de Oliveira, Kelly Cristine Santos Roballo, Clesio Gomes Mariano Junior, and Carlos Eduardo Ambrósio

1 Mitochondria in the Cell Metabolism Mitochondria are essential organelles present in most eukaryotic cells. They are considered energy transducers due to their ability to drive cellular metabolism, having as a key function the synthesis of adenosine triphosphate (ATP). This synthesis occurs through a process called oxidative phosphorylation (OXPHOS), which generates energy. Thus, this energy-generating mechanism must function properly if the entire organism is to obtain the energy needed to supply its various organs and tissues (Taanman 1999; Kang and Hamasaki 2005; Kang et al. 2007; Gammage and Frezza 2019). In addition to ATP, mitochondria produce metabolic precursors for macromolecules such as lipids, proteins, DNA, and RNA (Spinelli and Haigis 2018). They are also known as the main site of oxidative reactions as they generate metabolic byproducts such as reactive oxygen species (ROS) and ammonia (Spinelli and Haigis 2018; Chen et al. 2022). Mitochondria contain and control their own DNA, named mtDNA, which is responsible for synthesizing 15% of the proteins related to the cell respiratory chain. The nuclear DNA (nDNA) is responsible for protein synthesis, which will V. C. de Oliveira (✉) · C. G. Mariano Junior · C. E. Ambrósio Department of Veterinary Medicine, Faculty of Animal Science and Food Engineering, University of São Paulo, Pirassununga, SP, Brazil e-mail: [email protected]; [email protected]; [email protected] K. C. S. Roballo Biomedical Affairs and Research, Edward Via College of Osteopathic Medicine, Blacksburg, VA, USA Department of Biomedical Sciences and Pathobiology, Virginia Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA, USA e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_10

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help into other mitochondrial functions, such as control and transcription of mtDNA. The perfect organelle operation depends on the interaction between mtDNA and nDNA (Shadek and Clayton 1997). The mtDNA encodes essential proteins involved in carrying out mitochondrial metabolic functions. Compared to nDNA, mtDNA is more susceptible to acquiring somatic mutations, due to the accumulation of higher levels of the product of DNA oxidation, unlike genes encoded by the nucleus. The accumulated DNA damage, if not efficiently repaired, can increase ROS production and may cause mitochondrial dysfunction, inducing mutations and blockage of the electron transport chain, leading to cellular damage, which in turn can result in diverse pathogenesis of a variety of human diseases. Furthermore, when compared to nDNA repair mechanisms, which have been extensively studied, mtDNA repair mechanisms are much less understood (Tsutsui et al. 2009; Douglas et al. 2014; Cappa et al. 2020; Rong et al. 2021).

2 Mitochondrial Genome Mitochondria harbor their own genome, mtDNA, which encodes and regulates the expression of several factors that control mitochondrial DNA replication, transcription, and translation, as well as a molecular machinery responsible for the maintenance of the organelle (Gustafsson et al. 2016). The mitochondrial genetic system is composed of a circular DNA genome, with the human mtDNA comprising 16,569 base pairs and existing in multiple copies. The mtDNA contains 37 genes – 13 proteins that are subunits of the respiratory chain, 2 rRNAs, and 22 transfer RNA (tRNAs) – which are involved in the translation process of mitochondrial RNAs (mtRNAs) and are essential for the synthesis of proteins encoded by mtDNA (Larsson and Clayton 1995; Kang and Hamasaki 2005; Wallace 2005; Gustafsson et al. 2016). There are several copies of mtDNA in each cell, and the number of copies of mtDNA is related to the cell metabolism, function, and type, whereas the number of nDNA is still the same in all cells. Specialized cells that require more energy, such as muscle cells, will have more mtDNA molecules than less energy-consuming cells (Kang and Hamasaki 2005). The copy number of mtDNA is related to the ability to produce ATP, and this synthesis can be compromised when mutated (Chiaratti et al. 2010). Transcription factors, such as nuclear respiratory factors 1 and 2 (NRF1 and NRF2), help in the mitochondrial machinery. For example, NRF1 is involved in mitochondrial biogenesis, signal transduction, and protein synthesis and it functions as a transcription factor that activates the expression of some important genes that regulate cell growth and nuclear genes. When NRF1 and NFR2 are together, they both mediate the expression of several proteins, such as factors controlling mtDNA replication, transcription, and translation (Choi et al. 2011; Zhang et al. 2017).

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In relation to mtDNA replication and transcription, there is a specific site in mtDNA known as the D-loop, which contains the promoter sites for transcription initiation: the heavy-strand promoter (HSP) and light-strand promoter (LSP), as well as regulatory sequences that control mtDNA replication. The L-chain is rich in adenine and thymine, encoding the ND6 protein and eight tRNAs. The H-strand is rich in guanine and encodes the remaining 12 proteins and 14 tRNAs (Wang et al. 2021). Replication of mtDNA relies on DNA polymerase gamma (POLG), which is encoded in the nucleus. Bidirectional transcription and replication depend on factors that occur via the D-loop, with the basic transcription machinery consisting of TFAM, RNA polymerase (POLRMT), and transcription factors B1 and B2 (TFB1M and TFB2M) interactions (Ekstrand et al. 2004; Kang et al. 2007; Alonso-Montes et al. 2008).

3 Mitochondrial Diseases Several mutations in mitochondrial genes encoded in nDNA and mtDNA have been discovered since the 1960s, with many being related to an increasing number of pathologies. What characterizes these mitochondrial diseases are mutations that cause failures in the cell’s energy production process, compromising the electron transport chain and mitochondrial function (May-Panloup et al. 2005). Mitochondrial disorders are genetically heterogeneous and affect about 1 in 2000 individuals. They are caused by pathogenic variants in more than 300 genes encoded in both genomes, mtDNA and nDNA (May-Panloup et al. 2005). In mtDNA, the mutation rate can be 100 times more prevalent than in the nuclear genome, because mtDNA is more susceptible to damage, due to its limited repair mechanisms (Kang et al. 2007; Gorman et al. 2015; Lopes 2020). It is estimated that over 1500 different proteins are required to maintain the structure and function of mitochondria, indicating that the vast majority of them are being encoded by the nuclear genome and imported into the mitochondria. Of these, only 13 proteins participate in the mitochondrial genome (La Morgia et al. 2020). Variations in nuclear gene expression may generate mutations and epigenetic modifications that can affect mitochondrial functionality (Lopes 2020). The mitochondrial genome comprises between 1000 and 2000 nuclear DNA genes and thousands of copies of maternally inherited mtDNA, which at birth are usually all identical, which is called homoplasmy. Pathogenic mutations can be mixed with wild-type mtDNAs, which is known as heteroplasmy, a condition that can increase energetic defects within cells (Lopes 2020; Chinnery et al. 2000). Human diseases characterized by the presence of defects in mitochondrial activity can be hereditary and somatic. These can be caused by mutations in mtDNA as well as can occur due to mutations in nDNA (Ryzhkova et al. 2018). The discovery of mitochondrial diseases began with the reports of Zeviani et al. (1988) who identified large-scale deletions in mtDNA in patients with Kearns-Sayre

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syndrome (KSS) and with the reports of Holt et al. (1988) who showed in their findings deletions in the mtDNA of muscle tissue from patients with mitochondrial myopathies. In 1995, researchers identified a homozygous mutation in the succinate dehydrogenase A (SDHA) gene. This was the first report of a mutation in a disease-causing nuclear gene that encodes mitochondrial proteins, causing an impairment of the respiratory chain in humans. From then on, the list of disease-causing nuclear genes encoding mitochondrial proteins grew exponentially, and much attention has been paid to the investigation of nuclear genes associated with mitochondrial syndromes (Bourgeron et al. 1995). Moreover, the application of next-generation sequencing (NGS) has further increased the pace of discovery of nuclear genes involved in mitochondrial diseases, because it allows to reliably detecting low levels of heteroplasmy (Payne et al. 2013; Stenton and Prokisch 2020). Mitochondrial diseases occur systematically in humans and range from mild to lethal (Nasseh et al. 2001). The common molecular defects found are deletions and duplications, and the vast majority of mutations occur in transfer RNA genes (May-Panloup et al. 2005). Examples of mitochondrial diseases, including recent findings, clinical manifestations, affected ages, and molecular mechanisms, are summarized below. As previously mentioned, Kearns-Sayre syndrome (KSS) is a typical example of mtDNA-related disease, a progressive multisystemic mitochondrial myopathy-like disorder that occurs due to mutations comprising large-scale single and multiple mtDNA deletions and mtDNA duplication. This syndrome is characterized by muscle weakness, retinopathies such as ophthalmoplegia and retinitis pigmentosa, cardiac arrhythmia, neurosensory hearing loss, and ataxia (Goldstein and Falk 2003; Ryzhkova et al. 2018). Another example of mitochondrial-related disease is Pearson marrow-pancreas syndrome, a rare mitochondrial disorder caused by rearrangements of mitochondrial DNA. It was initially described as fatal in childhood; however, patients who survive childhood may also develop KSS later in life. It is characterized by sideroblastic anemia and variable exocrine pancreatic insufficiency. Clinical manifestations include short stature, cognitive deficit, sensorineural hearing loss, chronic renal failure, dementia, renal tubular acidosis, seizures, progressive myopathy, and endocrinopathies (Wild et al. 2020; Arena et al. 2022). Chronic progressive external ophthalmoplegia (CPEO) is a disorder characterized by drooping eyelids (ptosis) and bilateral paralysis of extraocular muscles (ophthalmoplegia). It causes limitation of eye movements, and sometimes with involvement of limbs and bulbar muscles. This disease can be caused by point mutations, single large-scale mtDNA deletions, duplications, or multiple mtDNA deletions secondary to a nuclear mutation, such as in the genes ANT1, POLG1, POLG2, OPA1, C10orf2, and SLC25A4 (Pfeffer et al. 2011; Arena et al. 2022). Myoclonic epilepsy with ragged red fibers (MERRF) is a chronic neurodegenerative disease that can appear in children and adults. MERRF is diagnosed with the presence of myoclonic epilepsy-associated encephalopathy, cerebellar ataxia, myopathy, and optic atrophy. Patients can carry several mtDNA point mutations, with a

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heteroplasmic m.8344A > G mutation being the most frequent. At a lower frequency, m.8356 T > C and m.8363G > A have also been described. Symptoms for this disease are dementia, cardiomyopathy, arrhythmia, optic atrophy, and neurosensory hearing loss (Wallace et al. 1988; Orsucci et al. 2021). Neuropathy, ataxia, and retinitis pigmentosa (NARP) is a disorder that is characterized by neuropathy, ataxia, and retinitis pigmentosa. Characteristic symptoms include seizures, migraines, learning disabilities, developmental delays, sensory neuropathies, and muscle weakness. It is a disease of maternal inheritance, usually developing in childhood, and is caused by mtDNA sequence variants leading to amino acid substitutions in the MT-ATP6 gene that codes for a subunit of the mitochondrial ATP synthase, complex V of OXPHOS. The m.8993 T > G variant substitutes a conserved leucine with an arginine in subunit 6 of the mitochondrial F1F0 ATP synthase. It can increase mitochondrial membrane potential leading to the increase of superoxide production, potentially triggering programmed cell death. In a different pathogenic variant, a leucine is converted to a proline at the same position (m.8993 T > C), which reduces the severity of interference with proton translocation. Furthermore, a heteroplasmic mutation (m.5789 T > C) in the tRNA-Cys gene of the mtDNA is related to late-onset NARP syndrome, which is additionally characterized by dementia, ataxia, pigmentary retinopathy, peripheral neuropathy, and epilepsy (Thorburn et al. 2003; Hippen et al. 2021). Leigh syndrome or subacute necrotizing encephalomyelopathy is a rare inherited neurometabolic disorder caused by damage to mitochondrial energy production that affects the nervous system. It is a heterogeneous neurological disorder that usually begins in early childhood. Genetically, alterations or mutations in the mitochondrial respiratory enzyme complex or the pyruvate dehydrogenase complex are believed to be responsible for the development of the syndrome. Clinical manifestations may include motor delay, mental retardation and/or progressive cognitive decline, hypotonia, dyskinesia, akinesia, ataxia, dystonia, and brainstem dysfunction (Ryzhkova et al. 2018; Chang et al. 2020). Leber hereditary optic neuropathy (LHON) syndrome is characterized by the sudden, complete, and painless loss of visual acuity (central and bilateral) due to optic nerve atrophy. It occurs in a sequential manner: first, the loss of vision in one eye and then the other and can occur at any age. It is characterized by the point mutation 11,778/G > A in mtDNA, affecting the ND4 subunit gene of complex I. Other mutations have also been found at positions 3460/G > A and 14,484/T > C, affecting complex I subunit genes (ND1 and ND6) (Sadun et al. 2013; Ryzhkova et al. 2018). Mitochondrial encephalomyopathy lactic acidosis and stroke-like episodes (MELAS) is a disease appearing during early childhood or early teen years. This syndrome is characterized by cardiomyopathy, diabetes, stroke, seizures, retinitis pigmentosa, renal tubule problems, and lactic acidosis. It is considered a neurodegenerative disease because demyelination of nerve fibers and gradual death of neurons occur. The mutation occurs in the gene for the leucine tRNA, swapping A for G in the mtDNA (A3243G) (Goto et al. 1991).

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4 The Regulatory Role of TFAM and Mitochondrial Function In this section, TFAM functions and “cell” responsibilities were summarized, including recent findings and molecular mechanisms. TFAM was first studied by Fisher and Clayton (1985). They noted that TFAM was extremely necessary for mitochondrial DNA transcription, along with mitochondrial DNA polymerase and transcription factor B2 (TFB2). Many studies in recent years have associated this gene with mtDNA function, precisely because it is involved in controlling mitochondrial mechanics. TFAM is a nuclear-encoded protein that is synthesized in the cytoplasm and imported into the mitochondria. It was first identified as part of the high-mobility group (HMG) chromosomal proteins. TFAM binding to DNA substrates is mostly mediated by electrostatic interactions via their HMG boxes, being an important mitochondrial transcription factor, as it is composed of two domains that stimulates transcription through specific binding to recognition sites on the LSP and HSP promoters (Garstka et al. 1994; Scarpulla 2002; Choi and Garcia-Diaz 2022). TFAM binds to its recognition site and promotes bidirectional transcription, facilitating interaction with mitochondrial RNA polymerase (POLRMT) and activating transcription at the two promoters, HSP and LSP. This activation results in the folding of mtDNA, which is of fundamental importance in activating mitochondrial transcription. The TFAM C-terminus is essential for the activation of transcription (Fig. 10.1) (Garstka et al. 1994; Falkenberg et al. 2002; Hallberg and Larsson 2011; Kukat and Larsson 2013). TFAM is characterized as a single-nuclear copy gene, regulated by NFR-1, as are several other genes. It also plays an important role in mtDNA replication, since replication initiation is dependent on an RNA primer formed by transcription from LSP (Kukat and Larsson 2013). In addition to performing transcription, TFAM also packages mtDNA and is therefore a dual function protein. Unlike nDNA, mtDNA is not associated with histones and within the mitochondrial matrix; there are nucleoprotein complexes associated with the inner membrane, called nucleoids (spherical nucleoprotein complex 100 nm in diameter). Each nucleoid can contain two to ten mtDNA copies. The most abundant structural component of nucleoids is TFAM, whose HMG domains facilitate mtDNA compaction, thereby regulating the accessibility of the mitochondrial genome to the replication and transcription machinery. The D-loop region of mtDNA, which serves as a regulatory locus for mtDNA replication and transcription, anchors mtDNA nucleoids in the inner mitochondrial membrane (Canugovi et al. 2010; Ryzhkova et al. 2018; Filograna et al. 2020). TFAM Responses to Inflammation and Diseases In this section, examples of mitochondrial diseases involving TFAM were summarized, including recent findings, clinical manifestations, affected ages, and molecular mechanisms.

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Fig. 10.1 Schematic representation of transcription and replication of mtDNA. TFAM is a nuclear gene that interacts with POLMRT and TFB2M and binds to the D-loop region of mtDNA and initiates a bidirectional transcription and replication

Recent studies have been relating TFAM to neurodegenerative diseases (Kang et al. 2018). Neurodegenerative diseases lead to the loss of functional neurons, with the most characteristic examples being Parkinson’s disease, Alzheimer’s disease, and Huntington’s disease. TFAM has been shown to play a central role in the mtDNA stress-mediated inflammatory response. Emerging evidence indicates that decreased mtDNA copy number is associated with several aging-related pathologies; however, little is known about the association of TFAM abundance and disease. There are evidences showing that in these neurodegenerative diseases, TFAM is overexpressed (Kim et al. 2010; Sheng et al. 2012; Kang et al. 2018), protecting affected cells against degeneration; furthermore, mtDNA has higher rate of mutation when compared to nDNA (Tuppen et al. 2010) which could lead to mitochondrial dysfunctions; thus TFAM also reduces the effect of these mutations. However, Kang et al. (2018) summarize that TFAM could also be leading to these neurodegenerative conditions, due to TFAM gene mutations or polymorphisms, but the mechanisms associated to these findings are still unclear. TFAM is also associated with other inflammatory diseases such as asthma (Luo et al. 2021). In patients with asthma, there is an increase of bronchial smooth muscle (BSM) cells, which also presents increased proliferation and mitochondrial biogenesis and mass, with TFAM overexpression (Trian et al. 2007). TFAM regulates mtDNA packing and homeostasis, thus indirectly regulating inflammatory response

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(West et al. 2015). In addition, TFAM is overexpressed repressing cytokinemediated signaling pathway, suggesting that the increased TFAM protects the lung epithelial cells from inflammation (Luo et al. 2021). TFAM is also associated with mitochondrial DNA depletion syndrome (Mehmedović et al. 2022), according to a study which used patient’s fibroblast to show that a mutation in TFAM leads to dysfunctional mtDNA transcription initiation, affecting the promoter sequence recognition and the bound between TFAM and tether helix of POLMRT. This study was performed in vitro, using biochemical characterization (Mehmedović et al. 2022). Mitochondrial DNA depletion syndrome is a reduction of mtDNA in specific tissues, such as myocytes (Wang et al. 2022). Clinical manifestations include muscle weakness, dysphagia, hypertrophic cardiomyopathy (Wang et al. 2022), and liver failure in neonates due to hepatocerebral mtDNA depletion syndrome (Stiles et al. 2016). TFAM is also involved in immunomodulatory diseases, for example, in myasthenia gravis, which is an autoimmune disease with incidence of 1.7–21.3/1,000,000 individuals per year (Berrih-Aknin et al. 2014). A recent study showed that TFAM was significantly lower in the peripheral blood mononuclear cells of patients with myasthenia gravis (Li et al. 2022).

5 Gene Editing The possibility of editing mtDNA both in vivo and in vitro opens new perspectives through different gene editing tools that may help model mitochondrial diseases and disorders, with potential development of future treatments for these diseases (Yang et al. 2021). The main methods of editing mitochondrial genes are restriction endonuclease (ER) technology, zinc finger nucleases (ZFN), transcription activator-like effector nucleases (TALENs), and the development of specific mitochondria-targeted nucleases known as mitoTALENS. These are expressed from the nuclear-cytoplasmic compartment, and a mitochondrial localization signal directs the monomers to the mitochondrial matrix, where they bind and cleave the mutant mtDNA (Bacman and Moraes 2020). Recently the clustered regularly interspaced short palindromic repeats and CRISPR-associated protein 9 (CRISPR-Cas9 system), mitoCas9, and DddA-derived cytosine base editor (DdCBE) technique were developed as promising technologies to install targeted mutations or introduce transmissible base conversion mutations in mtDNA (Bacman and Moraes 2020; Yang et al. 2021; Wei et al. 2022). Mitochondrial DNA editing technologies were employed for several mitochondrial disease models and functional analysis of mtDNA genes, such as ER in Leigh disease models and Leber’s hereditary optic neuropathy (LHOND) (Tanaka et al. 2002; Reddy et al. 2015); ZFN for the neuropathy, ataxia, and retinitis pigmentosa (NARP) disease model (Gammage et al. 2016); TALENS/mitoTALENS

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technologies in diseases such as osteosarcoma and Leber’s optic neuropathy dystonia (Bacman et al. 2013); and CRISPR/mitoCas for analyses of the genes ND1, ND4, and noncoding D-loop region (Bian et al. 2019) and COX1 and COX3 (Jo et al. 2015). Unlike CRISPR tools developed for nuclear genome editing, effective CRISPR approaches for mtDNA editing are still limited due to the inability of the guide RNAs to cross mitochondrial membranes and enter the organelle (Yang et al. 2021). In addition to the various strategies used to edit mtDNA, nuclear genes that are directly related to the mitochondrial genome are also being investigated and manipulated, such as the TFAM gene. Different editing technologies have been employed to better understand the relationship of these nuclear genes to mtDNA.

5.1

Gene Editing of TFAM

The TFAM gene is a prime candidate for further investigation because it directly affects mtDNA transcription and replication, as well as being required to regulate mtDNA copy number (Kang et al. 2018; Chen et al. 2018). Several studies have shown that TFAM may also be associated with DNA damage repair. Furthermore, it is known that p53 stimulates repair when it is in the presence of TFAM, and TFAMrelated mtDNA packaging may play a protective role against oxidation, such as ROS (reactive oxygen species), as well as TFAM playing a key role in maintaining mitochondrial stability (Canugovi et al. 2010). The TFAM gene has been disrupted before, for example, in the study by Larsson et al. (1998), who using Cre-loxP observed that mice with a homozygous TFAM mutation exhibited embryonic lethality after mtDNA depletion. The TFAM knockout (KO) mice died soon after birth. Such embryos were smaller, had delayed neural development, and showed cardiac problems caused by oxidative stress and mitochondrial depletion. TFAM was also found to be lethal after its disruption in mice (Cre-Lox recombination) and chickens (targeted gene disruption by expression vectors), a consequence of depletion of mtDNA (Kang and Hamasaki 2005). In order to generate an animal model for mitochondrial myopathy, KO mice for the TFAM gene (skeletal-muscle-specific disruption) were generated using the Crerecombinase technique. The KO animals developed a myopathy with irregular red muscle fibers, accumulation of abnormal appearing mitochondria, and progressive deterioration of respiratory chain function in skeletal muscle (Wredenberg et al. 2002). The same KO animals were further studied, and alterations in cellular handling of Ca2+ were involved in muscle dysfunction caused by mitochondrial myopathy, with results showing reduced sarcoplasmic reticulum storage capacity and lower muscle force in the TFAM KO mice. Tissue-specific TFAM knockout mice with Cre-loxP revealed a progressive cardiac phenotype, mtDNA depletion, and decreased rate of ATP production (Hansson et al. 2004). Another TFAM knockdown by siRNA mice had 60–95% reduction in TFAM gene expression and 50–90% reduction in cytochrome b (Cyt b)

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gene expression, as well as a decrease in cytochrome C subunit I (COX I) protein (Jeng et al. 2008). In vitro KO of TFAM was already performed, for example, the TFAM gene was edited in bovine fibroblasts by CRISPR-Ca9, and the edited heterozygous knockout cells showed decreased activity and lower levels of mitochondrial DNA copy number, indicating that TFAM is directly related to maintaining mtDNA integrity (Oliveira et al. 2019, 2020). Another example is the TFAM knockdown in MKN45 cells by siRNAs showing that TFAM mRNA levels and mtDNA copy number had a decline between 80 and 90% (Lee et al. 2017). A recent study reported a suppression of up to 80% of TFAM expression in a zebrafish study to better understand the role of TFAM in embryonic development, which resulted in a 42% drop in mtDNA-CN. This degree of knockdown was sufficient to alter muscle, heart, brain, and eye development (Otten et al. 2020). Two extremely uncommon cases of a homozygous missense mutation in TFAM were also reported, where an 89% reduction in mtDNA in the patient’s liver and a 79% reduction in mtDNA in muscles caused hepatic failure and newborn death, paired with postnatal developmental abnormalities (Otten et al. 2020). Another study obtained heterozygous TFAM knockout mice with reduced mtDNA and increased oxidative mtDNA damage (Woo et al. 2012). Crossing these mice to the adenomatous polyposis coli multiple intestinal neoplasia mouse cancer model exhibited increased tumor growth in the small intestine in the resulting offspring (Woo et al. 2012). The same process has been demonstrated by other methods of examining this relationship between TFAM disruption and mtDNA content. Human mammary epithelial cells of non-carcinoma (MCF10A) and carcinoma (MCF7) origin had a decrease in mtDNA copies of up to 75–80% when TFAM gene expression was silenced by about 90% (Guha et al. 2014). A recent study performed in 2021 showed a putative tumor-suppressing role of TFAM in head and neck cancer by silencing TFAM with small hairpin RNAs, which resulted in enhanced cancer cell growth, chemoresistance, and motility (Hsieh et al. 2021). Further tests with mouse tongue cancer and human head and neck cancer tissues also showed a negative correlation between TFAM and disease progression (Hsieh et al. 2021). In order to elucidate how alterations in mtDNA are related to cardiovascular disease, using CRISPR-Cas9, heterozygous TFAM KO human embryonic kidney (HEK) 293 T clones were generated. These KO cells demonstrated decreased TFAM levels and reduced mitochondrial protein and copy number. This gives some indication that there was a connection between low levels of mtDNA-CN and abnormally high levels of nDNA CpGs methylation at specific loci, which may be one of the processes underlying how mitochondrial dysfunction influence the incidence and progression of various disorders (Castellani et al. 2020). Also performed in vitro in HEK293T, the effects of CRISPR-Cas9 gene editing of TFAM cells on mtDNA copy number was investigated. Results showed that the mtDNA copy number is directly related to TFAM control, and its disruption results in interference with mitochondrial stability and maintenance (Oliveira et al. 2021).

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When TFAM dysfunctions were analyzed in patients, it is clear how important this gene is. For example, a homozygous missense mutation in TFAM was reported (Stiles et al. 2016). The affected patient had an early onset of fatal liver disease, presenting with severe intrauterine growth restriction, hypoglycemia, hyperbilirubinemia, cholestasis, and ascites in the neonatal period. There is also a connection between Krüppel-like transcription factors – DNA-binding proteins that are essential for the differentiation and growth of malignant tumors – and the control of carcinogenesis, with TFAM either promoting or suppressing the appearance of tumors, depending on the cellular environment. For example, a study in which the levels of KLF16 were altered showed that it aids in the modulation of TFAM in glioma cells, inhibiting (or promoting) the proliferation of malignant cells by acting on this key transcription factor in mitochondria (Chen et al. 2018). Recently TFAM was reported as a candidate gene for premature ovarian insufficiency (POI) in a patient of Pakistani origin with a pathogenic predicted homozygous missense mutation in TFAM (NM_003201.3: c.694C > T, NP_003192.1: p. (Arg232Cys)) (Tucker et al. 2020). The data suggests that pathogenic TFAM variants, and consequently defects in mtDNA maintenance, can be fatal, causing early-onset liver disease and other rare genetic disorders such as Perrault syndrome (Tucker et al. 2020). In another study, a recessive missense variant c.694C > T, p. Arg232Cys in TFAM that segregates with POI disease was identified. After performing TFAM gene editing in zebrafish using CRISPR-Cas9, they noticed that homozygous-mutated TFAM carried a deletion mimicking the ovarian dysgenesis phenotype observed in affected patients (Ullah et al. 2021). Lastly, it has also been shown that TFAM is required for normal nephron differentiation and that the loss of TFAM activity in renal epithelial cells produces molecular and metabolic features associated with a polycystic kidney disease. TFAM disruption in kidney epithelial cells resulted in decreased mitochondrial gene expression, a 63% reduction in mtDNA copy number – which is consistent with mtDNA depletion, a hallmark of TFAM deficiency – which in turn led to inhibition of nephron maturation and development of severe postnatal cystic disease (Ishii et al. 2021). As shown above and summarized in Table 10.1, there are several studies looking for the major functions of TFAM, as well as its interaction with mtDNA. Most of these studies have been performed in vitro, with cell lines, patient-derived immortal cell lines, using techniques such as KO, knock-in, siRNAs, simple DNA sequencing (Table 10.1). Thus, there is a need of performing TFAM analyses into primary cell culture models and in vivo models showing the systemic interactions of TFAM.

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Table 10.1 Study models of edition of TFAM Model type Mouse/in vivo

Technique Cre recombinase

Species Mouse

Mouse/in vivo

Cre recombinase

Mouse

Mouse/in vivo

Cre recombinase

Mouse

143B TK cells/in vitro

siRNA

Human

HeLa cells/in vitro

siRNAs

Human

Mouse/in vivo

Cre recombinase

Mouse

MCF10A e MCF7/in vitro

shRNA

Human

HEK-293 T/in vitro Human gastric cancer cell line MKN45/in vitro Melanoma cell lines/ in vitro Melanocyte (FM308)/ in vitro Fibroblasts/in vitro

CRISPR/Cas9 siRNAs

Human Human

Mitochondrial genome sequences

Human

Mitochondrial genome sequences

Human

CRISPR-Cas9

Bovine

Zebrafish/in vivo

Zebrafish

HEK293/in vitro

Knockdown by splicing modificated morpholino oligonucleotides CRISPR-Cas9

Zebrafish/in vivo

CRISPR-Cas9

Zebrafish

Renal epithelial cells/ in vitro

Cre recombinase

Mouse

Human

Reference Larsson et al. (1998) Wredenberg et al. (2002) Hansson et al. (2004) Jeng et al. (2008) Canugovi et al. (2010) Woo et al. (2012) Guha et al. (2014) Jo et al. (2015) Lee et al. (2017) Araujo et al. (2018) Araujo et al. (2018) Oliveira et al. (2019, 2020) Otten et al. (2020) Oliveira et al. (2021) Ullah et al. (2021) Ishii et al. (2021)

6 Conclusion and Perspectives Understanding mitochondrial mechanisms and cellular function is crucial for identifying dysfunctionalities within this organelle. Numerous studies have focused on investigating the relationship between mtDNA and nDNA, specifically the genes and proteins that play a significant role in maintaining mitochondrial functionality. This chapter provides a comprehensive overview of primary mitochondrial functions and diseases while emphasizing the significance of TFAM in mitochondrial processes. The TFAM gene is indispensable for mitochondrial function and survival, serving as a key factor in maintaining mtDNA and its copy number, making it an excellent candidate for clinical studies. Although there are still many aspects of mitochondrial maintenance and function yet to be discovered, the advent of novel

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techniques like CRISPR-Cas9 for editing mtDNA and mitochondria-related nDNA holds promise for addressing these areas in the near future. This progress has the potential to lead to enhanced in vitro and in vivo models for mitochondrial diseases.

References Alonso-Montes C, Castro MG, Reguero JR et al (2008) Mitochondrial transcription factors TFA, TFB1 and TFB2: a search for DNA variants/haplotypes and the risk of cardiac hypertrophy. Dis Markers 25(3):131–139 Araujo LF, Siena ADD, Plaça JR et al (2018) Mitochondrial transcription factor a (TFAM) shapes metabolic and invasion gene signatures in melanoma. Sci Rep 8(1):14190 Arena IG, Pugliese A, Volta S, Toscano A, Musumeci O (2022) Molecular genetics overview of primary mitochondrial myopathies. J Clin Med 11(3):632 Bacman SR, Moraes CT (2020) MitoTALENs for mtDNA editing. In: Giuseppe G. Academic Press, Porcelli AM The Human Mitochondrial Genome, pp 481–498 Bacman SR, Williams SL, Pinto M, Peralta S, Moraes CT (2013) Specific elimination of mutant mitochondrial genomes in patient-derived cells by mitoTALENs. Nat Med 19(9):1111–1113 Berrih-Aknin S, Frenkian-Cuvelier M, Eymard B (2014) Diagnostic and clinical classification of autoimmune myasthenia gravis. J Autoimmun 48-49:143–148 Bian WP, Chen YL, Luo JJ, Wang C, Xie SL, Pei DS (2019) Knock-in strategy for editing human and zebrafish mitochondrial DNA using Mito-CRISPR/Cas9 system. ACS Synth Biol 8(4): 621–632 Bourgeron T, Rustin P, Chretien D et al (1995) Mutation of a nuclear succinate dehydrogenase gene results in mitochondrial respiratory chain deficiency. Nat Genet 11:144–149 Canugovi C, Maynard S, Bayne ACV, Sykora P, Tian J, de Souza-Pinto NC, Croteau DL, Bohr VA (2010) The mitochondrial transcription factor a functions in mitochondrial base excision repair. DNA Repair 9:1080–1089 Cappa R, de Campos C, Maxwell AP, McKnight AJ (2020) “Mitochondrial toolbox” a review of online resources to explore mitochondrial genomics. Front Genet 11:439 Castellani CA, Longchamps RJ, Sumpter JA et al (2020) Mitochondrial DNA copy number can influence mortality and cardiovascular disease via methylation of nuclear DNA CpGs. Genome Med 12(1):84 Chang X, Wu Y, Zhou J, Meng H, Zhang W, Guo J (2020) A meta-analysis and systematic review of Leigh syndrome: clinical manifestations, respiratory chain enzyme complex deficiency, and gene mutations. Medicine (Baltimore) 99(5):e18634 Chen X, Li S, Ke Y et al (2018) KLF16 suppresses human glioma cell proliferation and tumourigenicity by targeting TFAM. Artif Cells Nanomed Biotechnol 46: 608–615 Chen Q, Li ZH, Song WQ et al (2022) Association between single nucleotide polymorphism of rs1937 in TFAM gene and longevity among the elderly Chinese population: based on the CLHLS study. BMC Geriatr 22:16 Chiaratti MR, FERREIRA CR, MEIRELLES FV, MÉO SC, PERECIN F, SMITH LC et al (2010) Xenooplasmic transfer between buffalo and bovine enables development of homoplasmic offspring. Cell Reprogram 12(3):231–236 Chinnery PF (2000) Primary mitochondrial disorders overview. In: Adam MP, Everman DB, Mirzaa GM et al (eds) GeneReviews® [Internet]. University of Washington, Seattle, pp 1993–2022 Choi WS, Garcia-Diaz M (2022) A minimal motif for sequence recognition by mitochondrial transcription factor A (TFAM). Nucleic Acids Res 50(1):322–332 Choi YS, Kim S, PAK YK (2011) Mitochondrial transcription factor A (mtTFA) and diabetes. Diabetes Res Clin Pract 2: 3–9

186

V. C. de Oliveira et al.

Douglas AP, Vance DR, Kenny EM, Morris DW, Maxwell AP, McKnight AJ (2014) Nextgeneration sequencing of the mitochondrial genome and association with IgA nephropathy in a kidney transplant population. Sci Rep 4:7379 Ekstrand M, Falkenberg M, Rantanen A, Park CB, Gaspari M, Hultenby K, Rustin P, Gustafsson CM, Larsson NG (2004) Mitochondrial transcription factor a regulates mtDNA copy number in mammals. Hum Mol Genet 13:935–944 Falkenberg M, Gaspari M, Rantanen A, Trifunovic A, Larsson NG, Gustafsson CM (2002) Mitochondrial transcription factors B1 and B2 activate transcription of human mtDNA. Nat Genet 31(3):289–294 Filograna R, Mennuni M, Alsina D, Larsson N (2020) Mitochondrial DNA copy number in human disease: the more the better? FEBS Lett 595:976–1002 Fisher RP, Clayton DA (1985) A transcription fator required for promoter recognition by human mitochondrial RNA polymerase. Accurate initiation at the heavy and light strandpromoters dissected and reconstituted in vitro. J Biol Chem 260: 11330–11338 Gammage PA, Gaude E, Van Haut L et al (2016) Near-complete elimination of mutant mtDNA by iterative or dynamic dose-controlled treatment with mtZFNs, Nucleic Acids Research 44(16): 7804–7816 Gammage PA, Frezza C (2019) Mitochondrial DNA: the overlooked oncogenome? BMC Biol 17: 53 Garstka HL, Facke M, Escribano JR, Wiesner RJ (1994) Stoichiometry of mitochondrial transcripts and regulation of gene expression by mitochondrial transcription factor A. Biochem Biophys Res Commun 200(1):619–626 Goldstein A, Falk MJ (2003) Mitochondrial DNA deletion syndromes. In: Adam MP, Everman DB, Mirzaa GM, Pagon RA, Wallace SE, LJH B, Gripp KW (eds) Amemiya A GeneReviews. University of Washington, Seattle, pp 1993–2022 Gorman GS, Schaefer AM, Ng Y, Gomez N, Blakely EL et al (2015) Prevalence of nuclear and mitochondrial DNA mutations related to adult mitochondrial disease. Ann Neurol 77(5): 753–759 Goto Y, Nonaka I, Honai S (1991) A new mtDNA mutation associated with mitochondrial myopathy, encephalopathy, lactic acidosis and stroke-like episodes (MELAS). Biochemical and Biophysical Acta 1097(3):238–240 Guha M, Srinivasan SP, Ruthel G (2014) Mitochondrial retrograde signaling induces epithelial– mesenchymal transition and generates breast cancer stem cells. Oncogene 33:5238–5250 Gustafsson CM, Falkenberg M, Larsson NG (2016) Maintenance and expression of mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160 Hallberg BMH, Larsson NG (2011) TFAM forces mtDNA to make a U-turn. Nat Struct Mol Biol 18(11):1179–1181 Hansson A, Hance N, Dufour E, Rantanen A, Hultenby K, Clayton DA, Wibom R, Larsson NG (2004) A switch in metabolism precedes increased mitochondrial biogenesis in respiratory chain-deficient mouse hearts. Proc Natl Acad Sci USA 101(9):3136–3141 Hippen M, Zsurka G, Peeva V, Machts J, Schwiecker K, Debska-Vielhaber G, Wiesner RJ, Vielhaber S, Kunz WS (2021) Novel pathogenic sequence variation m.5789T>C causes NARP syndrome and promotes formation of deletions of the mitochondrial genome. Neurol Genet 8(2):e660 Holt IJ, Harding AE, Morgan-Hughes JA (1988) Deletions of muscle mitochondrial DNA in patients with mitochondrial myopathies. Nature 331(6158):717–719 Hsieh YT, Tu HF, Yang MH, Chen YF et al (2021) Mitochondrial genome and its regulator TFAM modulates head and neck tumourigenesis through intracellular metabolic reprogramming and activation of oncogenic effectors. Cell Death Dis 12(11):1–11 Ishii K, Kobayashi H, Taguchi K, Guan N, Li A et al (2021) Kidney epithelial targeted mitochondrial transcription factor a deficiency results in progressive mitochondrial depletion associated with severe cystic disease. Kidney Int 99(3):657–670

10

Gene Editing Technologies Targeting TFAM and Its Relation. . .

187

Jeng JY, Yeh TS, Lee JW, Lin SH, Fong TH, Hsieh RH (2008) Maintenance of mitochondrial DNA copy number and expression are essential for preservation of mitochondrial function and cell growth. J Cell Biochem 103(2):347–357 Jo A, Ham S, Lee GH, Lee YI, Kim S, Lee YS, Shin JH, Lee Y (2015) Efficient mitochondrial genome editing by CRISPR/Cas9. Biomed Res Int 2015:305716 Kang D, Hamasaki N (2005) Mitochondrial transcription factor a in the maintenance of mitochondrial DNA overview of its multiple roles. Ann N Y Acad Sci 1042:101–108 Kang D, Kim SH, Hamasaki N (2007) Mitochondrial transcription factor a (TFAM): roles in maintenance of mtDNA and cellular functions. Mitochondrion 7:39–44 Kang I, Chu CT, Kaufman BA (2018) The mitochondrial transcription factor TFAM in neurodegeneration: emerging evidence and mechanisms. FEBS Lett 592(5):793–811 Kim J, Moody JP, Edgerly CK, Bordiuk OL, Cormier K, Smith K, Beal MF, Ferrante RJ (2010) Mitochondrial loss, dysfunction and altered dynamics in Huntington’s disease. Hum Mol Genet 19:3919–3935 Kukat C, Larsson N-G (2013) mtDNA makes a U-turn for the mitochondrial nucleoid. Trends Cell Biol 23:457–463 La Morgia C, Maresca A, Caporali L, Valentino ML, Carelli V (2020) Mitochondrial diseases in adults. J Intern Med 287:592–608 Larsson NG, Clayton DA (1995) Molecular genetic aspects of human mitochondrial disorders. Annual Review Genetics 29: 151–178 Larsson NG, Wang J, Wilhelmsson H, Oldfors A, Rustin P, Lewandoski M, Barsh GS, Clayton DA (1998) Mitochondrial transcription factor a is necessary for mtDNA maintenance and embryogenesis in mice. Nat Genet 18:231–236 Lee WR, Na H, Lee SW, Lim WJ, Kim N, Lee JE, Kang C (2017) Transcriptomic analysis of mitochondrial TFAM depletion changing cell morphology and proliferation. Sci Rep 7:17841 Li L, Cai D, Zhong H, Liu F, Jiang Q, Liang J, Li P, Song Y, Ji A, Jiao W, Song J, Li J, Chen Z, Li Q, Ke L (2022) Mitochondrial dynamics and biogenesis indicators may serve as potential biomarkers for diagnosis of myasthenia gravis. Exp Ther Med 23(4):307 Lopes A (2020) Mitochondrial metabolism and DNA methylation: a review of the interaction between two genomes. Clin Epigenetics 12(1):182 Luo J, Liu H, Li DKJ et al (2021) Repression of the expression of proinflammatory genes by mitochondrial transcription factor a is linked to its alternative splicing regulation in human lung epithelial cells. BMC Immunol 22:74 May-Panloup P, Vignon X, Chretien MF, Heyman Y, Tamassia M, Malthiery Y et al (2005) Increase of mitochondrial DNA content and transcripts in early bovine embryogenesis associated with upregulation of mtTFA and NRF1 transcription factors. Reprod Biol Endocrinol 3:65 Mehmedović M, Martucci M, Spåhr H, Ishak L, Mishra A, Sanchez-Sandoval ME, PardoHernández C, Peter B, van den Wildenberg SM, Falkenberg M, Farge G (2022) Disease causing mutation (P178L) in mitochondrial transcription factor a results in impaired mitochondrial transcription initiation. Biochim Biophys Acta Mol Basis Dis 1868(10):166467 Nasseh IE, Tengan CH, Kiyomoto BH, Gabbai AA (2001) Doenças mitocondriais. Revista Neurociências 9 (2): 60–69 Oliveira VC, Moreira GSA, Bressan FF, Junior CGM, Roballo KCS, Charpentier M, Concordet JP, Meirelles F, Ambrósio CE (2019) Edition of TFAM gene by CRISPR/Cas9 technology in bovine model. PLoS One 14:e0213376 Oliveira VC, Junior CGM, Belizário JE, Krieger JE et al (2020) Characterization of post-edited cells modified in the TFAM gene by CRISPR/Cas9 technology in the bovine model. PLoS One 15(7):e0235856 Oliveira VC, Roballo KCS, Junior CGM, Santos SIP, Bressan FF, Chiaratti MR, Tucker EJ, Davis EE, Concordet JP, Ambrósio CE (2021) HEK293T cells with TFAM disruption by CRISPRCas9 as a model for mitochondrial regulation. Life (Basel) 12(1):22 Orsucci D, Caldarazzo Ienco E, Rossi A, Siciliano G, Mancuso M (2021) Mitochondrial syndromes revisited. J Clin Med 10(6):1249

188

V. C. de Oliveira et al.

Otten ABC, Kamps R, Lindsey P et al (2020) Tfam knockdown results in reduction of mtDNA copy number, OXPHOS deficiency and abnormalities in zebrafish embryos. Front Cell Dev Biol 8: 381 Payne BA, Wilson IJ, Yu-Wai-Man P, Coxhead J, Deehan D, Horvath R, Taylor RW, Samuels DC, Santibanez-Koref M, Chinnery PF (2013) Universal heteroplasmy of human mitochondrial DNA. Hum Mol Genet 22:384–390 Pfeffer G, Sirrs S, Wade NK, Mezei MM (2011) Multisystem disorder in late-onset chronic progressive external ophthalmoplegia. Can J Neurol Sci 38(1):119–123 Reddy P, Ocampo A, Suzuki K, Luo J et al (2015) Selective elimination of mitochondrial mutations in the germline by genome editing. Cell 161(3):459–469 Rong Z, Tu P, Xu P, Sun Y, Yu F, Tu N, Guo L, Yang Y (2021) The mitochondrial response to DNA damage. Front Cell Dev Biol 9:669379 Ryzhkova AI, Sazonova MA, Sinyov VV et al (2018) Mitochondrial diseases caused by mtDNA mutations: a mini-review. Ther Clin Risk Manag 14:1933–1942 Sadun AA, La Morgia C, Carelli V (2013) Mitochondrial optic neuropathies: our travels from bench to bedside and back again. Clin Exp Ophthalmol 76:702–712 Scarpulla RC (2002) Nuclear activators and coactivators in mammalian mitochondrial biogenesis. Biochim Biophys Acta 1576(1–2):1–14 Shadek GS, Clayton DA (1997) Mitochondrial DNA maintenance in vertebrates. Annu Rev Biochem 66:409–435 Sheng B, Wang X, Su B, Lee HG, Casadesus G, Perry G, Zhu X (2012) Impaired mitochondrial biogenesis contributes to mitochondrial dysfunction in Alzheimer’s disease. J Neurochem 120: 419–429 Spinelli JB, Haigis MC (2018) The multifaceted contributions of mitochondria to cellular metabolism. Nat Cell Biol 20:745–754 Stenton SL, Prokisch H (2020) Genetics of mitochondrial diseases: identifying mutations to help diagnosis. EBioMedicine 56:102784 Stiles AR, Simon MT, Stover A, Eftekharian S et al (2016) Mutations in TFAM, encoding mitochondrial transcription factor a, cause neonatal liver failure associated with mtDNA depletion. Mol Genet Metab 119(1–2):91–99 Taanman JW (1999) The mitochondrial genome: structure, transcription, translation and replication, Biochimica et Biophysica Acta (BBA). Bioenergetics 1410(2):103–123 Tanaka M, Borgeld HJ, Zhang J, Muramatsu S et al (2002) Gene therapy for mitochondrial disease by delivering restriction endonuclease SmaI into mitochondria. J Biomed Sci 9(6):534–541 Thorburn DR, Rahman J, Rahman S (2003) Mitochondrial DNA-associated leigh syndrome and NARP. In: Adam MP, Ardinger HH, Pagon RA, Wallace SE, LJH B, Mirzaa G, Amemiya A (eds) GeneReviews. University of Washington, Seattle, pp 1993–2021 Trian T, Benard G, Begueret H, Rossignol R, Girodet P-O, Ghosh D, Ousova O, Vernejoux J-M, Marthan R, Tunon-de-Lara J-M (2007) Bronchial smooth muscle remodeling involves calciumdependent enhanced mitochondrial biogenesis in asthma. J Exp Med 204(13):3173–3181 Tsutsui H, Kinugawa S, Matsushima S (2009) Mitochondrial oxidative stress and dysfunction in myocardial remodelling. Cardiovasc Res 81:449–456 Tucker EJ, Rius R, Jaillard S, Bell K, Lamont PJ, Travessa A, Dupont J, Sampaio L, Dulon J, Vuillaumier-Barrot S et al (2020) Genomic sequencing highlights the diverse molecular causes of Perrault syndrome: a peroxisomal disorder (PEX6), metabolic disorders (CLPP, GGPS1), and mtDNA maintenance/translation disorders (LARS2, TFAM). Qual Life Res 139:1325–1343 Tuppen HA, Blakely EL, Turnbull DM, Taylor RW (2010) Mitochondrial DNA mutations and human disease. Biochim Biophys Acta 1797:113–128 Ullah F, Rauf W, Khan K, Khan S, Bell KM, de Oliveira VC, Tariq M, Bakhshalizadeh S, Touraine P, Katsanis N et al (2021) A recessive variant in TFAM causes mtDNA depletion associated with primary ovarian insufficiency, seizures, intellectual disability and hearing loss. Qual. Life Res 140:1733–1751

10

Gene Editing Technologies Targeting TFAM and Its Relation. . .

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Wallace DCA (2005) A mitochondrial paradigm of metabolic and degenerative diseases, aging, and cancer: a dawn for evolutionary medicine. Annu Rev Genet 39:359–407 Wallace DC, Lott MT, Hodge JA, Schur TG, Lezza AMS, IILJ ELSAS, Nikoskelainen EK (1988) Mitochondrial DNA mutation associated with Leber’s hereditary optic neuropathy. Science 242:1427–1430 Wang F, Zhang D, Zhang D, Li P, Gao Y (2021) Mitochondrial protein translation: emerging roles and clinical significance in disease. Front Cell Dev Biol 9:675465 Wang H, Han Y, Li S, Chen Y, Chen Y, Wang J, Zhang Y, Zhang Y, Wang J, Xia Y, Yuan J (2022) Mitochondrial DNA depletion syndrome and its associated cardiac disease. Front Cardiovasc Med 8:808115 Wei Y, Li Z, Xu K, Feng H, Xie L, Li D, Zuo Z, Zhang M, Xu C, Yang H, Zuo E (2022) Mitochondrial base editor DdCBE causes substantial DNA off-target editing in nuclear genome of embryos. Cell Discov 8(1):27 West AP, Khoury-Hanold W, Staron M, Tal MC, Pineda CM, Lang SM, Bestwick M, Duguay BA, Raimundo N, MacDuff DA (2015) Mitochondrial DNA stress primes the antiviral innate immune response. Nature 520(7548):553–557 Wild KT, Goldstein AC, Muraresku C, Ganetzky RD (2020) Broadening the phenotypic spectrum of Pearson syndrome: five new cases and a review of the literature. Am J Med Genet A 182(2): 365–373 Woo DK, Green PD, Santos JH, D'Souza AD, Walther Z, Martin WD, Christian BE, Chandel NS, Shadel GS (2012) Mitochondrial genome instability and ROS enhance intestinal tumorigenesis in APC(Min/+) mice. Am J Pathol 180(1):24–31 Wredenberg A, Wibom R, Wilhelmsson H, Graff C, Wiener HH, Burden SJ, Oldfors A, Westerblad H, Larsson NG (2002) Increased mitochondrial mass in mitochondrial myopathy mice. Proc Natl Acad Sci USA 99(23):15066–15071 Yang X, Jiang J, Li Z, Liang J, Xiang Y (2021) Strategies for mitochondrial gene editing. Comput Struct Biotechnol J 19:3319–3329 Zeviani M, Moraes CT, DiMauro S, Nakase H, Bonilla E, Schon EA, Rowland LP (1988) Deletions of mitochondrial DNA in Kearns-Sayre syndrome. Neurology 38(9):1339–1346 Zhang GM, Deng MT, Lei ZH, Wan YJ, Nie HT, Wang ZY, Fan YX, Wang F, Zhang YL (2017) Effects of NRF1 on steroidogenesis and apoptosis in goat luteinized granulosa cells. Reproduction 154(2):111–122

Chapter 11

The Gene-Edited Babies Controversy: Reactions in the Scientific Community, Social Media, and the Press Morgan Meyer and Frédéric Vergnaud

1 Introduction At the end of November 2018, biophysicist He Jiankui, then at the Southern University of Science and Technology in Shenzhen, posted five videos on YouTube. The most infamous of these videos is titled About Lulu and Nana: Twin Girls Born Healthy After Gene Surgery As Single-Cell Embryo (duration: 4′43″). In the video – whose narrative style is a mix of public announcement, ethical justification, and personal confession – He announces the birth of Lulu and Nana1 who underwent gene surgery at the single-cell stage. He states that the mother’s pregnancy was “normal,” that the gene surgery worked “safely,” and that both Lulu and Nana are “as healthy as any other babies.” The main motivation behind the experiment, he explains, was to provide the babies an “equal chance and a healthy life” as their father suffers from HIV. While describing and justifying his experiment, He draws upon medical, scientific, ethical, and social arguments. He makes, for instance, a historic comparison with in vitro fertilization in order to dedramatize his announcement, he refers to social problems (such as exclusion), and he tries to defend a family-centered ethics. Aware of the potential negative backlash, he declares “I understand my work will be controversial, but I believe families need this technology and I’m willing to take the criticism for them.” He’s announcement has indeed caused an intense “storm” (Lovell-Badge 2019; Daley et al. 2019) and quickly became a hotly debated “affair” (Greely 2019) within the scientific world. But also beyond the scientific world, He’s announcement has led 1

Both names are pseudonyms.

M. Meyer (✉) · F. Vergnaud Centre for the Sociology of Innovation, Mines Paris (Paris Sciences & Lettres, CNRS), Paris, France e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5_11

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to numerous debates, articles, and discussions. Both in the scientific and the public sphere, there has been a rather strong condemnation of He’s experiment. “Irresponsible,” “illegal,” and “unethical” are some of the terms frequently used to disqualify his experiment. But at a closer look, we see that reactions differ between the spheres where He’s experiment is discussed. In this chapter, we therefore analyze how the controversy sparked by He Jiankiu has unfolded in three different spheres: the scientific community, the press, and social media. The reaction of the scientific community will be analyzed by focusing on the second international summit on human genome editing (held from 27 to 29 November 2018, at the University of Hong Kong). Reactions in social media will be discussed via a literature review. And the press will be analyzed through a selection of articles compiled via the Europresse database. This chapter thus builds upon and complements previous work on the topic: our own work analyzing scientific publications and meetings (Meyer 2022; Meyer and Vergnaud 2021); analyses of social media such as Twitter or Weibo (Calabrese et al. 2020; Zhang et al. 2021; Ni et al. 2022); and analyses of the press (Lee 2020; Marcon et al. 2019). But while these academic articles have so far analyzed the controversy within one given world, our chapter aims to compare and contrast discussions across three worlds (science, the press, and social media).

2 Scientific Community The Second International Summit on Human Genome Editing was convened in 2018 by the Academy of Sciences of Hong Kong, the Royal Society, the US National Academy of Sciences, and the US National Academy of Medicine. We have analyzed the recorded video webcasts (a total of 22 h of discussion) and the final statement of the summit in detail. While the 2018 summit resembled the first summit held in 2015 in terms of scholarly format, interdisciplinary focus, publicness, and the writing of a final statement, the circumstances were entirely different. The most intense discussion was He Jianjui’s talk and the subsequent question-and-answer session. He had been invited to speak at the summit, but the organizers didn’t know the story that was going to break when inviting him, and there were discussions within the organizing committee about whether to disinvite him. When introducing He’s talk, the chair of the session, biologist Robin Lovell-Badge, gave some guidelines: He would have “a chance to explain what he’s done” and the audience needed to allow him to speak “without interruptions.” In the question-and-answer session that followed He’s talk (see Picture 11.1), a large number of questions were asked. The first questions concerned the CCR5 gene, its possible effects on other bodily functions, and the reasons for doing research on this specific gene. Before the question period was opened to the floor, David Baltimore, the chair of the summit, came to the stage and made a few comments. He argued that the use of gene editing of human embryos “would still be considered

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Picture 11.1 Image extracted from the video About Lulu and Nana: Twin Girls Born Healthy After Gene Surgery As Single-Cell Embryos which shows He Jiankui explaining his experiment. (Author: The He Lab. License: Creative Commons Attribution 3.0 Unported)

irresponsible,” that it was not a “transparent” process, and that it was not medically necessary. He concluded by saying that He’s experiment represented a “failure of self-regulation by the scientific community.” After Baltimore’s statement, the floor was opened up to questions from the audience. A large number of questions were posed: Why were alternative techniques such as sperm washing not used? With whom was the trial discussed? How were ethics reviewed? What was He’s responsibility? How did he manage to recruit and convince parents? What were the sources of funding? Did He expect and anticipate the “fuss”? How was informed consent gained? How could the wellbeing of the babies be proven if their identity remained secret? Was He willing to openly share the paper he had submitted and the consent forms that he had used? In sum, the questions and criticisms revolved around the technical, ethical, practical, financial, legal, and communicational aspects of He’s experiment. In his responses, He defended his research and gave some more details about the process. He also explained that he felt “proud” of his achievement – a term quoted in many press articles reporting on the summit. One of the final questions was, why, despite there being an international consensus, had he “crossed this line” and done it “in secret.” Why, in other words, had He not openly shared his plans and why did he choose to do something that the scientific community considered so controversial? The question addressed to He Jiankui were almost all critical, and Baltimore’s comments were unequivocal. The response from scientists – at least those who spoke at the summit – was a strong condemnation and disqualification of He’s work, representing a clear exclusion of his work from what is considered “responsible” research practice.

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In their closing comments, the organizers of the summit came back to the issue of responsibility. Victor Dzau, the president of the US National Academy of Medicine, said that not following guidelines such as those created by the Nuffield Council on Bioethics (2016) and those by the US National Academies of Sciences, Engineering, and Medicine (2017) “will be an irresponsible act” and he called for “responsible scientific conduct.” John Skehel, the vice-president of the UK Royal Society, argued that “there are clear responsibilities for all science of this sort, which begin with the individual but then go to the institution and eventually to the national regulatory authorities.” And Baltimore emphasized the need to “take the responsibility” and read out the final statement of the organizing committee, which holds that “proceeding with any clinical use of germline editing remains irresponsible at this time.” About He’s research in particular, the final statement contained the following lines: At this summit we heard an unexpected and deeply disturbing claim that human embryos had been edited and implanted, resulting in a pregnancy and the birth of twins. We recommend an independent assessment to verify this claim and to ascertain whether the claimed DNA modifications have occurred. Even if the modifications are verified, the procedure was irresponsible and failed to conform with international norms. Its flaws include an inadequate medical indication, a poorly designed study protocol, a failure to meet ethical standards for protecting the welfare of research subjects, and a lack of transparency in the development, review, and conduct of the clinical procedures. (Organizing committee of the Second International Summit on Human Genome Editing 2018)

He’s announcement was thus not treated as a proven fact, but as a provisional knowledge claim that needed further verification. The statement not only disqualified He’s announcement; it also criticized his whole experiment on a variety of scientific, medical, legal, and moral grounds. It did so, in particular, by saying what the experiment was not: transparent, ethical, and safe. In sum, we see that this disqualification was based upon a threefold process: There was distrust in He’s results, the entirety of the process that led to these results was criticized, and the lack of connection to existing collective entities (norms and standards) was condemned. The severe tone of the statement and the fact that He’s results were disqualified on so many grounds (epistemic, procedural and normative) reveal just how “disturbing” the situation was for the scientific community. As a response, the organizing committee produced a strict demarcation between, on the one hand, the disciplined and visible spaces of “proper” science and, on the other, the undisciplined and secretive space in which He’s experiment was carried out. The final statement of the summit was not the only collective reaction to He’s announcement. The German Ethics Council stated that using gene editing on embryos was “irresponsible” and represented “a serious violation of ethical obligations” (26 November 2018), the Genetics Society of China and the Chinese Society for Stem Cell Research issued a joint statement to “strongly condemn it for the extreme irresponsibility, both scientifically and ethically” (27 November), and the French National Consultative Ethics Committee issued a statement declaring that “a red line has been crossed” and called for a “stronger global governance” (29 November). In December 2018, the World Health Organization established an expert panel on the governance and oversight of human gene editing (that stated that

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it would be irresponsible to proceed with clinical applications). The Southern University of Science and Technology in Shenzhen distanced itself from He’s work, and investigations were launched to determine the legality of the experiment. Then, on 21 January 2019, He was fired from his university (for an extended discussion on reactions in China see Yan and Mitcham (2022)). Calling He’s experiment “irresponsible” was a means to do demarcation work between “bad” and “good” scientific practice. He’s experiment was positioned outside of proper science, in an illegitimate and undisciplined space. Responsible science was, at the same time, positioned as an activity carried out in a public and disciplined space, a space shaped by collective and transparent norms, values, and ethics. One can speculate that many scientists were concerned that the controversy was likely to have a chilling effect on public attitudes, values, and preferences regarding human gene editing, as well as negative consequences regarding funding and regulation. This demarcation work was arguably a means for the scientific community to restate and protect its moral authority and to reassure the public. If so, then calling He Jiankui “irresponsible” would be not only a form of critique. It represented, in essence, an excommunication of He by the scientific community – and the 2018 summit provided the perfect stage to do so. It is interesting to note, here, that if we look at the content of scientific publications concerned with the politics of gene editing, we observe a striking homogeneity (Meyer and Vergnaud 2021). Several themes are recurrent, such as ethics, governance, benefits, risks, and public debate. The general framing of the debate about the politics of gene editing is something almost universally shared (even though the specific kinds of benefits being discussed might differ and some countries are more precautious than others).

3 Social Media While the controversy around He Jiankui has been discussed at conferences and in many articles, only a few articles have specifically analyzed discussions in social media. Using semantic network analysis, Calabrese et al. (2020) have examined discussions on Twitter. They show that different clusters of words can be differentiated. Before He’s announcement, the main cluster (73%) revolves around the applications of gene editing. After the event, the main cluster revolves around the “CRISPR babies” news (59%) – with the most central words being baby, CRISPR, scientist, twin, and edit – and the second largest cluster is about the negative concerns related to the story (26%) (Calabrese et al. 2020: 961). Overall, their sentiment analysis reveals a clear shift after He’s announcement, with more negative comments (42%) after He’s announcement, than before (35%) (Calabrese et al. 2020: 962). The comments made about the video About Lulu and Nana: Twin Girls Born Healthy After Gene Surgery As Single-Cell Embryo have been analyzed by Meyer (2018). He has noted the presence of negative terms such as “shame” and the use of

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positive terms such as “brave” and “proud” and “congratulations.” Via a multidimensional text analysis (using the software iRaMuTeq), three main clusters of words are identified: a cluster called “ontologies and effects of gene editing” that revolves around the modification of genes and its effects on human health; a cluster called “responsible research and medicine” concerned with the ethics and the wider historical and social context; and a cluster called “irresponsible research” that contains many negative qualifications. Zhang et al. (2021), on their hand, have analyzed frames, emotions, and metaphors in discussions on the platform Weibo. They show that frames and emotions differ according to social actors (government, professionals, journalists, laypeople). For instance, laypeople are more concerned with social issues and display fear and anger, while the government is more concerned with scientific developments and displays – like professionals – more hope. They also show how the debates differ before and after He’s announcement: issues such as scientific development decrease, whereas scandals, ethics, laws, and regulations increase in frequency and emotions such as hope and rejoicing decrease while anger, disgust, and shame increase. Ni et al. (2022) compared discussions across Weibo, Twitter, Reddit, and YouTube. Like the previous studies, they reveal variety across time. They also show that Twitter and Weibo include more critical posts than – in that order – YouTube and Reddit. And they note a difference in terms of the language used, with, for instance, comments on Reddit being more “multidimensional” and “thoughtful” and those on YouTube being more “dramatic” (Ni et al. 2022: 11). Also comparing discussion across various platforms, Ji et al. (2022) find that discussions are more nuanced and diverse on Twitter than on Weibo. They show that different subjects are discussed: on Weibo, only three topics are present (the international summit and the controversy, scientific aspects, and the Chinese investigation), while on Twitter, there are nine topics (including ethical, technological, communicational, and legal aspects which together account for about two-thirds of the topics). The above-quoted studies all point to the same direction: There is a significant variety of comments in and across social media. While the previous section has shown how the scientific community distanced and demarcated itself from He Jiankui, this section shows that in social media, discussions are not so clear cut. We find negative comments as much as positive comments. And while the themes discussed revolve around similar issues – ethics, science, technology, legal frameworks, and politics – in social media, there is more emotionality and dramatization, with more emphasis on national identity and reputability. A number of limits must, however, be highlighted. First, posts in social media are rather short: for instance, an average Tweet is 28 characters long (its limit being 280 characters) and an average post on Weibo is 63 characters long. The press articles we compiled (see next section) are much longer, with an average of 4770 characters. Second, lexical forms are usually not put in context in existing analyses. For example, a given theme and keyword, such as “ethics,” can be linked to different issues and controversies. Ethics can be discussed in a very general and global sense and be discussed in terms of the actors concerned (institutions, collectives, patients,

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scientists, decision-makers, etc.). At the same time, ethics can also refer to the rules (to be) established by ethics committees and/or the more or less implicit rules of the scientific community: How has He Jiankui positioned himself regarding these rules? How does the scientific community position itself? Which rules have been challenged and which ones need to be changed? This shows that we also need to examine the what and the how of the controversy. We need to examine what is at stake and which actors, groups, institutions, and arguments are visible in the debate. This is why we turn now to articles in the press. We will focus on one country, France, to be able to provide an in-depth analysis.

4 The Press In order to analyze the reaction in the press, we proceeded as follows. We searched the Europresse database with the term “He Jiankui” from the period from 1 January 2018 to 30 June 2022. We selected as country of origin and language France/French. Our first request yielded 366 articles, and after several rounds of cleaning, deleting articles that are not pertinent and doublons, we ended up with a final corpus of 309 articles (104 in 2018, 133 in 2019, 45 in 2020, 13 in 2021, and 14 in 2022). In what follows, we present the main themes discussed in this corpus of articles, while indicating the most representative lexical forms in quotation marks (see Fig. 11.1).

Fig. 11.1 Descending hierarchical classification of the 309 articles of our corpus. (36 forms on average per segment, 96.45% of the segments are classified)

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Three Main Classes

Unsurprisingly, the descending hierarchical classification (see Souza et al. 2018) performed on the 309 articles of our corpus reveals a first set of lexical forms about the announcement made by “He Jiankui” in “November” 2018 of the “birth” of the “twins” “Lulu and Nana” (Class 3, ≃ 25% of the classified forms). The French general press (see Table 11.1, Appendix), whether national or regional, echoes the posting on YouTube of the video of the “Chinese” “father,” a “researcher” at the University of “Shenzhen,” who “genetically” manipulated two “babies” by “modifying” their CCR5 gene in order to make it “resistant” to the “AIDS” virus. Class 2 (≃36% of the classified forms) gathers the lexical forms related to the technology used by He Jiankui, the “tool” of the “genetic scissors” “CRISPR” associated with the “Cas9 enzyme,” as well as the consequences that result from it when curing certain “diseases,” in particular those which have as origin a “genetic mutation” and certain cancers. The articles about this subject detail both the technique of “targeting” a particular “DNA sequence” with the aim of “cutting” it, the preliminary experiments on “mice” and the difficulty of moving to the human scale, while contextualizing their remarks by mentioning the Nobel Prize in Chemistry won by Emmanuelle Charpentier and Jennifer Doudna in October 2020. The final classification of lexical forms in our corpus of news articles brings together a more analytical set of themes (≃39,5% of the classified forms). What are the consequences of He Jiankui’s experience on scientific actors, from doctors to researchers in biology? What are the social and political consequences? What moral, legal, and juridical barriers have been crossed as a result of the manipulation and what are the future ethical rules to adopt in order to prevent such an event from happening again? Where should they be adopted? What are the economic repercussions for biotechnology companies but also for the nations that host them? If we observe here the voice of scientific actors who severely criticize the actions and arguments of He Jiankui as to the validity, ethics, and legality of his experiment, the press articles further stress (i) the need to clarify the gray areas of international bioethics but also (ii) the need for a debate on the economic aspects of modern genetics.

4.2

An Epistemic Turning Point

Notwithstanding the controversial nature of the experiment, it still represents a major revolution in science. He Jiankui’s experience takes place in a context in which the ecosystem of “research” is already being disrupted by the penetration of “artificial intelligence” in many scientific activities, including biomedicine. His experience has impacted most social spaces, and, according to our analysis of the French press, “biomedical” scientific research, whether “fundamental,” “genetic,” and “pharmaceutical,” in “bacteriology” or “virology,” is experiencing the most significant shocks.

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He Jiankui is said to have betrayed the scientific world (a betrayal leading to a “storm” in the scientific world, see Lovell-Badge 2019). While a member of the scientific community, he carried out research in fields that are considered to be “forbidden,” that were “not authorized,” “not validated” by his peers. He “trampled” on the “ethical” rules, the “responsibilities” collectively defined at the local and global levels by his colleagues, using “forgery” to achieve his ends. As a researcher, He Jiankui has discredited, at startling speed, via a short YouTube video, a whole profession, a whole set of norms built together over time. The press articles are in line with the conclusions of the scientific community gathered at the 2018 summit while condemning “ethically disturbing” and “dangerous” research. This quasi-traumatic episode can be considered as an epistemological turning point, and it required a rapid response (as was the case at the 2018 summit). The academic community mobilized and positioned itself and tried to clarify the situation. A “debate” must take place. The birth of genetically modified twins poses a “dilemma” and a “challenge”: what “values,” “norms,” and “standards” should researchers and their institutions adopt? What “structures” should biology researchers have in place to define a new “ethical and moral framework” for genetic editing? The event represents an epistemic turning point since from now on, there is a before and an after “Lulu and Nana.” Before November 2018, press articles – at least in the USA and Canada – show a rather “strong promotion” of gene editing (Marcon et al. 2019). An event of this magnitude arises animosity and the search for those responsible. According to He Jiankui himself, states are pointed at concerning the “control of the consequences” of this technological upheaval. He Jiankui’s betrayal could only have been committed in a favorable context: the existence of regulatory disparities between nations. At the time of the events, China did have a text, the “Ethical Guiding Principle on Human Embryonic Stem Cell Research” drafted in 2003, which prohibits the implantation of a genetically modified human embryo in a womb. However, according to the French government, this text is directly subject to the very political interpretations of the Ministry of Health and the Ministry of Science and Technology, which greatly limits its scientific and legal value (Maesen 2019).

4.3

Geopolitics

The articles in our corpus point out that the debate also lies beyond the scientific sphere of biomedical research, as nation states hold a responsibility in “controlling the consequences” of the technological upheaval. Disparities in regulations oppose nations. On the one hand, there are those nations who have “national ethics committees” and who have adopted texts “prohibiting human cloning,” thus proscribing “any reimplantation of genetically modified embryos,” and more generally any “risky experimentation.” In this group we find, among others, the signatories of the Oviedo 2 convention which “protects human rights.” On the other hand, we find

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countries with “less strict legislation” such as China, or which have not “adhered to the Oviedo 2 convention,” such as the USA, the United Kingdom, and Russia. This group of nations thus draws the contours of a “Wild West of medical research,” in which a “soft law” reigns, with “sub-calibrated laws,” which calls into question the “definition of the person.” These “nonbinding norms” result in experiments that only lead to “public disapproval.” We find here an extension of the opposition described by the scientists at the 2018 summit between “good science” and “bad science”: Here, the dichotomy puts “good countries” and “bad countries” back to back in terms of scientific ethics. In addition to these existing divergences between different geopolitical areas, the absence of an “international” framework for the “regulation” of genome editing methods seems to have facilitated the performance of an extraordinary experiment by a researcher based in a country, which, at the time, had no normative and legally binding text on “genetic modifications carried out on embryos for reproductive purposes.” Texts that have been adopted after the event of the birth of Lulu and Nana, like the final statement of the organizing committee of the 2018 summit, are far from being unanimously accepted across nations. The WHO “has set up a registry” and has committed itself to making available the information it has gathered and has “announced that it will set up a group of experts” (which eventually materialized in March 2019). The WHO is therefore called upon to accentuate its role as an international regulator, by creating “a bioethics hotline” and by “assisting in the drafting” of bioethics laws. In addition, the WHO is also called upon to educate and achieve greater “public education” on genome editing. In addition, in the articles we have selected, we see reflections on the place of countries in the competition for innovation. China, a “technologically ambitious and independent” nation, is described as being “at the forefront of innovation.” What is also at stake in the gene editing controversy is the role that those territories and firms that are able to reproduce these experiments will play in the international competition of biomedical research. The dispute over the paternity of the CRISPR-Cas9 method – which opposes the two Nobel Prize winners Doudna and Charpentier to Zhang since 2016 – is not the only one: Issues of “intellectual property and patents” filed in the future by new “biotech startups” are bringing to light the increased competition over the “exploitation rights” of the new technology. France, a country with a “tradition of moratoriums” which, from this point of view, has accumulated a “significant delay” due to “very strict legislation,” is experiencing “difficulties in competing internationally.” Its “economic competitiveness and national security” are “threatened,” but a “revision of the laws of bioethics can change this.” Finally, the articles in our corpus address the social consequences of the experiments conducted by He Jiankui, and of gene editing in general, which have raised “hopes” and “expectations” in “civil” and “human” societies. Hopes to cure certain genetic diseases or cancers, as we have seen, but also expectations in terms of equality: “Genome editing could accentuate discrimination and divisions within society.” The method of gene editing could be a “danger for society” and lead to a “society of castes” in which the income of individuals will determine different rights, such as “increasing one’s intelligence” (since the modification of the CCR5 gene is

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likely to improve memory and cognitive faculties and since gene therapies tend to be expensive). Thus, “citizen consultations” and “debates” are needed, in order to render “acceptable” the new methods of genomic manipulation. This concern with ethics and democracy also raises legal issues. The articles in our corpus question the “fundamental rights of human beings” to “life, health, and respect”. He Jiankui’s genetic manipulation “violated human rights,” which in turn must be better “protected,” especially the rights of children who were genetically manipulated at the embryonic stage.

5 Discussion Across the scientific community, the press, and social media, we observe several similarities. In these three spheres, the debate about He Jiankui is multilayered. Scientific and technical aspects are discussed, as much as legal frameworks, ethics, responsibilities, politics, and social issues. As a whole, the debates make explicit the various kinds of entanglements between genetics and society. In all three spheres, we also see that He Jiankui’s experiment is “hotly” debated. The event is a very prominent and visible controversy across spheres and has taken people by surprise. Beyond these similarities, there are three notable differences. The first concerns geopolitics. This aspect is discussed most explicitly in the press, with debates about scientific competition between countries, about legal and regulatory differences and divergences between nation states. While not entirely absent in social media, the geopolitical nature of the controversy is, however, much less visible – at least in the scientific articles we have reviewed here. And within the scientific community, we see a different kind of politics being articulated. At the 2018 international summit, we observed efforts of self-regulation and demarcation to protect the moral and epistemic authority of the scientific community and to critique – and eventually “excommunicate” – He Jiankui. At the same time, the need for “international” regulation was stressed (but the competition between nation states is less visible than in the press). While the 2018 summit drew a clear line between “good science” and “bad science,” in the press, the dichotomy is rather expressed in terms of “good countries” versus “bad countries” in terms of their ethics and regulation. The second difference concerns emotions and affect. In the scientific sphere, emotions are rather absent. He Jiankui’s experiment is discussed in a serious, technical, and factual way. Despite the fact that the scientific community strongly condemned and criticized He Jiankui, it nevertheless expressed its criticisms in a rather impersonal way. In the press and social media, however, various emotions are visible: fear, anger, pride, hope, rejoicing, disgust, shame, etc. At the same time, the discussion is more dramatic, with references to the “Wild West,” security, threat, and danger. Third, the use of positive terms is also notable: While virtually absent in the scientific sphere and in the press, we do find positive assessments in social media. Words such as “brave” and “proud” are used in comments that approve of He’s experiment, and the word “congratulations” appears several times.

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In this chapter we have presented and discussed empirical data of two spaces: the 2018 international summit on genome editing and press articles from France. Further research could examine how the controversy unfolded in other scientific spaces and in other countries.2 The controversy sparked by He Jiankui could also be further compared to other major controversies in the life sciences, such as the one around the birth of Dolly the sheep. The latter also took the scientific community by surprise and led to heated discussions around regulations (and their gray areas), risk, ethics, and governance (see Petersen 2002). Yet, unlike the controversy sparked by He Jiankui, Dolly the sheep and, especially, the prospect of human cloning, also led to debates about nature, individuality, integrity, genetic uniqueness and was framed in terms of “unacceptable” research (while He Jiankui’s experiment was framed as “irresponsible”). All in all, both cases show that a strong decoupling between innovation and the governance of research is a key motor in controversies. We thus want to argue that the political facets of controversies need to be taken much more into account: further analyses need to trace the arguments and effects in terms of policy, both at the national and the international level.

Appendix Table 11.1 Journals and number of articles containing “He Jiankui” in the French press. (Source: Europresse, date range: 01/11/2018–30/06/2022) Journal Sciences et Avenir Le Monde La Croix Libération Le Figaro Courrier International Les Echos Le Quotidien du Médecin Ouest France Aujourd hui en France HuffPost La Recherche Le Point.fr L’Obs La Tribune

Number of articles 34 33 29 20 19 13 13 12 11 9 9 9 8 6 6 (continued)

2

The limits of the database we used, Europresse, also need to be highlighted: It contains a lot of data scrapped from the web, French and English sources are predominant, and sources from Asia, Africa, and South America are underrepresented.

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Table 11.1 (continued) Journal 20 minutes L’Express Science et Vie Sud-Ouest Atlantico Centre Presse Challenges Charente Libre Revue juridique personnes et famille Corse Matin Courrier picard L’Usine Nouvelle La Dépêche du Midi Le Courrier de l’Ouest Le Journal de Saône et Loire Le Maine Libre Le Progrès Le Télégramme Midi Libre Presse Océan Télérama Valeurs Actuelles Historia Industrie et Technologies L’Echo républicain L’Est Républicain L’Humanité L’Union La Nouvelle République La Vie La Voix du Nord Le Berry républicain Le Monde diplomatique Le Nouveau Magazine Littéraire Manière de voir Pèlerin Planet Science et Vie Junior Yahoo Total

Number of articles 5 5 5 5 3 3 3 3 3 2 2 2 2 2 2 2 2 2 2 2 2 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 309

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References Calabrese C, Ding J, Millam B, Barnett GA (2020) The uproar over gene-edited babies: a semantic network analysis of CRISPR on twitter. Environ Commun 14(7):954–970 Daley GQ, Lovell-Badge R, Steffann J (2019) After the storm—a responsible path for genome editing. N Engl J Med 380(10):897–899 Greely HT (2019) CRISPR’d babies: human germline genome editing in the ‘He Jiankui affair’. J Law Biosci 6(1):111–183 Ji J, Robbins M, Featherstone JD, Calabrese C, Barnett GA (2022) Comparison of public discussions of gene editing on social media between the United States and China. PLoS One 17(5): e0267406 Lee YY (2020) Genome editing or genome cutting? Communicating CRISPR in the British and German press. Yearbook German Cogn Linguist Assoc 8(1):45–66 Lovell-Badge R (2019) CRISPR babies: a view from the Centre of the storm. Development 146(3): dev175778 Maesen S (2019) Le scandale des bébés CRISPR fait encore couler beaucoup d’encre. France Diplomatie Marcon A, Master Z, Ravitsky V, Caulfield T (2019) CRISPR in the north American popular press. Genet Med 21(10):2184–2189 Meyer M (2018) Irresponsible research? Dis/qualifying the gene editing of human embryos. i3 Working Papers Series, 18-CSI-01 Meyer M (2022) Taking responsibility, making irresponsibility: controversies in human gene editing. Soc Stud Sci 52(1):127–143 Meyer M, Vergnaud F (2021) The geographies and politics of gene editing: framing debates across seven countries. Front Polit Sci 3:731496 Ni C, Wan Z, Yan C, Liu Y, Clayton EW, Malin B, Yin Z (2022) The public perception of the# GeneEditedBabies event across multiple social media platforms: observational study. J Med Internet Res 24(3):e31687 Organizing Committee of the Second International Summit on Human Genome Editing (2018) On Human Genome Editing II, 29 November 2018 Petersen A (2002) Replicating our bodies, losing our selves: news media portrayals of human cloning in the wake of Dolly. Body Soc 8(4):71–90 Souza M, Wall M, Thuler A, Lowen I, Peres A (2018) The use of IRAMUTEQ software for data analysis in qualitative research. Rev Esc Enferm USP 52 Yan P, Mitcham C (2022) The gene-edited babies controversy in China: field philosophical questioning. Soc Epistemol 35(4):379–392 Zhang X, Chen A, Zhang W (2021) Before and after the Chinese gene-edited human babies: multiple discourses of gene editing on social media. Public Understanding Sci 30(5):570–587

Index

A A549, 135, 140, 160, 167 Acampomelic campomelic dysplasia (ACD), 18 ACE2, 160, 164 Acquired immune deficiency syndrome (AIDS), 25, 198 Acronym, vi, 2 Adaptation, 2, 3, 7, 118 Adaptive defense, 2 Adeno-associated virus (AAVs), 16, 22–24, 26, 31, 89, 90, 165 Adenosine triphosphate (ATP), 173, 174, 177, 181 Adipocytes, 25 Adoptive transfer, 85, 100 Allogeneic, 87, 89, 90, 92, 96–100 Alzheimer’s disease, 24, 140, 179 Amyloid beta (Aβ), 24 Amyotrophic lateral sclerosis (ALS), 22, 23, 29 Animal models, 13, 14, 17, 19, 67, 68, 74, 132, 133, 135, 136, 138–140, 142, 145, 150, 181 AntagomiRs, 78 Anti-CD19, 90–92, 95–98 AntimiRs, 78 Antisense oligonucleotides (ASOs), 47, 78 Antitumoral, 76, 78, 79, 100 APOBECs, 26

B Babies, vii, 191–202 Bacterial artificial chromosomes (BACs), 19, 23

Bacteriophage, vi, 2, 3, 157 Bad science, 200, 201 Base editors, 18, 25, 26, 66, 89, 99, 132, 146, 148, 180 Base excision repair (BER), 15 Bioinformatics, 163–164, 170 Biotechnology, 1, 4, 198 Blood clotting, 22, 24 Breast cancer, 45, 49, 50, 66

C Calu-3, 160, 167, 168 Campomelic dysplasia (CD), 18 Cancer, vii, 13, 18, 19, 29, 43–50, 52, 53, 59– 69, 73–80, 85–88, 92, 93, 95, 96, 142, 182, 184, 198, 200 Cardiomyocytes, 23, 139 Cas (associated genes), 2, 3, 14, 32, 46, 66–69, 87, 131, 132 Cas1, 3 Cas1-Cas2 enzymatic complex, 3 Cas2, 3 Cas9, 4, 14, 46, 61, 74, 87, 114, 132, 157, 184, 198 Cas9-expressing pigs, 34 Cas9n, 15, 78, 79 Cas12a, 7, 8, 15, 16 Cas13, 7, 8, 48, 51, 52 CCR5, 31, 135, 140, 144, 148, 166, 192, 198, 200 Chagas disease (CD), 111, 122 Charpentier, E., vi, 198, 200

© The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 G. A. Passos (ed.), Genome Editing in Biomedical Sciences, Advances in Experimental Medicine and Biology 1429, https://doi.org/10.1007/978-3-031-33325-5

205

206 Chimeric antigen, 85 Chimeric antigen receptors (CARs), 52, 85–100 Clustered, vi, 1, 46, 114, 131, 180 Clustered regularly interspaced short palindromic repeats (CRISPR), vi, vii, 4–6, 47, 49, 50, 59–69, 90, 93, 115, 132, 157–170, 181, 195, 198 Coagulation, 24 Combinatorial screen, 63, 65 Controversy, vii, 191–202 COVID-19, 53, 165, 167, 169 Cpf1, 15, 16, 90 Cre-recombinase, 15, 28, 29, 181, 184 CRISPRable, 47 CRISPR activation (CRISPRa), 7, 8, 26, 47–50, 64–65, 158, 160, 165, 169 CRISPR array, 3 CRISPR-Cas system, 2, 4–6, 8, 46 CRISPR expression, 2, 3 CRISPR interference (CRISPRi), 7, 8, 26, 47– 51, 64–65, 157, 160 CRISPRko, 158, 160–162, 164, 165 CRISPRoff, 51 CRISPR RNA (crRNA), vi, 3–5, 32, 46, 90 crRNA:tracrRNA, 5 Cytotoxic T lymphocyte-associated molecule-4 (CTLA-4), 85, 86, 91, 92

D Dead Cas9 (dCas9), 6, 7, 26, 48, 49, 51, 52, 64, 65, 158 Dengue, 8, 165, 166 DGCR8, 77 Diabetes, 19, 26, 177 DICER, 77 Disease aetiology, 13, 23 DNA endonuclease targeted CRISPR transreporter (DETECTR), 7, 8 Donor DNA, 6, 113, 158 Double-strand breaks (DSBs), vi, 3–6, 14–17, 26, 27, 47, 48, 65, 66, 87, 88, 90, 99, 113–116, 131, 132, 158 Doudna, J.A., vi, 5, 113, 198, 200 DROSHA, 77 Duchenne muscular dystrophy (DMD), 18, 21, 23, 26 Dystrophin, 21, 23

E Electroporation, 15, 16, 19, 67, 68, 93, 113, 117, 144, 148 Embryo, 15, 17, 27, 32, 44, 68, 181, 191–195, 199, 200

Index Embryonic stem cells, 13, 16, 17, 23, 27, 28, 68, 199 Endocrine, 73 Endonuclease, 3–7, 14, 15, 23, 25, 27, 28, 32, 77, 87, 89, 90, 113, 132, 157, 158, 160, 161, 180 Engineering, 4, 15, 46, 67, 85–100, 194 Epidermal growth factor receptor (EGFR), 76, 80 Epigenome, 50–52, 65 Escherichia coli, vi, 1, 46, 115, 117 Ethics, 191, 193–202 Eukaryotes, 4 Exportin-5, 77

F FACS-based selection, 163 Factor VIII, 22 Fluorescence-activated cell sorting (FACS), 62 Frontotemporal dementia (FTD), 22, 23

G GENCODE, 47 Gene editing tool, 32, 65, 132–144, 146, 180 Gene-targeting, 13 Gene therapies, vi, 14, 19, 21–26, 30, 46, 64, 86, 88, 147, 150, 201 Genetically engineered mouse models (GEMMs), 15, 17, 26 Genetic memory, 4 Genome editing, vi, vii, 4, 6, 13–34, 46, 47, 50, 85–100, 111–115, 127–150, 181, 192, 194, 200, 202 Genome sequence, 74, 111, 116, 132, 184 Glioblastoma, 18, 19, 93, 94, 139 Glycosylphosphatidylinositol (GPI), 21 Good science, 200, 201 Graft-versus-host disease (GvHD), 22, 87, 89, 97, 99 Green fluorescent protein (GFP), 52, 63, 116, 117, 161 Gt(ROSA)26Sor, 20

H H19, 44, 45, 49 Haemophilia A, 22 Haemophilia B, 24 HEK cells, 161, 162 HEK293T, 66, 169, 182 Hepatic steatosis, 19 HNH, 4, 5, 14, 15 Hodgkin’s disease, 45

Index Homeobox C locus (HOXC), 45 Homology-directed repair (HDR), 5, 6, 14, 16– 17, 23, 24, 28, 29, 31, 32, 47, 48, 52, 68, 87–90 Homozygous, 32, 138, 141, 176, 181–183 HOXD, 45, 46 HOX Transcript Antisense RNA (HOTAIR), 45 Human Brunello, 160 Human diseases, 13, 14, 17–19, 23–25, 29, 30, 43, 165, 169, 174, 175 Human GeCKO, 160 Human immunodeficiency virus (HIV), 25, 31, 165, 166, 191 Humanised mouse models, 19–23 Human leukocyte antigen (HLA), 86, 89 Hunter syndrome, 128, 143 Huntingtin gene (HTT), 23 Hyperglycaemia, 19 Hypomorphic missense mutation, 21 Hypothalamus, 19

I i-GONAD, 16 Immune checkpoint (IC), 85–87, 89, 91–93 Inflammation, 24, 44, 139, 140, 178, 180 Insertions/deletions (indels), 5, 6, 14, 21, 47, 48, 87, 146 In silico, 46, 111, 161 In situ, 25, 52 Interference, 2, 3, 47, 50, 52, 177, 182 In vitro, 7, 8, 31, 32, 34, 59–64, 74–76, 90–92, 97, 98, 100, 114, 115, 117, 118, 120, 121, 132, 133, 138, 139, 144, 146, 164, 170, 180, 182–184, 191 In vivo, vii, 14, 18, 28, 32–34, 64, 66–69, 75, 78, 91, 95–100, 121, 132, 135, 141, 142, 144, 145, 147, 164, 165, 170, 180, 183, 184

K Knock-in, 6, 13, 15, 18, 27, 28, 30, 31, 33, 34, 90, 91, 117, 141, 142, 183 Knockouts (KOs), vi, 6, 13, 16, 19, 22, 45, 47, 48, 50, 52, 60, 62, 63, 68, 90, 92, 94, 95, 99, 112, 113, 117–121, 133–138, 140, 141, 143, 144, 158, 161–164, 166–169, 181–183 Kras, 18, 68 Krüppel-associated box (KRAB), 7, 26, 49, 51, 64

207 L Lateral meningocele syndrome (LMS), 17 Lentivirus, 16, 61, 161, 162 Leptin, 19, 25 Leukaemias, 18, 29 Library, vi, 49, 60–62, 157–170 LincRNAs, 42, 45, 46 Lkb1, 18, 68 Long non-coding RNA (lncRNA), 20, 41–52, 77–80 LoxP, 15, 16 Lysosomal, 127–130, 134, 136, 137, 139, 140, 147, 149 Lysosomal disorders, 127–150 Lysosomal storage, 127–150 Lysosomes, 127–131, 137–139, 141, 143, 145

M Major histocompatibility complex (MHC), 22, 86, 98 Mammalian genomes, 14, 33–34, 41, 43 MAPK, 49, 75–76 Mdx mouse, 23, 26 Medulloblastoma, 18 Metastasis, 24, 43–46, 64, 78 Metastasis Associated Lung Adenocarcinoma Transcript 1 (MALAT1), 45, 49, 50 Methylation, 30, 44, 45, 50, 51, 182 Methyltransferases, 48, 51 Microhomology, 65, 114 Microinjection, 15–18, 22, 27, 33 MicroRNAs (miRNAs), 43, 51, 77–80, 140 Mitochondria, 43, 173–175, 178, 181, 183, 184 Mitochondrial, 43, 95, 118, 120, 136, 142, 174– 176, 184 Mitochondrial diseases, 173, 175–178, 180, 184 Mitochondrial DNA (mtDNA), 173–184 Mitochondrial transcription factor A (TFAM), 173, 175, 178–184 Mobile genetic elements, 3 Mosaicism, 17, 68 Motif, 4, 93, 113, 118 Mouse for Actively Recording Cells 1 (MARC1), 33 Mouse models, 17–19, 21–26, 29, 30, 45, 64, 75, 94, 95, 141, 142 Mucopolysaccharidosis, 128, 129, 146, 149 Multiplicity of infection (MOI), 61, 159, 162, 163, 167, 168 Myofibers, 18, 23, 24

208 N Negative selection, 61–62, 163 Neurodegenerative diseases, 22, 29, 140, 142, 176, 177, 179 Nickase, 15, 27, 65, 78, 99, 132, 149 Non-coding RNAs (NcRNAs), 41–43, 50, 52 Non-homologous end joining (NHEJ), 5, 6, 14, 16–17, 29, 32, 33, 47, 48, 52, 65, 87, 114, 158 Nuclear DNA (nDNA), 173–175, 178, 179, 182, 184 Nuclear Enriched Abundant Transcript 2 (NEAT2), 45 Number of genes, 41, 92, 170

O Obesity mouse model, 25 Off-target, 7, 15, 27, 32, 49, 66, 92, 98, 118, 149, 165 Oncology, 59, 60 Open read frame (ORF), 21, 47 Orthologues, 19–23, 28, 118 Osteogenesis imperfecta (OI), 18 Oxidative phosphorylation (OXPHOS), 173, 177

P P53, 68 Palindromic, vi, 1, 46, 114, 131, 180 Parents, 193 Pathogen-associated molecular patterns (PAMPs), 159 Phage infections, 3 Plasmids, vi, 2, 3, 17, 22, 25, 31, 46, 60, 61, 68, 112–117, 149, 157, 158, 160–163 Point mutations, vi, 20, 65, 66, 88, 136, 176, 177 Polyadenylation (PolyA), 42, 47–49 Positive selection, 62, 148, 164 Precursor crRNA (pre-crRNA), 3 Pre-implantation, 32 Prime-editors (PE), 65–66, 132 Prokaryotic genomes, 1 Prokaryotic immune system, 3 Proto-oncogenes, 93 Protospacer adjacent motif (PAM), 4–6, 14, 15, 17, 23, 27, 47, 51, 78

R RAS mutations, 75 Reactive oxygen species (ROS), 136, 142, 173, 174, 181

Index Recombinase polymerase amplification (RPA), 7 Repair, 5, 6, 8, 14, 15, 17, 18, 22, 23, 25–27, 29, 32, 47, 48, 65, 66, 87, 113–119, 131, 132, 148, 174, 175, 181 Reverse transcriptase, 27, 66, 132 Reverse-transcription, 8 RNA, 3, 14, 41, 66, 77, 90, 112, 132, 157, 173 RNA-induced silencing complex (RISC), 77 RNA interference (RNAi), 3, 4, 47, 112, 119 RNA polymerase II, 41 RNase III, 3 RNP complex, 3, 5, 117, 118, 144, 148 Rosa26, 20, 26, 28–32, 34, 147 RuvC, 4, 5, 14, 15

S Safe harbour loci, 20, 27–31, 33 SARS-CoV-2, 8, 160, 167, 169, 170 Sensitivity enzymatic reporter unlocking (SHERLOCK), 7, 8 sgRNA:DNA, 4 Single-strand DNA (ssDNA), 6, 8 Single stranded oligo DNA nucleotides (ssODNs), 15–17 Skill factors, 16 Social media, 191–202 SRY-box 9 (Sox9), 18 Spacer-decoded RNA, 3 Spacers, 2, 3, 52, 157 Spacer sequences, 1, 2 Staphylococcus aureus (SaCas9), 115, 117, 118 Streptococcus pyogenes (SpCas9), 4, 46, 114–118 Subretinal injection, 24 Suppressor, 43–46, 49, 50, 64, 76–78, 80, 93

T Target, v, vi, 4–8, 14–17, 22, 24, 26, 32, 47, 48, 50–53, 60–64, 66, 74, 75, 77–79, 86–90, 95, 97–99, 112–118, 131–133, 140, 144, 147, 157–162, 164, 165, 168–170 Target DNA, 5–7, 46, 88, 113, 131 T cell receptors (TCRs), 66, 86, 89–94, 97–100 T cells, vii, 25, 26, 29, 52, 64, 66, 85–100 Therapies, 7, 25, 30, 43, 52, 53, 64, 67, 69, 75, 80, 85–100, 131, 140–142, 145, 146, 149, 159, 165–170 Thyroid, 73–80 Toolbox, vi, 6–9, 13–34 Topoisomerase 1 (Top1), 32 Trans-activating crRNA (tracrRNA), 3–5, 46, 90

Index Transcription activator-like effector nucleases (TALENs), 4, 30, 59, 68, 74, 87, 89, 90, 97, 131, 132, 165, 180 Transcription factors, 18, 42, 49, 94, 95, 158, 174, 175, 178, 183 Transduction, 61, 90, 94, 158, 161, 162, 164, 174 Transgenesis, 19 Transposons, 3, 157 Trypanosoma cruzi, 111 Tumor necrosis factor (TNF), 92 Tumour suppressor genes, 18 Tyrosinemia type I, 25

U Upstream, 4–6, 27, 41, 47, 63 UTR, 77

V Vascular endothelial cells, 24 Viral DNA, 3, 7 Viral infections, 2, 8, 29, 140, 160, 163, 165, 166, 169, 170 Virus-host, 159, 160, 163, 164

209 W Whole-genome screening, 159–164

X X-chromosome inactivation, 43, 45, 79 Xenotransplant, 76 X-fragile syndrome, 51 X-inactive-specific transcript (XIST), 45, 79 X-linked Dystrophin (DMD), 21

Z Zebrafish, 33, 68, 135, 136, 138, 140, 141, 144, 182–184 Zika, 8, 165, 166 Zinc-finger nucleases (ZFNs), 4, 30, 68, 87, 97, 113, 114, 131, 132, 144, 146, 149, 180 Zinc finger protein (ZFP), 87, 119 Zygotes, 13, 15–18, 22, 27, 28, 32–34, 67, 68