**Methods in Molecular Biology 2434**

Virginia Arechavala-Gomeza Alejandro Garanto *Editors*

# Antisense RNA Design, Delivery, and Analysis

# M ETHODS IN M OLECULAR B IOLOGY

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

For further volumes: http://www.springer.com/series/7651 For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

# Antisense RNA Design, Delivery, and Analysis

Edited by

# Virginia Arechavala-Gomeza

Ikerbasque, Basque Foundation for Science, Bilbao, Spain; Neuromuscular Disorders Research Group, Biocruces Bizkaia Health Research Institute, Barakaldo, Spain

# Alejandro Garanto

Department of Pediatrics, Department of Human Genetics, Radboud University Medical Center, Nijmegen, The Netherlands

Editors Virginia Arechavala-Gomeza Ikerbasque, Basque Foundation for Science Bilbao, Spain

Neuromuscular Disorders Research Group Biocruces Bizkaia Health Research Institute Barakaldo, Spain

Alejandro Garanto Department of Pediatrics Department of Human Genetics Radboud University Medical Center Nijmegen, The Netherlands

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-0716-2009-0 ISBN 978-1-0716-2010-6 (eBook) https://doi.org/10.1007/978-1-0716-2010-6

© The Editor(s) (if applicable) and The Author(s) 2022. This book is an open access publication.

Open Access This book is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this book are included in the book's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the book's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use.

The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

This Humana imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature.

The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

# Preface

The field of RNA therapeutics has rapidly changed over the last few years. Particularly, two mRNA vaccines for SARS-CoV-2 and a customized oligonucleotide for an n-of-1 trial have revolutionized the expectations in the field. Although the use of RNA-based molecules as treatments has been investigated for several decades, this recent surge of applications has seen more and more researchers embarking on the design and assessment of such therapies. The main objective of this book is to provide basic knowledge and a large collection of methods to facilitate the work of these newcomers to the field who want to exploit antisense technology as a therapeutic intervention.

This book was conceived thanks to the network DARTER (Delivery of Antisense RNA Therapeutics, https://www.antisenserna.eu). DARTER is funded by the EU Cooperation of Science and Technology (COST), which aims to enhance interaction and collaborations between researchers in Europe and other countries (https://www.cost.eu/). Within DARTER, we aim to exchange knowledge and protocols and explore the full potential of antisense technology in an environment that combines academia, industry, and patient representatives. This book depicts the variety of models, delivery systems, and approaches that can be used to assess RNA technology and has combined the expertise of researchers located in ten different countries.

Among the 28 chapters included in this book, we have commissioned several review chapters that cover aspects from the historical development of nucleic acid therapeutics, the clinical applications of antisense oligonucleotides, and considerations to include in the preclinical evaluation of the oligonucleotide-mediated toxicology, to patent issues that may need to be contemplated. The remaining chapters follow a classical protocol structure, and we have divided them attending to the subject covered: the design of antisense technology and their delivery (Part II), the description of the model systems developed to evaluate their efficacy, both in vitro (Part III) and in vivo (Part IV), methods to evaluate preclinically the toxicity associated with these new potential drugs (Part V), and intellectual property considerations (Part VI).

We thank all contributing authors for their tremendous effort including their secret tips in the notes of their protocols. We also thank COST for facilitating the cooperation between the research groups and for making it possible for this book to be published Open Access. We have learned a lot during the revision of this book, and we hope that readers will, too.

Barakaldo, Spain Virginia Arechavala-Gomeza Nijmegen, The Netherlands Alejandro Garanto

# Acknowledgments

This book is based upon work from COST Action DARTER (CA 17103), supported by COST (European Cooperation in Science and Technology).

COST (European Cooperation in Science and Technology) is a funding agency for research and innovation networks. Our Actions help connect research initiatives across Europe and enable scientists to grow their ideas by sharing them with their peers. This boosts their research, career, and innovation.

www.cost.eu

# Contents


#### PART III IN VITRO MODEL SYSTEMS


x Contents


#### PART V SAFETY AND TOXICOLOGY



# Contributors


ALEJANDRO GARANTO • Department of Paediatrics, Amalia Children's Hospital, Nijmegen, The Netherlands; Department of Human Genetics, Radboud University Medical Center, Nijmegen, The Netherlands; Radboud Institute for Molecular Life Sciences (RIMLS), Radboud University Medical Center, Nijmegen, The Netherlands

JAMES W. GILBERT • RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, MA, USA

BRUNO M. D. C. GODINHO • RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, MA, USA

IRENE GONZALEZ-MARTINEZ • Interdisciplinary Research Structure for Biotechnology and Biomedicine (ERI BIOTECMED), Universidad de Valencia, Valencia, Spain; Translational Genomics Group, Incliva Health Research Institute, Valencia, Spain; Joint Unit Incliva-CIPF, Valencia, Spain

REMKO GOOSSENS • Department of Human Genetics, Leiden University Medical Center, Leiden, The Netherlands


STEFANO GUSTINCICH • Central RNA Laboratory, Istituto Italiano di Tecnologia, Genova, Italy; Area of Neuroscience, Scuola Internazionale Superiore di Studi Avanzati (SISSA), Trieste, Italy

KWAN-LEONG HAU • UCL Institute of Ophthalmology, London, UK

PABLO HERRERO-HERNANDEZ • Department of Clinical Genetics, Erasmus Medical Center, Rotterdam, The Netherlands; Department of Pediatrics, Erasmus Medical Center,

Rotterdam, The Netherlands; Center for Lysosomal and Metabolic Diseases, Erasmus Medical Center, Rotterdam, The Netherlands


# Part I

Overview

# Chapter 1

# Introduction and History of the Chemistry of Nucleic Acids Therapeutics

### Michael J. Gait and Sudhir Agrawal

#### Abstract

This introduction charts the history of the development of the major chemical modifications that have influenced the development of nucleic acids therapeutics focusing in particular on antisense oligonucleotide analogues carrying modifications in the backbone and sugar. Brief mention is made of siRNA development and other applications that have by and large utilized the same modifications. We also point out the pitfalls of the use of nucleic acids as drugs, such as their unwanted interactions with pattern recognition receptors, which can be mitigated by chemical modification or used as immunotherapeutic agents.

Key words Antisense, siRNA, Nucleic acid therapeutics, Oligonucleotides, Toll-like receptors, Pattern recognition receptors, Gapmer, Splice switching

#### 1 Introduction to Synthetic Antisense Oligonucleotides and siRNA

Oligonucleotides are short single-stranded sections of DNA or RNA that contain 2<sup>0</sup> -deoxyribo-nucleosides or ribo-nucleosides, respectively, which are linked by 3<sup>0</sup> –5<sup>0</sup> phosphodiester linkages (Fig. 1a). Antisense oligonucleotides are those that are complementary to a section of naturally occurring RNA, such as an mRNA or a viral RNA, to form Watson–Crick base pairs and to thus inhibit a biological function of that RNA. Zamecnik and Stephenson pioneered this concept in 1978 by utilizing antisense oligodeoxyribonucleotides (ODNs) to bind and inhibit the replication of Rous sarcoma virus (RSV) RNA [1]. This work followed much earlier (1969) studies of De Clercq et al. on interferon induction by synthetic polynucleotides and their phosphorothioate analogues [2] and together these early studies heralded the new field of nucleic acids therapeutics that began to accelerate in the mid to late 1980s.

Many further chemistry developments since then in the use of synthetic oligonucleotide analogues, as outlined below, as well as advances in molecular biology, such as in the newer fields of short

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_1, © The Editor(s) (if applicable) and The Author(s) 2022

Fig. 1 Chemical structures. (a) DNA oligonucleotides (ODN), (b) phosphorothioate ODN (PS-ODN), (c) <sup>R</sup>p-PS-ODN, (d) <sup>S</sup>p-PS-ODN, (e) methylphosphonate ODN (PM-ODN), (f) phosphoramidate ODN (PN-ODN) R <sup>¼</sup> H or alkyl, (g) phosphomorpholidate, (h) phosphopiperazidate, (i) phosphorodiamidate morpholino (PMO), (j) peptide nucleic acids (PNA). B <sup>¼</sup> heterocyclic base

interfering RNA (siRNA) and non-coding RNAs, such as micro-RNA (miRNA), have led to the widespread and convenient use of synthetic oligonucleotides as antisense and siRNA reagents for gene ablation or targeting of non-coding RNA, as well as their use in animals and in humans leading to the approval of 12 drugs to date. In this chapter we outline the history of oligonucleotide chemistry in antisense and siRNA that has led to preclinical studies that have guided their development with drug-like properties and hence clinical trials (Fig. 2). We go on to discuss the development of the principles of widely used antisense gapmers and siRNAs as well as their immune responses by triggering pattern recognition receptors (PRRs) and how such activities can be controlled or harnessed for

Fig. 2 Evolution of chemical modifications during the development of antisense as therapeutic agents. In the nineties, various modifications of ODNs and ORNs were studied and based on RNase H activation, PS-ODNs became the choice as first-generation antisense agents. Soon it was realized that PS-ODN had off-target activities including complement activation and also sequence specific immune activation. This led to questions on the mechanism of action of PS-ODNs and associated safety signals, and clinical development of most of the PS-ODN ASOs was discontinued. In parallel use of modified ORN for splicing correction in the cells was established. Based on the early work conducted in early nineties, gapmer antisense design provided the key attributes and became the choice as second-generation antisense agents. Studies of chemical modifications in antisense has facilitated development of other therapeutic oligonucleotides. The key modifications which have been identified in the development of antisense, e.g. PS-PDN and PS-ORN, 20 -modified or 2<sup>0</sup> -O-substituted ribo-nucleosides, bridged ribo-nucleosides, and PMO which are being employed in various nucleic acid therapeutics. In the last few years, a number of drugs based on gapmer ONs (mipomersen, inotersen, volanesorsen), 2<sup>0</sup> -MOE PS-ORN (nusinersen), PMO (eteplirsen, golodirsen, vitolarsen), and siRNA (patisiran, givosiran) have been approved

> use as drugs (Fig. 3). Readers are also referred to a recent book edited by us on recent Advances in Therapeutic Nucleic Acids [3].

#### 2 Oligonucleotide Synthesis

Early work in the 1960s and 1970s on phosphodiester and phosphotriester chemistry for the synthesis of ODNs required armies of nucleic acids chemists for painstaking oligonucleotide synthesis in solution phase that took months to years for each synthesis (for

Fig. 3 Pattern recognition receptors (PRRs). The number of PRRs listed in the right-side column which is known to recognize patterns of nucleic acids and induce appropriate immune responses. This recognition is applicable to all use of nucleic acids for therapeutics, and depending on the engagement could affect the mechanism of action and also safety signals. This recognition could be due to PAMPs of the nucleic acid agent being used or due to DAMPs, due to the buildup of administered agent in the tissues and organs, being recognized as endogenous nucleic acids

example [4]). The revolution for molecular biologists came in the late 1970s and early 1980s with the development of solid-phase ODN synthesis first suggested by Letsinger and Mahadevan in 1965 [5] and later developed into working methods in the laboratories of Gait [6–8] and Itakura [9]. These methods were then superseded by the more efficient phosphoramidite chemistry of Caruthers and colleagues [10], which went on to be automated, such as by Applied Biosystems and other companies. The rapid and automated synthesis allowed molecular biologists to obtain synthetic ODNs readily for biological purposes such as for sequencing, cloning, and gene synthesis. The history of oligonucleotide synthesis chemistry has been reviewed [11, 12]. Today standard and modified ODNs can be obtained rapidly and efficiently on a small to large scale through highly automated solid-phase DNA oligonucleotide synthesis for many biological and diagnostic purposes.

Oligoribonucleotide synthesis is also now well established for the synthesis of siRNA or for aptameric RNAs. Currently, several oligonucleotide manufacturing plants are operational to produce oligonucleotides under GMP conditions.

#### 3 Synthetic Oligonucleotide Analogues in Antisense and siRNA

The early work of Zamecnik and colleagues utilized RSV since this was the only viral RNA sequenced at this time. Zamecnik noticed that both ends of the linear RNA genome bore the same primary sequence in the same polarity and that DNA might be synthesized from the RNA by reverse transcription via circularization of the 50 -end with the 30 -end through base pairing. Zamecnik and Stevenson were able to block this circularization by use of a synthetic ODN via hybridization with the 30 -end of the viral RNA. By use of a cell-free system, translation of the RSV mRNA was impaired, thus leading to inhibition of viral replication. This work was the birth of the antisense concept [1, 13].

Further progress in the antisense field awaited the mid to late 1980s for the availability of genomic DNA (or RNA) sequences for antisense targeting as well as the new automated methods of oligonucleotide synthesis as described above. There was also some scepticism regarding the stability and eukaryotic cell entry ability of unmodified oligonucleotides. Nevertheless Zamecnik and Gallo were able to employ unmodified antisense ODNs to inhibit human immunodeficiency virus 1 (HIV-1) replication [13] and to suppress expression of HIV-1 related markers [14]. Cellular uptake of the ODN was not a limiting factor since experiments were carried out in primary human cells and non-targeted control oligonucleotides showed minimal inhibition of HIV-1 replication, thus demonstrating sequence-specificity. This paper reignited the therapeutic potential of the antisense approach.

The next step was to provide drug-like properties to the unmodified antisense ODNs through chemical modifications. In this context the key consideration was to provide nucleolytic stability to antisense ODNs without affecting their hybridization and affinities with the RNA target. Not much was known at the time about the in vivo characteristics of unmodified ODN, or any modified ODNs, which could have guided the study of potential modifications. The first step was to see if modifications of the internucleotide linkages would provide nucleolytic stability to ODNs, while preserving the hybridization affinity to the target RNA. Later on, discovery of PRRs further provided insights into recognition of pathogen associated molecular patterns (PAMPs) of nucleic acids, and how sequence of antisense and nucleic acid-based therapeutic approaches could affect the mechanism of action (Fig. 3).

3.1 Backbone Analogues 3.1.1 Phosphorothioates The antisense field took inspiration from the very early work of De Clerq, Eckstein, and Merigan [2] where phosphorothioate (PS) modifications were studied in homopolynucleotides as stabilizing agents. By the mid-1980s new chemical synthesis methods for the PS linkage in ODNs became available [15]. Here, a simple sulfur atom replaces a non-bridging oxygen atom (Fig. 1b). However, standard automated synthesis, which in the phosphoramidite method involves treatment with a sulfurizing agent in place of oxidation by iodine, produced a mixture of diastereomeric oligonucleotide products (Rp and Sp) (Fig. 1c, d) and thus there was found to be a lower binding affinity to target RNA compared to phosphodiesters. However, PS-linked ODNs are much more resistant to nuclease degradation than phosphodiesters. Optimization of the synthesis methodology allowed the synthesis of milligram quantities of PS-ODNs for use in cell and in vivo experiments.

Early studies showed dose-dependent inhibition of viral replication and antiviral activity in HIV-1 infected cells by use of antisense PS-ODNs targeted to several regions of HIV-1 mRNA [16, 17]. Surprisingly, homopolymers were also effective and antiviral activity depended on the base composition, suggesting that PS-ODNs also had off-target activity. Longer PS-ODNs were more effective than shorter ones and cellular uptake was efficient in primary human cells without a carrier. In addition, antisense PS-ODNs showed potent and durable inhibition of HIV-1 replication in chronically HIV-1 infected cells [18–20]. Soon after, studies with PS-ODN targeted to influenza virus showed inhibition of virus replication [21]. Based on these early studies and promise, PS-ODNs became the choice for first-generation antisense agents.

Following these studies, work on antisense was pursued in many laboratories around the world for a broad range of applications [22–24]. In addition, several new companies were founded to advance therapeutic applications of antisense, such as Gilead Sciences, Isis Pharmaceuticals (now Ionis Pharmaceuticals), Hybridon (now Idera Pharmaceuticals), and others. Numerous reports appeared on the use of antisense PS-ODNs to target viruses [25, 26], oncogenes [24, 27], and kinases [27, 28], etc. Cellular uptake of PS-ODNs in transformed cells in culture was found to be poor but could be improved substantially by use of cationic lipid carriers. It was also shown that an antisense PS-ODN bound to its target RNA engaged RNase H [19, 20] to excise the RNA strand and this was therefore likely to be the mode of action and not steric block inhibition of translation as observed with other modified ODNs [29, 30]. Nevertheless RNase H cleavage activity was poorer than for a PO-ASO [20].

The first in vivo study in mice of a systemically delivered PS-ODN showed that plasma half-life was very short but that there was a broad tissue disposition with most delivered to liver and kidneys and the lowest amounts in the brain [31]. The PS-ODN was stable in tissues for several days and excreted primarily in urine in degraded form mostly through exonuclease cleavage from the 30 -end. Chemical modifications at the 30 -end increased the stability [32, 33]. The PS-ODN was bound by serum proteins, which increased the plasma half-life and improved tissue disposition [34, 35].

Antisense PS-ODNs showed very potent activity in animal models of viral diseases and cancer [36, 37]. However, in some cases a control PS-ODN also showed some activity, leading to the possibility of off-target effects [38]. For example, a PS-ODN targeted to human papillomavirus inhibited papillomavirus-induced growth of implanted human foreskin in a mouse xenograft model but unexpectedly was also active in a mouse cytomegalovirus (CMV) model [39]. Studies in immune-compromised mice showed that the effect of the PS-ODN was largely due to immune activation of the host. Sequence-dependent immune stimulation was confirmed during non-clinical safety evaluations of drug candidates. Repeated systemic administration of PS-ODN candidates in mice and rats caused inflammation, splenomegaly, and histological changes in multiple organs [40, 41]. Further in non-human primates, bolus administration of the first antisense PS-ODN (GEM91) led to severe hemodynamic changes due to activation of the alternative complement pathway [42]. Stimulation of the alternative pathway complement activation cascade became the first documented off-target effect, which was due to a plasma concentration effect of the poly-anionic nature of PS-ODNs. It could be mitigated by subcutaneous administration or by slow intravenous infusion. Thereafter, the US Federal Drug Agency (FDA) published guidelines and required the use of non-human primates for non-clinical safety studies of all oligonucleotide drugs [43].

In the 1990s a number of antisense PS-ODN clinical candidates were advanced to human trials [44] through intravenous infusion, intravitreal or subcutaneous delivery [45–47]. Humans showed similar pharmacokinetics and excretion data to those of non-human primates [48, 49]. However, most clinical studies were discontinued due to the lack of activity or a poor therapeutic index [50]. For example, the subcutaneous administration of GEM91 in humans caused flu-like symptoms, swelling of the draining lymph nodes, prolongation of activated partial thromboplastin time (aPTT), and thrombocytopenia [29]. Rather than suppression of HIV-1, HIV-1 RNA levels were increased in blood [51]. However, intravenous delivery had minimal effect on these parameters. There were a few reports of immunostimulatory properties of DNA/ODNs containing CG nucleotides [52, 53]. It only became clear much later that PS-ODNs containing an unmethylated CpG motif activated the immune responses by binding to Toll-like receptor 9 (TLR9), an innate immune receptor present in immune cells that recognizes DNA containing CpG dinucleotide motifs [54]. It became clear that the flu-like symptoms and injection site reactions seen with most of the PS-ODN drug candidates in clinical trials, such as the clinically approved drug fomivirsen, administered intra-vitreally to treat AIDS-related CMV-induced retinitis, contained a CpG motif [55]. Thus, the true mechanism of action of this first-generation drug, now no longer marketed, remains unclear. Altogether, preclinical and clinical studies have provided important insights into the properties of PS-ODNs and its use as drugs [35, 56, 57].

Also of debate for some time has been whether the presence of a mixture of Rp and Sp diastereoisomers in the synthetic PS-ODNs (Fig. 1c, d) bears any influence on their biological properties. For example, a 20-mer would have 2<sup>19</sup> isomers. The stereospecificity of enzymes that act on nucleoside phosphates was well known from early work of Eckstein (reviewed in [58]). Since PS-ODN interacting enzymes, such as nucleases, also utilize only a single diasterioisomeric isomer [59], it was plausible that there might be a significant biological effect in cells of utilizing mixed PS diastereomers in antisense PS-ODNs. Testing of this only awaited the solidphase synthesis of stereo-enriched and stereo-pure PS-ODNs. This became possible through pioneering work of Stec and later by use of nucleoside bicyclic oxazaphospholidinium synthons [15, 60]. It is now known that binding strength and recognition by RNase H is generally higher for antisense oligonucleotides containing Rp linkages but depends crucially on the placement of these with respect to Sp linkages and overall stereospecific PS-ODNs have had limited therapeutic utility [61]. Recently certain stereo-pure antisense oligonucleotides were shown to have improved activity in cell culture and in vivo [62] but the therapeutic significance of such stereospecificity is currently hotly disputed [63]. Even more recently, the clinical development of a stereo-pure PS-ODN, WVEN-531 targeted to DMD has been discontinued due to lack of clinical activity [64]. Furthermore, dosing of this antisense ODN also led to transient increases in complement factors and C-reactive protein [65].

3.1.2 Charge-Neutral Analogues Two phosphate-containing, charge-neutral oligonucleotide analogues that were particularly used in early antisense studies are the methylphosphonate [66] (Fig. 1e) and the phosphoramidate linkages [67] (Fig. 1f). They both consist of a mixture of diastereoisomers. Methylphosphonate ODNs (MP-ODNs) are stable at physiological pH and are resistant to nucleolytic degradation but are less strongly bound to target RNA compared to PO-ODNs [66, 68]. MP-ODNs targeted to HIV-1 showed dose-dependent inhibition of HIV-1 replication [16], but they are less active than PS-ODNs due to their lack of RNase H activation [19] but instead inhibit protein translation, which is generally a weaker activity in cells. Limited in vivo studies with a MP-ODN showed that while this modification is very resistant to nucleolytic degradation, due to poor protein binding, there was a very poor in vivo disposition and the majority of the administered ODN was eliminated in urine rapidly (Agrawal, unpublished data). In addition, longer MP-ODNs, which bind more strongly to RNA and which are therefore more potent, are poorly soluble under physiological conditions and thus have not been advanced toward clinical trials. By contrast, a 13-mer antisense oligonucleotide containing all phosphoramidate linkages is more soluble. An anticancer agent (GRN163) inhibits the enzyme telomerase [67] and did get into a clinical trial, however, clinical development was discontinued due to lack of clinical activity. In early work, phosphoramidate-linked antisense oligonucleotides (Fig. 1g, h) targeted to HIV-1 showed similar results in cell-based assays to an MP-ODN and were not pursued [16].

Phosphorodiamidate morpholino oligonucleotides (PMOs) are also charge-neutral but here a morpholino ring replaces the sugar unit (Fig. 1i) [69]. PMOs inhibit translation by a steric block mechanism [70] as they are not recognized by RNase H. They are completely resistant to nucleases but are not taken up well by cells and thus require very high doses for in vivo delivery. They were found to be strong antiviral agents, for example, against Ebola, Marburg, and Chikungunya viruses [71]. Three exon skipping PMO drugs, eteplirsen, golodirsen, and vitolarsen designed to induce alternative splicing and restore the reading frame of mutant dystrophin in patients with Duchenne muscular dystrophy (DMD) [72] have been approved but requires the use of high doses (50 mg/kg or higher). Its therapeutic effectiveness, based on biomarkers, is limited [73], but it is a safe drug at the therapeutic dose. PMO and other chemistries used in exon skipping and other steric block activities have been reviewed [74].

Another initially highly promising, charge-neutral analogue are peptide nucleic acids (PNA), where the sugar-phosphate backbone is replaced by aminoethylglycine units linked by amide bonds (Fig. 1j) [75]. PNA binds strongly to target RNA and, like PMO, they are also completely resistant to degradation by nucleases as well as proteases. Also similar to PMO, duplexes with RNA are not recognized by RNase H and thus PNA acts by a steric block mechanism. Antisense PNAs have been broadly studied as anticancer [76, 77], antiviral [78, 79], and antibacterial agents [80, 81] as well as inhibitors of micro-RNAs [82]. However, once again very high doses are needed in in vivo applications, due to poor cellular uptake and unfavorable pharmacokinetics. Poor in vivo biodistribution is a likely reason for why antisense PNAs have not to date found utility as clinical candidates.


Fig. 4 Chemical structures of the ribo-nucleoside units of therapeutically useful RNA and RNA analogues. (a) ribo-nucleoside (ORN) (b) 2<sup>0</sup> -O-methyl (2<sup>0</sup> -OMe), (c) 2<sup>0</sup> -O-methoxyethyl (2<sup>0</sup> -MOE), (d) bridged/locked nucleic acid (LNA), (e) 2<sup>0</sup> -O,4<sup>0</sup> -C-ethylene linked nucleic acid (ENA), (f) tricyclo-DNA (tcDNA), and (g) constrained ethyl (cET). B <sup>¼</sup> heterocyclic bases

solid-phase synthesis became available commercially in the early 1990s [29]. Studies with 2<sup>0</sup> -O-methyloligoribonucleotide phosphorothioates (2<sup>0</sup> -OMe PS-ORN) showed enhanced stability to nucleases as compared to PS-ORN and showed a higher affinity to target [84]. However, they also showed lower antisense activity compared to PS-ODNs, demonstrating that activation of RNase H was key for this activity [30, 51, 85, 86]. Since then, very many additional 2<sup>0</sup> -O-alkyl analogues have been synthesized and tested in antisense oligonucleotides (ONs), predominantly in gapmer studies (see below) to allow recognition by RNase H. From these studies 2<sup>0</sup> -O-methoxyethylribonucleoside (2<sup>0</sup> -MOE) (Fig. 4c) has been employed widely in clinical gapmer candidates (Chapters 3 and 4 of Agrawal and Gait [3]).

As described earlier, dose-dependent activation of complement and prolongation of aPTT were found to be unwanted side effects of PS-ODNs. These effects as well as strong binding to serum proteins were thought to be due to the poly-anionic nature of the PS linkage. However, there were found to be significantly less side effects when PS-ORN or 20 -OMe-PS-ASO where used, suggesting that the poly-anionic nature of the PS backbone in PS-ORN and PS 20 -OMe is not as pronounced when placed in the context of an RNA or RNA-like sugar conformation [40, 85–87]. This became crucial to their use in later gapmer antisense studies.

Uniformly 20 -O-alkyl modified PS-RNA has also found very high therapeutic use in splice switching (exon skipping or exon inclusion) and other steric blocking applications due to their high binding strength to nuclear pre-mRNA [88, 89]. However, the exon skipping 20 -OMe PS-ORN antisense drisapersen drug candidate failed to show clinical benefit in patients with DMD and also caused significant adverse side effects and was, therefore, not approved for clinical use [90]. By contrast, the 18-mer 20 -MOE phosphorothioate (20 -MOE PS-ORN, nusinersen), which redirects the splicing of the SMN-2 gene to generate active SMN protein (exon inclusion), administered intrathecally only few times a year was approved by the FDA for the treatment of spinal muscular atrophy (SMA) [91, 92]. The thrombocytopenia and renal toxicity observed with the use of drisapersen could be largely due to the need for repeated subcutaneous dosages of 20 -OMe PS-ORN, which is very stable to nucleolytic degradation and therefore may accumulate in tissues due to its long half-life and potentially interact with PRRs. Nusinersen, a 2<sup>0</sup> -MOE PS-ORN, is also quite stable to nucleolytic degradation, but its intrathecal administration and need for infrequent and lower doses minimizes the impact of tissue accumulation and avoids the need for passage into the brain and spinal cord from the circulation.

3.2.2 Locked/Bridged Nucleic Acids A major step forward in the design of antisense ONs was the development in the laboratories of Wengel and also of Imanishi of bicyclic sugar analogues known as locked or bridged nucleic acids (LNA/BNA). Here the conformational flexibility of nucleotides is significantly reduced by linkage of the 2<sup>0</sup> -oxygen atom to the 4- 0 -carbon atom in the ribose ring (Fig. 4d). This results in a significant increase in the binding affinity of ONs to complementary RNA targets with an increase in the melting temperature of 2–8 -C per residue [93]. Unfortunately, LNA oligomers of 8 units or longer tend to self-aggregate. Therefore they became more useful as mixmers with 2<sup>0</sup> -deoxynucleotides and here miravirsen, the first microRNA-targeting drug, which acts by sterically blocking microRNA-122, highly expressed in liver, was developed for the treatment of hepatitis C virus infection, a debilitating liver disease [94]. Unfortunately, this drug's clinical development was discontinued because of safety issues. LNA has also been used as mixmers with 20 -OMe nucleotides targeting various RNAs in cells (e.g. [93]) and has also found utility in the flanking sequences of gapmers. This had the effect also of modulating the binding strength of the ON and increasing the specificity of the interaction (reviewed in Chapter 3 of Agrawal and Gait [3]).

Another bicyclic analogue that became useful is 20 -O,4<sup>0</sup> -Cethylene linked nucleic acid (ENA) [95] (Fig. 4e). In a recent study an antisense ON DS-5141, containing segments of 20 -OMe PS-RNA and ENA, showed good activity in an mdx mouse model of DMD and a phase 1/2 clinical trial was carried out in Japan [96]. A further analogue useful in steric block applications is tricyclo-DNA (tcDNA) [97] (Fig. 4f). However, perhaps the most important bicyclic derivative of LNA that has found considerable therapeutic utility is the methylated analogue known as "constrained Ethyl" (cET), which is being employed in shorter gapmers [98] (Fig. 4g) and being evaluated in preclinical and clinical studies by Ionis Pharmaceuticals Inc. All these types of bridged nucleic acids have shown very strong affinity to target RNA and increased nucleolytic stability, but none of them are substrates for RNase H. Thus, these types of bicyclic sugar analogues are mostly used in steric block/splicing modulation approaches and in the flanking sequences of gapmers.

3.3 Heterocyclic Base Analogues In early antisense studies it was thought that increased antisense activity might be achievable by improving the affinity of an ODN to target RNA through modification of the heterocyclic bases, for example, by adding an extra hydrogen bond in the base pairing between an ODN and its RNA target or by increasing the base stacking potential in a DNA–RNA duplex. Chemically this was simplest through modifications in the pyrimidine rings, for example, by modifications at positions C-2, C-4, C-5 or at C-6, and many of these base analogues were incorporated into antisense ODNs. However, few of these proved to be of significant value. Incorporation of modified purines generally resulted in a reduced binding affinity of an antisense ODN. Perhaps the most useful study of antisense activity was of incorporation of various heterocyclic bases in ODNs including the increased base stacking analogues C-5 propynyl and 5-methyl cytosine (5-MeC) and the increased hydrogen-bonding analogues phenoxazine, and G-clamp. These studies showed that the increased hydrogen-bonding analogue G-clamp had potent dose-dependent antisense activity [99]. Unfortunately, these antisense ODNs containing G-clamps were found to be highly toxic in in vivo studies. Currently, the only significantly used nucleoside base analogue in antisense ODNs is 5-methyl-20 -deoxycytidine (5-MedC) [100]. This methylated base analogue is used mainly to mitigate immune activation in CpG dinucleotide sequences rather than for changing binding strength [101].

#### 4 RNase H Active Gapmer Chemistry for Use as Drugs

Early studies conducted with various modified ODNs, ORNs, and 20 -substituted ORNs as antisense agents provided great insights into what is important for providing drug-like properties to antisense oligonucleotides [3]. Studies with PS-ODNs showed that increased nucleolytic stability and activation of RNase H were key. However, polyanion-related side effects and sequence-dependent immune activation were limiting factors in their broad applicability [50, 57]. Studies with MP-ODNs showed that polyanion-related side effects could be completely mitigated (Agrawal, unpublished data), and had significant nucleolytic stability. However, with lower affinity and lack of RNase H activation, there was a loss in antisense potency [16]. These observations led to the concept of combining desirable properties of the two modified ODNs to provide druglike properties to antisense oligonucleotides [29]. The first studies were carried out with antisense containing segments of PS-ODN and PM-ODN or PN-ODN referred to as mixed backbone antisense ONs. These antisense designs showed increased nucleolytic stability and RNase H activation [19, 40]. However, reduced affinity limited their potency. Further insight was obtained from in vivo studies in which a mixed backbone ON containing PS-ODN and PM-ODN showed wide tissue disposition and increased stability and longer half-life in tissues [102].

This led Agrawal and colleagues to design antisense oligonucleotides in which the segments of PS-ODN and 2<sup>0</sup> -substituted PS-ORN were combined at the appropriate positions [29, 84, 85]. These types of antisense oligonucleotides were referred to as Hybrid ONs, now commonly referred to as gapmers (Fig. 5). In the original design of antisense, a segment of PS-DNA was placed in the middle and segments of 2<sup>0</sup> -O-alkyl PS-ORN or a combination of PO- and PS-linkages were placed at both 3<sup>0</sup> - and 5<sup>0</sup> -ends [29, 84, 85, 87, 103]. This design of antisense combined the desirable properties of PS-DNA and 2<sup>0</sup> -O-alkyl PS-ORN, and provided increased affinity to targeted RNA, activation of RNase H, increased nucleolytic stability, and reduced polyanion-related side effects. Furthermore, inflammatory responses were also reduced [35, 40, 56]. In vivo administration in mice showed similar plasma half-life and tissue disposition similar to that observed with PS-ODNs, and with increased in vivo stability and retention in tissues [104]. Also, due to increased in vivo stability, oral and rectal delivery of gapmer antisense was possible [34]. It was postulated that the increased stability and in vivo persistence may allow less frequent dosing to obtain therapeutic benefits.

There was also a concern that increased retention of gapmer antisense in tissues may lead to tissue build up following repeated dosing, which would induce local inflammatory responses and side effects, thereby limiting its therapeutic potential.

Other configurations of gapmer antisense were also evaluated, including the configuration in which a segment of 20 -O-alkyl PS-ORN was placed in the center and segments of PS-DNA were placed at both 30 - and 50 -ends. This design of gapmer antisense showed increased potency compared to PS-DNA and reduced polyanion-related side effects. In general, the specificity of RNase H mediated cleavage and its efficiency and excision sites were dependent on the position of the PS-DNA in gapmer antisense ONs [105].

Based on these encouraging results, gapmer antisense became the choice for second-generation antisense agents. In 2001, a

Fig. 5 Design of gapmer antisense oligonucleotides. In a gapmer antisense, segments of PS-DNA and modified RNA are appropriately placed to combine desirable characteristics for antisense agent with both of these modifications. PS-DNA segment provides RNase H activation, and modified RNA segments provide increased nucleolytic stability, affinity to target RNA, decreased polyanionic characteristics and inflammatory responses

licensing agreement between the companies allowed the technology to be widely available [44]. Over the years, studies have been carried out to establish the optimal size of the window of a central PS-DNA segment [105]. Similarly, studies have been carried out to optimize the size of the modified ORN wings at both 3<sup>0</sup> - and 50 -ends. In the wings of the gapmer antisense, various modified ORNs have been incorporated and evaluated (see Chapter 3 of Agrawal and Gait [3]). To date, most promising results have been obtained with gapmer antisense containing segments of 2<sup>0</sup> -Omethyl or 2<sup>0</sup> -O-methoxyethyl at both 3<sup>0</sup> - and 5<sup>0</sup> -ends. Over 30 gapmer antisense drug candidates containing 2<sup>0</sup> -O-methoxyethyl or LNA segments have been advanced to clinical evaluations following systemic delivery. To date, three candidates have been approved for clinical use. These include inotersen [106], volanesorsen [107], and mipomersen [108]. Clinical development of several gapmer antisense drug candidates have been discontinued, due to lack of clinical activity and or safety signals. These include ISIS-FXIRx, ISIS-EIF4ERx, ATL1103, ATL1102, ISIS-GCGRRx, ISIS-PTBRx, ISIS-APOARx, ISIS-SOD1Rx, ISIS-FGFR4Rx, ISIS-405879, OGX-011, OGX-427, LY2181308, ATL1103, ATL1102, etc.

As discussed above, many of the bicyclic sugar analogues including locked/bridged nucleic acids have been studied as antisense agents. These analogues have also been studied as part of the wings in gapmer antisense. These include LNA (Fig. 2d) [109], constrained ethyl 20 -40 bridged nucleic acid (cEt) (Fig. 2g) [110], anhydrohexitol [91], fluorocyclohexenyl (F-CeNA) [111], altritol nucleic acids [112], and tricyclo-DNA (tcDNA) (Fig. 2f) [97]. These modifications provide very high affinity and have allowed the length of the gapmer antisense to be reduced. A few of these shorter gapmer ASOs are being employed to achieve allele specific knockdown [113]. While the use of shorter antisense may be cost effective, it increases the possibility of off-target effects by binding to non-targeted RNA [114, 115]. Also selected LNA and cET ASOs have been associated with liver toxicity [116, 117].

Several gapmer antisense drug candidates employing LNA in the wings have advanced toward the clinic but development of most of these candidates have been discontinued, primarily due to safety issues and lack of therapeutic index.

Other modifications in the gapmer antisense studies include 20 -deoxy-2<sup>0</sup> -fluoro-beta-D-arabinonucleic acid (FANA) [118], 3<sup>0</sup> fluorohexitolnucleic acid (FHNA) [119], 2<sup>0</sup> -thiothymine, 5-modified pyrimidine bases, etc. These studies are limited to preclinical evaluations.

#### 5 siRNA Chemistry for Use as Drugs

The lessons learned in the development of antisense ONs have allowed the development of siRNA therapeutics to be speeded up. siRNAs have a well-defined structure: a short double stranded RNA of 20–25 base pairs with phosphorylated 5<sup>0</sup> -ends and hydroxylated 3<sup>0</sup> -ends and also usually containing two 3<sup>0</sup> -overhanging nucleotides, although blunt ends are sometimes used. While the key requirement is to provide nucleolytic stability to a siRNA candidate, it requires an understanding of the function of each strand. One strand is called the passenger strand and the other is the active component and is called the antisense or guide strand. It is the guide strand that is incorporated into the enzyme complex called RISC in order to be directed to cleave the target RNA strand, while the passenger strand is displaced.

Studies of various chemical modifications in antisense and their impact on providing drug-like properties have allowed the use of some of these modifications in development siRNA therapeutics. These include PS-linkages, various modified ribose sugars such as 20 -O-methyl, 2<sup>0</sup> -fluoro 20 -deoxy (20 -F), LNA as well as the sugar ring-opened analogues unlocked nucleic acid (UNA), and glycol nucleic acid (GNA). In siRNA chemical modifications are introduced strategically to provide nucleolytic stability. In addition the passenger strand is usually heavily modified in order to block passenger strand entry into RISC, while to promote RISC loading of the guide strand only light modification is used, such as 20 -F replacement of 20 -OH groups in pyrimidines. At the same time modifications must not be placed centrally in the guide strand so as to block RISC-associated cleavage of the target RNA. The exact locations of such modifications in guide and passenger strands are generally closely guarded secrets by reagent suppliers. In addition, the 50 -phosphate of a siRNA guide strand is essential for recognition by RISC. Phosphatase-resistant analogues of the 50 -end phosphate have been shown to improve the in vivo efficacy and are used in clinical candidates [120].

In siRNA candidates, chemical modifications provide nucleolytic stability, however, delivery to a desired tissue or cell type requires use of carrier or conjugation with delivery moieties [121]. To date, two main delivery platforms—ionisable lipid nanoparticles (iLNPs) and trivalent N-acetylgalactosamine (GalNAc) conjugates—have been employed for delivery to liver hepatocytes. To date, two siRNA drugs have been approved for clinical use patisiran, which uses lipid delivery, and givosiran, which uses a GalNAc conjugation. Third siRNA drug candidate, inclisiran, a GalNac conjugate has shown positive results in phase 3 trial [122]. Details of these structure activity relationship studies have been discussed in two chapters from a previous book [123, 124].

#### 6 Immune Responses to Nucleic Acids

Over the last five decades there have been several reports on observations that certain nucleic acid sequences showed immune stimulatory properties [52, 125, 126]. In the mid-1990s subcutaneous administration of an antisense PS-ODN targeted to HIV-1 (GEM91) in HIV-1 infected individuals caused flu-like symptoms and systemic immune responses [51]. This observation alone could not be explained until the discovery of PRRs. These receptors are part of the immune system and PAMPs and host-derived damageassociated molecular patterns (DAMPs). These PRRs play an essential role in establishing antiviral and antibacterial responses by recognizing PAMPs. However, PRRs could also induce development of autoimmune and inflammatory diseases by recognizing DAMPs [127, 128].

PAMPs are highly conserved motifs in pathogens, such as bacteria and viruses. There are several PRRs that are known to recognize motifs, sequences, and patterns of nucleic acids and induce receptor-mediated immune responses. These include members of Toll-like receptors (TLRs), of which four TLRs respond to nucleic acids. TLR3, TLR7/8, and TLR9 recognize doublestranded RNA (dsRNA), single-stranded RNA with certain sequence composition and modified bases (ssRNA), and DNA containing unmethylated CpG sequences (CpG DNA), respectively (Chapter 13 of Agrawal and Gait [3]; [129]). These TLRs are localized in endosomes and expressed on various cell types. The type of immune response induced varies dependent on the receptor and the nature of the nucleic acid [130]. In addition to TLRs, additional receptors are present in the cytoplasm known to recognize nucleic acid-based PAMPs. These include retinoic acidinducible gene-I (RIG-I), melanoma-associated gene-5 (MDA-5), absent in melanoma 2 (AIM2), cyclic-AMP synthase (cGAS), and stimulator of the interferon gene (STING) (Chapter 13 of Agrawal and Gait [3]).

Following the discovery of TLR9, it became clear that immune activation observed with administration of GEM91 was due to the presence of unmethylated CpG dinucleotides in the antisense sequence [131]. This also provided insights into many of the preclinical studies that the chosen antisense may be exerting antiviral or anticancer activity due to immune activation and not by an antisense mechanism [51, 57]. Interestingly, most of the antisense PS-ODNs in clinical development contained unmethylated CpG motifs, raising questions on the intended mechanism of action [51, 57]. Clinical development of all these antisense drug candidates was discontinued due to lack of activity but also due to safety signals. Similar observations have been made with a few initial siRNA candidates and once again mechanisms of action have been correlated with activation of immune responses [132, 133].

The discovery of PRRs has provided key insights into many of the observations made with use of PS-ODN antisense. For example, TLR9 is a receptor for synthetic ODNs containing unmethylated CpG motifs [131]. Activation of TLR9 leads to induction of Th1 type immune responses in mice, primates, and in humans (Chapter 14 of Agrawal and Gait [3]). Inductions of Th1 type immune responses, which include type interferon (IFN) and interleukin 12 (IL-12), have shown therapeutic potential as antiviral and anticancer agents. This explains the activity of a PS-ODN antisense containing the CpG motif targeted to HPV, also showing activity for CMV and loss of activity in immune compromised mice [39]. This also explains the reason for flu-like symptoms with administration of GEM91, a PS-ODN antisense containing the CpG motif target to HIV-1 [55]. Interestingly, most of the PS-ODN antisense that were advanced to clinical development in the early 1990s contained unmethylated CpG motifs [51]. Thus, their mechanisms of action could be largely due to immune activation or side effects were caused by immune activation.

Detailed structure activity relationships have been carried to elucidate the interaction of PS-ODNs with TLR9. These studies have provided great insights. For example, (a) the presence of unmethylated CpG motif is required, although its position in the sequence is equally important [134], (b) accessibility of the 50 -end is required [111, 135], (c) modifications of the flanking sequence on the 50 -end impacts the immune activity [136], (d) methylation of C in the CpG motif neutralizes immune activation and causes it to act as an antagonist, and (e) certain modified bases could be used in the CpG motif without inducing immune responses. These insights have been very helpful in designing antisense candidates. These lessons have provided the basis for the creation of optimized agonists and antagonists of TLR9. These classes of compounds have been studied extensively in preclinical models of cancer [106], vaccines [137], viral infection [138], and autoimmune diseases [139], and clinical proof of concept has been established in multiple diseases [3]; Chapters 5 and 14 of Agrawal and Gait [3].

Detailed structure–activity relationship studies have been carried out for TLR3 [134], TLR7 and TLR8 [127, 140], RIG-I [141], and AIM2 [142]. It is important to take these insights into consideration when selecting a sequence and prioritizing chemical modification for use in therapeutic applications.

#### 7 Conjugates and Delivery

The in vivo efficacy of ONs is defined by plasma half-life, tissue uptake, nucleolytic stability, and elimination. Systemic administration of several gapmer ASOs has shown a similar profile, i.e., short plasma half-life, wide tissue dispositions, and the presence of intact ASO for longer durations [121]. Even though the delivered gapmer ASO is present in targeted tissues including liver, for sustained clinical activity weekly dosing has been employed. This suggests that the administered ASO is not present in the right cells or cell compartment. Further insights came from intrathecal delivery of the 2<sup>0</sup> -O-methoxyethyl PS-ASO nusinersen (Spinraza) to treat SMA. Patients are being treated with IT delivery, administered only four to five times a year. This suggests that in a local compartment, a delivered ASO exerts pharmacodynamic activity for a longer duration and thereby requires less frequent dosing. In recent studies, both preclinical and clinical, conjugation of gapmer ASO with a GalNac cluster has been shown to improve potency and frequency of treatment for liver targeted RNA/gene targets. Efforts are being made to improve delivery of ASOs to muscles to treat muscular disorders employing antibody conjugates [143, 144].

Peptide conjugation has been researched extensively in recent years in efforts to increase the delivery of oligonucleotides. Numerous cell-penetrating peptides (CPPs) have been developed, which are beyond the scope of this Introduction. Readers are referred to a book describing methods that use cell-penetrating peptides [145]. However, the only peptides that have reached clinical development are Arginine-rich CPPs. These are not suitable for conjugation with negatively charged oligonucleotides because of the tendency of such conjugates to aggregate due to charge–charge interactions between the positively charged peptide part and the negatively charged oligonucleotide part. Instead they have found clinical utility for use as conjugates with charge-neutral PMOs. Here the company AVI Biopharma (now called Sarepta) developed a series of Arginine-rich CPPs that were taken to toxicological testing in monkeys but were found to have renal toxicity at elevated doses leading to a poor therapeutic index [146]. Recently Sarepta has advocated use of an alternative and shorter Arg-rich peptide, which is (Arg)6-Gly, as a PMO conjugate as a treatment for the neuromuscular disease DMD. This gave rise to significant improvements in delivery of an attached PMO and increased exon skipping [147]. The peptide-PMO conjugate is currently in Phase 2 clinical trials. Similar Arg-rich peptides, eg ones known as Pip having a short internal hydrophobic domain, have given rise to increased exon skipping for an attached PMO in muscles as well as in heart in an mdx mouse model of DMD [148]. Pip peptides and similar derivatives are currently being evaluated as potential therapeutics for other neuromuscular diseases, for example, in myotonic dystrophy [149]. Once again, shorter Arg peptide derivatives as PMO conjugates are likely to be the future clinical candidates in neuromuscular and neurodegenerative diseases.

Delivery of siRNA has been facilitated by the use of lipid complexes or as conjugates with GalNAc, mainly to the liver. Use of lipids provides stability to an siRNA candidate by encapsulating them, along with preferential delivery to the liver. Several lipid encapsulated siRNA candidates have advanced to clinical development. For example, patisiran formulated with lipid nanoparticles (LNP) has received regulatory approval. It is important to note that lipid-nucleic acid mixtures form complexes that create virus-like particle structures and engage PRRs to induce immune responses. In the case of patisiran, subjects were pre-treated with steroids to mitigate inflammatory responses.

The application of the use of GalNAc to hepatocytes has been known for some time and was employed for oligonucleotide delivery more than two decades ago [150]. GalNAc is a ligand for the asialoglycoprotein receptor 9 (ASGPR), which is very abundant on the surface of hepatocytes [151]. Conjugation with GalNAc generally leads to preferential delivery to the liver. However, depending on the nature of modifications of ONs, delivery to other compartments including the kidney has been observed. An siRNA-GalNAc conjugate givosiran has been approved for clinical use, another GalNAc conjugate. Inclisiran has shown positive results in a phase 3 clinical trial (Chapter 11 of Agrawal and Gait [3]).

#### 8 Further Developments in Therapeutics

Over the years many other applications of nucleic acid-based therapeutics have been pursued. These include aptamers, CRISPR/ Cas9, and use of modified mRNA for protein overexpression. While the construct and sequence of DNA or RNA employed in these uses may differ, one common aspect is the need to provide drug like properties to the selected agent. Most of the lessons learned in the development of chemistry in antisense field have facilitated the development of these approaches. In aptamers, modified nucleosides, such as 20 -O-methyl, 2<sup>0</sup> -fluoro or 20 -amino, and modified internucleotide linkages such as PS-linkages or boranophosphate are regularly employed [152, 153]. In the case of mRNA therapy, considerations of use of chemical modifications are different than in other approaches. The 50 -cap and 30 -poly(A) tail are the key contributors to provide long half-life and for efficient translation. New capping agents such as 1,2-dithiodiphosphonate modified caps have been shown to improve RNA translation [154]. Several modified nucleosides including N<sup>1</sup> -methyl-pseudouridine and others have been useful in increasing efficiency of translation, and also mitigating immune stimulatory activity [155, 156]. The positional incorporation of modified bases in mRNA affects the secondary structure of the mRNA, which in turn influences its translation. Further stability to mRNA is provided by formulation with LNPs [157].

In studying CRISPR/Cas9-based therapeutic applications, several modifications are being evaluated. These include PS linkages and 2<sup>0</sup> -fluoro, LNA, c-Et [158], 2<sup>0</sup> -O-methyl [159], 2<sup>0</sup> ,4<sup>0</sup> -BNA (NC) [N-Me] [160], etc. These chemical modifications not only provide stability but also mitigate interactions with PRRs. CRISPRbased technologies have been described in a recent book [161].

#### 9 Summary

Nucleic acid-based therapies are now entering into their fifth decade (see Fig. 2 for a timeline of developments). Since the first report of the antisense principle in 1978 using unmodified ODNs, the technology has evolved, and drugs are now being approved. Based on the progress to date and the promise of the results, nucleic acid therapeutics are now being recognized as the third major drug discovery and development approach in addition to small molecules and protein/antibody approaches.

Nucleic acid therapeutic agents are built of A, C, G, T, and U nucleotides and connected through internucleotide bonds. Early work on chemical modifications to provide drug-like properties to antisense and lessons learned have been of tremendous value not only in creating antisense drugs but also in developing therapeutics using synthetic nucleic acids with other mechanisms of action (Fig. 4). Nucleic acid therapeutics could be broadly divided into two classes, the first in which an agent is created to target RNA or DNA and modulate its expression, and in the second an agent is created to bind to proteins or cellular factors. In both of these categories, agents could be recognized by PRRs thereby inducing immune responses, either unintended or intended affecting the mechanism of action.

The work on the chemistry of antisense has provided us with a few key modifications that have become important tools in nucleic acid therapeutics. The most important of these include PS linkages in ODN and ORN, gapmer design, selected 20 -O-sustituted nucleosides, and various bridged/locked nucleic acids, etc. The art of creating a nucleic acid agent lies in the understanding of putting together the nucleotide sequence and various modifications for its intended mechanism of action without interacting with PRR (Fig. 3).

#### Acknowledgments

SA is indebted to Mike Gait for his mentorship during his postdoctoral training in Mike's laboratory and over the last three decades. SA is also grateful to all the colleagues and collaborators whose names appear in the references cited from his laboratory in this chapter.

#### References


IV. Improved solid phase synthesis of oligodeoxyribonucleotides through phosphotriester intermediates. Nucl Acids Res 8: 1080–1096


treatment of human leukemia in a scid mouse model with c-myb antisense oligodeoxynucleotides. Proc Natl Acad Sci U S A 89(24): 11823–11827


h t tp s://doi.org/10.2165/00003088- 199528010-00002


sec.gov/Archives/edgar/data/86183 8/000095013501501616/b39654hye8-k. txt. Accessed 9 Dec 2020


proliferation by bacterial DNA. J Immunol 147:1759–1764


Nucl Acids Res 48:1691. https://doi.org/ 10.1093/nar/gkaa031


escalation study using morpholino oligomer AVI-4658. Lancet 8:918


properties, stability studies, and biological activity. Bioorg Med Chem 4(10): 1685–1692. 0968089696001605 [pii]


bridged nucleic acids (ENA) with nucleaseresistance and high affinity for RNA. Nucl Acids Res Suppl 1:241–242


silencing of mutant Huntingtin. Mol Ther 19:2178–2185


TLR9 confers responsiveness to bacterial DNA via species-specific CpG motif recognition. Proc Natl Acad Sci U S A 98:9237–9242


IL-23-induced psoriasis. J Investig Dermatol 133:1777–1784


enhances translation through eIF2 α-dependent and independent mechanisms by increasing ribosome density. Nucl Acids Res 45:6023–6036


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 2

# Antisense RNA Therapeutics: A Brief Overview

Virginia Arechavala-Gomeza and Alejandro Garanto

#### Abstract

Nucleic acid therapeutics is a growing field aiming to treat human conditions that has gained special attention due to the successful development of mRNA vaccines against SARS-CoV-2. Another type of nucleic acid therapeutics is antisense oligonucleotides, versatile tools that can be used in multiple ways to target pre-mRNA and mRNA. While some years ago these molecules were just considered a useful research tool and a curiosity in the clinical market, this has rapidly changed. These molecules are promising strategies for personalized treatments for rare genetic diseases and they are in development for very common disorders too. In this chapter, we provide a brief description of the different mechanisms of action of these RNA therapeutic molecules, with clear examples at preclinical and clinical stages.

Key words RNA therapy, Antisense oligonucleotides, Clinical trials, Splicing, Personalized medicine

#### 1 Introduction

Nucleic acid therapeutics is still a growing field. With the irruption of the mRNA vaccines against SARS-CoV-2 special attention has been given to this type of therapies but other types of nucleic acid therapeutics, coined antisense oligonucleotides (AONs), have been studied for many years. Although only a dozen therapeutic oligonucleotides have been formally approved for clinical use, there are many new such drugs in the pipeline for a plethora of (mainly rare) diseases. These AON molecules interact with different nucleic acids (mRNA, non-coding RNA, and DNA) thanks to sequence specific Watson–Crick base pairing. Their mechanism of action, that may be designed to bind specific targets, makes these drugs easy to design, less likely to cause side effects and, therefore, potential candidates to lead the next wave of precision medicine. In this chapter, we describe the most frequently used AON-based therapeutic strategies, their mechanisms of action (Fig. 1), and the results of several clinical trials, with special emphasis in eye and muscle diseases.

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_2, © The Editor(s) (if applicable) and The Author(s) 2022

Fig. 1 Schematic representation of the multiple mechanisms of action of antisense oligonucleotide (AON) molecules. AONs can act at pre- and mRNA levels of the synthesis of a functional protein (left panel). They can be used to modulate splicing (upper right panel) or to degrade (pre-)mRNA (lower right panel). Splicemodulating AONs bind to pre-mRNA and promote the insertion or skipping of regular exons. In addition, they can redirect splicing when mutations in a gene lead to splicing defects (such as pseudoexon insertions). This splicing modulation causes the degradation of the transcript and a consequent reduction of protein levels. Alternatively, transcript degradation can also be achieved by using AONs binding to the pre-mRNA to disrupt the open reading frame and degrade transcripts via nonsense-mediated decay (NMD). Gapmers, in contrast, can bind to both pre-mRNA and mRNA and activate RNase-H1 RNA degradation. (Created with BioRender. com)

#### 2 Mechanisms of Action


effect using therapeutic AONs, and several of the recently approved AON molecules target different DMD exons [3–9]. Because there are many different DMD mutations, the skipping specific exons would be therapeutic for different subsets of patients.

This concept has also been employed in the development of new AONs to skip in-frame exons carrying single-nucleotide changes generating premature stop codons in large genes. Mutations in USH2A cause either Usher syndrome (deafness combined with blindness) or isolated blindness in the form of retinitis pigmentosa. Exon 13 of USH2A is prone to carry truncating variants and by deleting it, a protein with residual function is potentially produced [10, 11]. This is also the case of a stop codon introduced by a variant in exon 36 of CEP290, which is naturally skipped at low levels in the retina and involved in retinal dystrophy. AONs designed to skip exon 36 restored the reading frame and produced a functional protein able to rescue the cellular phenotype in patientderived cells [12]. Following the same strategy, AON molecules to skip different exons of COL7A1 have been developed for dystrophic epidermolysis bullosa, a skin disease inherited in both dominant and recessive fashion [13–16].

2.1.2 Exon Inclusion A seemingly opposite mechanism of action is at the core of nusinersen, an AON approved for the treatment of spinal muscular atrophy (SMA). In this case, mutations in the SMN1 gene cause low or lack of SMN protein production. However, SMN protein can be produced by two nearly identical genes, SMN1 and SMN2. The latter, however, contributes at very low levels due to the high rate of exon-7 skipping that disrupts the ORF. Nusinersen is used to alter the splicing of SMN2 and include exon 7, and therefore produce sufficient amounts of SMN protein to ameliorate the patient's disease [17, 18].

2.1.3 Splicing Redirection Variants close to the splice sites result often in either exon skipping or exon elongation. In the second scenario, mutations decrease the recognition of the original splice site and a cryptic splice site present in the intron is recognized; in the most extreme case, the entire intron is retained. Exonic variants may cause a synonymous or a predicted non-deleterious missense change at protein level, and in addition they can have a dramatic effect at RNA level by creating a novel splice site. In any case, these splicing defects are also amenable for AON intervention. For instance, a mutation in exon 3 of USH1C linked to deafness generates a novel splice donor site (SDS) upstream the regular SDS of the exon. This new SDS is preferentially used by the cells, leading to a disrupted reading frame. By using AONs to block the newly generated SDS, the normal transcript can be produced [19]. A similar approach has been used to target the exon elongation caused by near-exon intronic variants in ABCA4 linked to Stargardt macular degeneration. This study showed that by blocking the newly created SDS the normal splicing can be restored. However, this approach turned out to be not that successful when targeting exon elongations caused by variants in the splice acceptor site (SAS) [20].

Another elegant way to modulate splicing using AONs is by targeting the non-productive transcripts. These transcripts often are generated by (a) alternative splicing causing insertion or skipping of exons; (b) using alternative cryptic splice sites; and (c) retaining the introns. In any case, these splicing events lead to a disrupted ORF being the transcript degraded via nonsensemediated decay (NMD). A very recent study has shown that 1246 potentially disease-associated genes present at least one of these non-productive transcripts. By targeting these splicing events to insert or skip an exon, exclude a retained intron, or redirect splicing when a cryptic splice site is used, the overall protein levels can be increased and this might be a promising therapeutic tool for haploinsufficiency cases [21].


2.2 Transcript Degradation Antisense technology can be extremely useful to degrade transcripts and cause gene silencing (knockdown). From the therapeutic perspective, this might be a potential tool to treat autosomal dominant diseases caused by dominant-negative mutations. In this case, by degrading specifically the mutant allele, the correct protein can perform its function properly.

2.2.1 RNase H1- Activating Antisense Oligonucleotides (Gapmers)

These antisense molecules are characterized for being able to actively reduce the levels of the mRNAs in the nucleus and cytoplasm [31], therefore they are very useful to downregulate gene expression. These RNase H1-activating AONs or gapmers are chimeric molecules linked using a phosphorothioate (PS) backbone that usually present a conformation 5-10-5, where the two arms consist of five modified RNA nucleotides (20 -O-methoxyethyl (20 MOE), 20 -O-methyl (2<sup>0</sup> OMe) or locked nucleic acid (LNA)) flanking ten DNA nucleotides [32]. The first-ever AON approved by the FDA was fomivirsen, a first-generation RNase H1-activating AON [33–36] (see Subheading 3). However, this is the only RNase H1-activating AON that does not have the chimeric RNA/DNA structure. So far, four molecules using this mechanism of action have received FDA and/or EMA approval to treat different disease conditions [32].

Gapmers can be used to downregulate genes in alleleindependent and allele-specific manner. Below, we review some examples of each case.

Allele-independent mRNA degradation is often used to target genes or pathways that are overexpressed in certain disease conditions or can worsen the disease progression. Thus, reducing the levels of particular genes can be very beneficial. This is the case for two of the approved AON drugs: mipomersen and volanesorsen. These molecules target the mRNA of the apolipoprotein B-100 in familial hypercholesterolemia or apolipoprotein C3 in familial chylomicronaemia syndrome, hypertriglyceridemia and familial partial lipodystrophy, respectively, to lower the levels of specific lipids increased in these diseases [37–41].

In contrast, allele-specific mRNA degradation aims to target only the mutant allele. This way, specific mutations that cause a dominant-negative effect can be targeted. This is the case of inotersen, a gapmer designed to target the mRNA encoding the transthyretin (TTR) protein in autosomal dominant hereditary transthyretin amyloidosis [42, 43]. A single-nucleotide change in the gene produces misfolding of the TTR protein. As TTR protein needs to tetramerize in order to conduct its function, the addition of mutant monomers into the tetramer affects the overall function. Systemic amyloid depositions are formed, leading to progressive polyneuropathy of the sensory and motor systems with multiorgan dysfunction in late-disease stages. The therapeutic gapmer targets the mutant allele to reduce the amount of tetramers containing the mutant protein, and therefore prevent the aforementioned depositions [42, 43]. Another recent example is the use of gapmers to specifically degrade the mutant allele introduced by a mutation in the COCH gene, which causes autosomal dominant hearing impairment [44]. In this study, two strategies were used to degrade the mutant transcript: directly targeting the mutation or other single-nucleotide polymorphisms (SNPs) in cis with the mutation that are part of the mutant haplotype.

2.2.2 Disrupting Reading Frame Splice-switching AONs can also be used to induce transcript degradation. Skipping regular exons can also be used to knockdown the function of an undesired gene, by creating mRNA isoforms that encode non-functional proteins or trigger degradation of the mRNA by NMD [45]. For instance, exon skipping of hepatic APOB100 was able to sustainably reduce LDL cholesterol levels in mice [46], downregulation of MAPT gene has been proposed as a possible treatment for tauopathies [47], and skipping exon 2 of ALK5 may modulate the TGF-β signaling cascade, reducing the components related to the overproduction of extracellular matrix in hypertrophic scar [48].

#### 3 Therapeutic Potential

While years ago oligonucleotides were considered a useful research tool and just a curiosity in the clinical market, this has rapidly changed into approved therapeutic strategies for several diseases and promising personalized treatments for many other (rare inherited) diseases. In this section, we will focus on the development of AON-based therapeutic strategies for two particular tissues: muscle and retina.

3.1 Examples of Clinical Trials for Muscle Diseases The use of AONs to treat neuromuscular disorders has been at the forefront of the clinical development of AON-based therapies and more than half of the AONs currently in the market target either Duchenne muscular dystrophy (DMD) or spinal muscular atrophy (SMA). As previously described, AONs targeting the DMD gene aim to skip specific exons to restore the reading frame. This gene has 79 exons and patients present a large variety of mutations, mostly deletions and duplications, that require the design of specific AONs to treat a small subset to patients. The first such drug, eteplirsen, targeted exon 51 of DMD. Skipping this exon could potentially be therapeutic for 13% of DMD patients [3, 6]. Since then, golodirsen, viltolarsen, and casimersen have been approved, all applicable to decreasing percentages of patients [9, 49, 50].

> All DMD exon-skipping drugs currently in the market are phosphorodiamidate morpholino oligomers (PMO). In contrast, the development of the first AON drug in clinical trials for this disorder, drisapersen (a 20 OMe/PS oligonucleotide) [51] as well as that of many others targeting DMD with the same chemistry were halted due to side effects [52]. Despite the apparent success of PMO chemistries to reach the market, these drugs are yet not very efficient, and their clinical outcomes are still poor. This is the main reason why they are yet to be approved in Europe, while in the USA and Japan have been given "accelerated approval" based on dystrophin protein expression as a surrogate endpoint, which is very low and there is debate about its clinical relevance [53]. Currently, several efforts are driven toward increasing the delivery efficacy of these drugs to the target tissue [32, 54]. Several next generation AONs targeting the same exon as eteplirsen (exon 51)

have been or are being developed. This is the case of the stereopure suvodirsen, which was halted after poor results in a phase I clinical trial (NCT03907072) or the peptide-conjugated PMO currently in Phase I/II clinical trials (MOMENTUM, NCT04004065).

While AONs for DMD do not offer yet the clinical benefits that were hoped to achieve at initial stages, the journey to their development has provided very valuable lessons to stakeholders interested in developing these drugs, particularly in the context of orphan drugs [55]. A drug that benefited from some of the previous knowledge was nusinersen, a 20 MOE/PS AON targeting another neuromuscular disorder (SMA). Nusinersen was approved only months after eteplirsen and has been quickly approved worldwide due to the robust clinical data derived from the clinical trials [18, 56]. As described before, this AON is based on an exon inclusion approach to restore the expression of SMN protein in motoneurons. In this case, the target tissue is treated directly by intrathecal infusion, circumventing any delivery hurdles that may have hampered the efficacy of AONs targeting muscle or other organs when delivered systemically. Indeed, nusinersen's delivery approach, chosen chemistry and formulation has been replicated in several n-of-1 clinical trials of other AONs targeting motoneurons, such as milasen and jacifusen (NCT04768972) [57] (see Subheading 4).

3.2 Examples of Clinical Trials for Eye Diseases

The eye is one of the most promising organs for therapeutic development. Among other characteristics, it is contained, easily accessible, and immune-privileged [58]. In fact, the first-ever FDA-approved AON (fomiversen) was a first-class oligonucleotide to treat human cytomegalovirus retinitis, an eye condition in immunocompromised patients [33–36]. Furthermore, a growing group of genes and mutations causing retinal diseases have been targeted at preclinical level using AONs. This includes pseudoexon exclusion for CEP290 [59–63], OPA1 [64], CHM [65] USH2A [66], and ABCA4 [24–29]; splicing modulation for USH2A [10] and CEP290 [12]; or transcript degradation for NR2E3 [67] and RHO [68]. Three of these molecules are currently in different clinical trial phases detailed below.

The most advanced molecule in a clinical setting is sepofarsen (QR-110). This is a 17-mer 20 OMe/PS oligonucleotide aiming to correct the inclusion of a pseudoexon caused by a deep-intronic mutation in CEP290-associated autosomal recessive Leber congenital amaurosis [69]. In the phase 1/2 clinical trial (NCT03140969), all patients were injected with an initial loading dose of either 320 or 160 μg followed by a maintenance dose every 3 months (160 or 80 μg) [70, 71]. Interim results showed that sepofarsen was well tolerated and safe with no serious adverse events [70, 71]. Although the final results of the trial have not yet been published, the improvement that most patients showed led to the design and approval of a phase 2/3 clinical trial (Illuminate, NCT03913143). This is a multi-center, double-masked, randomized, controlled, multiple-dose study to evaluate efficacy, safety, tolerability, and systemic exposure in patients older than 8 years carrying the specific mutation in at least one of the two alleles. Two different doses and a sham-procedure group will be assessed, for a total period of 2 years. In addition, two other clinical trials for the same molecule are ongoing. One is the extension of the phase 1/2 clinical trial to continue treating the patients of the first trial by administering sepofarsen every 3 months in both the already intervened and the contralateral eye (NCT03913130). The second is a multi-center, open-label, dose-escalation, and double-masked randomized controlled trial to evaluate safety and tolerability in children below age of 8 years old (Brighten, NCT04855045).

A multi-center phase 1/2 clinical trial to assess safety and tolerability of QR-421a (Stellar, NCT03780257) is currently ongoing. This 21-mer 2<sup>0</sup> MOE/PS oligonucleotide aims to skip the frequently mutated exon 13 of USH2A [10] causing autosomal recessive Usher syndrome or isolated retinitis pigmentosa. Preliminary results, presented in a press release seem to indicate that QR-421a is well tolerated with no serious adverse events. Furthermore, after treatment with this molecule, improvements in several measures of vision were detected. With these encouraging results, two preliminary phase 2/3 clinical trials have been designed in order to study different patient populations based on the best corrected visual acuity. Both trials will be double-masked, randomized, controlled, 24-month, and multiple-dose study (Sirius and Celeste).

The third molecule in a clinical setting is QR-1123, a gapmer designed to degrade the mutant allele (known as P23H) in the RHO gene [68], which has a dominant-negative effect leading to autosomal dominant retinitis pigmentosa. Thus, the hypothesis is that by degrading the allele carrying the mutation, the other allele will be able to produce a functional protein. This molecule is in an early stage of a multi-center open-label, double-masked, randomized, phase 1/2 trial (NCT04123626).

Other molecules for eye-related genetic diseases in late stages of preclinical development are QR-504a for TCF4-associated Fuchs endothelial corneal dystrophy and QR-411 for pseudoexon exclusion in USH2A-associated Usher syndrome or isolated retinitis pigmentosa.

As well as to target specific mutations, AONs have also been explored for multifactorial eye conditions. This is the case of primary open angle glaucoma, in which TGF-β2 was targeted with a 14-mer 3 + 3 LNA-modified gapmer in a phase I clinical trial. Results showed that the molecule was tolerated, safe and potentially clinically efficacious [72]. Besides this, other type of antisense molecules (small interference RNA, siRNA) have been clinically tested for glaucoma [73], dry eye syndrome [74], diabetic macular edema [75], and age-related macular degeneration [73, 76, 77].

# 4 Future of AON Trials and Personalized Medicine: <sup>n</sup> <sup>¼</sup> 1 Trials?

In 2019, an AON molecule (milasen) to treat a single patient pushed the bounds of personalized medicine and raised many regulatory and ethical questions never explored before for genetic treatments [57, 78].

Milasen was customized exclusively for Mila, a child suffering from a form of Batten disease (neuronal ceroid lipofuscinosis 7) caused by the insertion of an SVA (SINE–VNTR–Alu) retrotransposon, with a detrimental effect on splicing, in the intron 6 of the MFSD8 gene [57]. Using a 22-mer 2<sup>0</sup> MOE/PS AON it was possible to redirect splicing avoiding the insertion of the SVA in the final mRNA transcript. Besides the exclusivity of this treatment, another extraordinary achievement was that it took only 13 months to go from the clinical diagnosis to the first dosing: Mila had a clinical diagnosis in mid-November of 2016, the genetic defect was identified in May 2017, approval to proceed was received in January 2018 and first patient dosing occur in the same month.

The AON delivery regime via intrathecal bolus injection was highly similar to the one of nusinersen, the AON used for SMA [17, 18, 56]. The treatment did not show any safety concerns and the frequency and duration of the seizures was reduced. Unfortunately, despite the treatment had some effect, Mila passed away early 2021. Nevertheless, this study is the hallmark of personalized medicine, and although not all diseases are amenable for this type of therapies, it has highlighted this as a possible approach and managed to re-evaluate the speed and type of safety studies and regulatory requirements. In a similar development, a drug was designed and provided to patient suffering from amyotrophic lateral sclerosis (ALS) with mutations FUS gene, following the same delivery route as nusinersen and milasen. Unfortunately, this patient, Jaci Hermstad, also died recently. However, the drug originally developed for this single patient, ION363 or jacifusen, is currently being tested in a phase III trial for patients with the same disease (NCT04768972). Thus, AON technology can be considered as a platform for individualized treatments which may, sometimes, be extended to other patients.

#### 5 Hurdles

A drawback when compared to small molecule drugs is the relatively large size of AON molecules which limits their delivery into the cells where they exert their action. Therefore, their distribution is limited, their naked uptake is poor, and it is highly determined by the chemistry of their backbones [54]. Often, these AON molecules are not even able to reach their target organ. To circumvent this, most of these AONs rely on their conjugation or formulation with different delivery systems to be able to reach and access their intracellular targets [32]. In addition, when delivered systemically, these molecules can barely reach the central nervous system due to the blood retina and brain barriers. However, as described before, local delivery of naked modified AONs to these specific organs have shown to be efficient and safe in several clinical trials [17, 18, 56, 70, 71, 79].

Another drawback is the high exposure of certain organs upon systemic delivery of AONs. For instance, after intravenous injection of AONs a significant proportion is taken by the liver and kidney. This limits the biodistribution to other tissues and derivate on toxic effects in these organs. However, many of the liver and kidney injuries were found when using high and not clinically relevant doses of AONs [32]. In that sense, novel delivery methods or conjugates are required to be able to target the organs of interest and bypass the high clearance by the liver and kidneys.

Finding proper models to assess the sequence-dependent efficacy and safety of AONs is still a pending issue. Their safety assessment is often performed in rodents, non-human primates, and human plasma. However, these studies only provide sequenceand chemical modification-specific effects. The generation of humanized models have provided very good results, however, generating a humanized animal model for every mutation to be targeted is not feasible nor ethical. It is also possible to generate almost any human cell from patient-derived cells reprogrammed to a pluripotent stage. While these models can provide good readouts at RNA, protein, or even functional levels the entire context will still be missing. Currently, significant efforts are being made in the generation of organ-on-chips. This technology allows the combination of multiple tissues or even organs to study the interaction between them and test therapeutic interventions [80, 81]. In addition, this technology enables other type of measurements that in the near future might be very valuable to perform drug screenings and evaluate the efficacy and safety of many molecules, including AONs [80–83].

Finally, clear guidelines and novel clinical trial designs are needed to explore the full therapeutic potential of AONs when investigated as treatments for rare diseases. The case of milasen has proven that this is possible and new of such trials are being planned.

#### 6 Conclusions

The therapeutic potential of AONs has been, for many years, subject of speculation and theoretical discussion and, while these molecules were widely applied in a research laboratory setting, their clinical application was anecdotal and limited to rare diseases. However, this landscape has recently changed completely thanks to several factors. On one hand, many of such drugs have been approved, being splice-switching AON and siRNA drugs at the forefront of this wave. Secondly, several breakthroughs in the delivery formulation of these drugs have increased the uptake of AONs targeting the liver and this has open wide open the field to consider these as reliable treatment options for several disorders where the liver is the target tissue. Thirdly, much more attention has been given to antisense technology due to the n-of-1 case of milasen. Lastly, RNA-therapies have gained extraordinary popularity due to vaccines against SAR-CoV-2 based on mRNA technology, highlighting the development of drugs based on nucleic acids. All of this will contribute to make these drugs a main resource in the therapeutic toolbox of the twenty-first century.

#### Acknowledgments

V.A-G holds a Miguel Servet Fellowship from the ISCIII (grants CP12/03057 and CPII17/00004), part-funded by ERDF/ FEDER. V.A-G also acknowledges funding from Ikerbasque (Basque Foundation for Science). A.G. group is financially supported by the Foundation Fighting Blindness (PPA-0517-0717- RAD), the Curing Retinal Blindness Foundation as well as the Landelijke Stichting voor Blinden en Slechtzienden and Stichting Oogfonds via Uitzicht 2019-17, together with Stichting Blindenhulp, Rotterdamse Stichting Blindenbelangen and Dowilwo. The funding organizations had no role in the design or conduct of this research and provided unrestricted grants. Both authors are members of the European COST Action DARTER (CA17103).

#### References


Hum Gene Ther 18(9):798–810. https://doi. org/10.1089/hum.2006.061


Popplewell L, Graham IR, Dickson G, Wood MJ, Wells DJ, Wilton SD, Kole R, Straub V, Bushby K, Sewry C, Morgan JE, Muntoni F (2009) Local restoration of dystrophin expression with the morpholino oligomer AVI-4658 in Duchenne muscular dystrophy: a singleblind, placebo-controlled, dose-escalation, proof-of-concept study. Lancet Neurol 8(10): 918–928. https://doi.org/10.1016/S1474- 4422(09)70211-X


J Med 377(18):1723–1732. https://doi.org/ 10.1056/NEJMoa1702752


Collin RWJ, Cremers FPM, Leroy BP, De Baere E (2019) ABCA4-associated disease as a model for missing heritability in autosomal recessive disorders: novel noncoding splice, cis-regulatory, structural, and recurrent hypomorphic variants. Genet Med 21(8): 1761–1771. https://doi.org/10.1038/ s41436-018-0420-y


cardiovascular events and are lowered by inhibition of APOC-III. J Am Coll Cardiol 69(7): 789–800. https://doi.org/10.1016/j.jacc. 2016.11.065


882–890. https://doi.org/10.1016/S1474- 4422(16)30035-7


RWJ, Garanto A (2018) Antisense oligonucleotide-based splicing correction in individuals with leber congenital amaurosis due to compound heterozygosity for the c.2991+1655A>G mutation in CEP290. Int J Mol Sci 19(3):753. https://doi.org/10.3390/ ijms19030753


6447–6454. https://doi.org/10.1167/iovs. 16-20303


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part II

# Design and Formulation of Antisense Technology

# Design of Bifunctional Antisense Oligonucleotides for Exon Inclusion

### Haiyan Zhou

#### Abstract

Bifunctional antisense oligonucleotide (AON) is a specially designed AON to regulate pre-messenger RNA (pre-mRNA) splicing of a target gene. It is composed of two domains. The antisense domain contains sequences complementary to the target gene. The tail domain includes RNA sequences that recruit RNA binding proteins which may act positively or negatively in pre-mRNA splicing. This approach can be designed as targeted oligonucleotide enhancers of splicing, named TOES, for exon inclusion; or as targeted oligonucleotide silencers of splicing, named TOSS, for exon skipping. Here, we provide detailed methods for the design of TOES for exon inclusion, using SMN2 exon 7 splicing as an example. A number of annealing sites and the tail sequences previously published are listed. We also present methodology of assessing the effects of TOES on exon inclusion in fibroblasts cultured from a SMA patient. The effects of TOES on SMN2 exon 7 splicing were validated at RNA level by PCR and quantitative real-time PCR, and at protein level by western blotting.

Key words Antisense oligonucleotide, Bifunctional antisense, Pre-mRNA splicing, TOES, Splice switching, Exon inclusion, Exon skipping

#### 1 Introduction

Harnessing antisense oligonucleotides (AONs) to redirect the altered pre-messenger RNA (pre-mRNA) splicing and modulate target gene expression is an efficient therapeutic strategy for genetic disorders associated with alternative splicing. A number of AON approaches have been investigated on redirecting pre-mRNA splicing. The original strategy is to use AONs complementary to a cryptic splice site to prevent its use and favored selection of the authentic site [1]. This approach has been used regularly to alter the proportion of splice isoforms produced from mutated genes or alternative splicing units. In addition to blocking the splice sites, alternative splicing events are often controlled by regulatory proteins bound to exonic and intronic elements located beyond the alternative splice sites. A valid approach is to use AONs to directly

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_3, © The Editor(s) (if applicable) and The Author(s) 2022

target exonic or intronic elements by blocking the binding of regulatory proteins to these elements that are involved in premRNA splicing. This strategy has been successfully used to augment the exon 7 inclusion in SMN2 gene by using a short AON to target an intronic splicing silencer (ISS) within the gene [2– 5]. Nusinersen, an 18-mer AON annealing to the ISS-N1 element in SMN2 intron 7 is the first antisense drug approved by the US Food and Drug Administration (FDA) for treatment of any types of spinal muscular atrophy (SMA) [6–8]. This strategy has also been proved to be very effective in Duchene muscular dystrophy (DMD) by promoting the skipping of an exon in the DMD gene to restore the interrupted reading frame hence partial rescue of the functional dystrophin protein [9, 10]. Three AON drugs, eteplirsen for exon 51 skipping and golodirsen and viltolarsen for exon 53 skipping in the DMD gene, have been approved by the FDA for treatment of DMD [11–13].

The other splice switching approach is the use of bifunctional oligonucleotides to increase the number of positively or negatively acting signals in an exon or intron and to regulate the alternative splicing. The oligonucleotides were designed with one domain (the antisense domain) annealing to the target exon or intron, and another domain (the tail domain) containing a sequence that either recruits RNA binding proteins involved in pre-mRNA splicing [14] or is made of a synthetic protein domain covalently linked to the antisense domain [15]. This approach may be designed as targeted oligonucleotide enhancers of splicing (TOES) for exon inclusion [14, 16], or as targeted oligonucleotide silencers of splicing (TOSS) for exon skipping [17].

The effectiveness of TOES as a potential therapy for SMA by augmenting exon 7 splicing in SMN2 gene has been approved both in vitro in cellular model [14, 16] and in vivo in mouse model [18– 20]. A bifunctional oligonucleotide targeted to SMN2 exon 7 was expressed in transgenic mice within a modified U7 snRNA gene. Expression of the TOES-U7 RNA in a mouse model of SMA produced a substantial improvement in function and lifespan [20]. Two other bifunctional oligonucleotides targeting the intronic splicing silencers in SMN2 intron 6 and intron 7, which have the dual effects of blocking the silencer and recruiting activator proteins, also showed the potential therapeutic effects in the transgenic mouse models of SMA [18, 19].

We describe here the details in design of bifunctional oligonucleotide for exon inclusion by correcting SMN2 exon 7 splicing as an example (Fig. 1). TOES oligonucleotides are designed to contain two domains, an antisense domain complementary to sequences of SMN2 gene and a tail domain comprising sequences known as binding moieties for splicing activator proteins. The following design principles for TOES oligonucleotides are followed: (1) the antisense sequence may anneal to the potential

Fig. 1 Design of TOES to promote SMN2 exon 7 inclusion. The sequence of SMN2 exon 7 is in upper case and the flanking introns in lower case. Nucleotide 6 in SMN2 exon 7 is T (in red). Two shaded sequences are the binding sites of Tra2<sup>β</sup> and SF2/ASF, respectively. TOES is designed with two functional parts, the antisense domain to anneal to nucleotides 2–16 in SMN2 exon 7, the tail domain containing 3 repeats of "GGAGGAC" motifs to recruit the SR protein SRSF1. Cap contains five nucleotides at the 5<sup>0</sup> -end of the tail (in green, which is chemically modified)

> splicing silencer binding sites in either intron 6, exon 7 or intron 7, and should avoid any splicing enhancer binding sites; (2) a number of splicing enhancer motifs (e.g. SF2/ASF, SRSF1, and hTra2β1) may be included in the tail domain to improve the effectiveness of the oligonucleotides; (3) chemical modification can be applied to the antisense sequence, but not to the tail domain, which may inactive protein binding to the tail domain. The effects on exon inclusion are evaluated at RNA and protein levels in fibroblasts cultured from a patient with type II SMA carrying three copies of SMN2 gene.





#### 3 Methods

3.1 Design of Bifunctional Oligonucleotides 1. Predict the potential binding motifs of the negative splicing regulator heterogeneous nuclear ribonucleoprotein A1 (hnRNP A1) in the target intron or exon sequences, using Human Splicing Finder online software.

	- 2. Cells are cultured in 2 mL of growth medium for 24 h.
	- 3. 24 h later, change the growth medium to 1 mL Opti-MEM and leave the cells in the incubator during the preparation of transfection mixes.
	- 4. Prepare the transfection reagent mixes in sterile 1.5 mL tubes. For each sample, prepare two mixes: the first mix (Mix A) contains 100 μL Opti-MEM and 1 μL AON at desired concentration (e.g. 1 μL AON at 100 μM to get a 100 nM final concentration). While for the mock control add only 100 μL Opti-MEM. The second mix (Mix B) contains 100 μL Opti-MEM and 5 μL Lipofectamine 2000.
	- 5. Mix the AON-containing tube (Mix A) with the lipofectaminecontaining tube (Mix B) at a ratio of 1:1 (100 μL + 100 μL).
	- 6. Incubate the transfection mix for 20 min at room temperature (RT).
	- 7. Add 800 μL Opti-MEM in the transfection mix to top it up to a final volume of 1 mL.
	- 8. Remove Opti-MEM from the 6 well plate and replace with 1 mL transfection mix in each well.
	- 9. Incubate the plate for at least 6 h at 37 -C with 5% CO2 (see Note 4).

Table 2 The reported bi-functional AONs designed for SMN2 exon 7 inclusion


#### 3.3 Splicing Assay of Bifunctional AONs on SMN2 Exon 7 Inclusion at RNA Level


3.4 Bifunctional AONs on Restoring SMN Protein Measured by Western Blotting


#### 4 Notes


#### Acknowledgments

This work was supported by the Wellcome Trust, University College London, UK Medical Research Council (MRC), SMA-Europe, SMA Trust, Muscular Dystrophy UK and NIHR Great Ormond Street Hospital and Institute of Child Health Biomedical Research Centre.

#### References


antisense oligonucleotide design to suppress aberrant SMN2 gene transcript processing: towards a treatment for spinal muscular atrophy. PLoS One 8:2–11


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 4

# Design and Delivery of SINEUP: A New Modular Tool to Increase Protein Translation

### Michele Arnoldi, Giulia Zarantonello, Stefano Espinoza, Stefano Gustincich, Francesca Di Leva, and Marta Biagioli

#### Abstract

SINEUP is a new class of long non-coding RNAs (lncRNAs) which contain an inverted Short Interspersed Nuclear Element (SINE) B2 element (invSINEB2) necessary to specifically upregulate target gene translation. Originally identified in the mouse AS-Uchl1 (antisense Ubiquitin carboxyl-terminal esterase L1) locus, natural SINEUP molecules are oriented head to head to their sense protein coding, target gene (Uchl1, in this example). Peculiarly, SINEUP is able to augment, in a specific and controlled way, the expression of the target protein, with no alteration of target mRNA levels. SINEUP is characterized by a modular structure with the Binding Domain (BD) providing specificity to the target transcript and an effector domain (ED) containing the invSINEB2 element—able to promote the loading to the heavy polysomes of the target mRNA. Since the understanding of its modular structure in the endogenous AS-Uchl1 ncRNA, synthetic SINEUP molecules have been developed by creating a specific BD for the gene of interest and placing it upstream the invSINEB2 ED. Synthetic SINEUP is thus a novel molecular tool that potentially may be used for any industrial or biomedical application to enhance protein production, also as possible therapeutic strategy in haploinsufficiency-driven disorders.

Here, we describe a detailed protocol to (1) design a specific BD directed to a gene of interest and (2) assemble and clone it with the ED to obtain a functional SINEUP molecule. Then, we provide guidelines to efficiently deliver SINEUP into mammalian cells and evaluate its ability to effectively upregulate target protein translation.

Key words SINEUP, Long non-coding RNA, Antisense, Translational increase, Physiological increase, Therapeutic tool, Haploinsufficiency, Protein manufacturing

#### 1 Introduction

The quantitative improvement of protein production in mammalian systems is a compelling need for the industrial manufacturing of commercially available enzymes, antibodies and supplements, but also for gene therapy-based treatments of medical conditions. Several technologies are available to address such a need, however, they usually consist of introducing exogenously constructs containing the protein of interest or directly the target

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_4, © The Editor(s) (if applicable) and The Author(s) 2022

peptide [1]. These approaches still struggle to overcome hazardous but invariable hurdles, especially when used as therapeutic tools, such as ectopic expression and protein quantity modulation, sometimes associated with toxicity [2]. As an alternative, newly identified RNA-based techniques such as small activating RNAs (RNAa) are able to target and upregulate endogenous gene transcription [3]. In 2012, Carrieri et al. discovered a new class of lncRNAs, belonging to the category of natural antisense transcripts (NATs), that have the property to increase the protein translation of the target mRNA [4]. These transcripts were named SINEUP based on their ability to upregulate target protein translation by means of an invSINEB2 repeat, leaving unaltered the transcriptional levels of the target mRNA. They were first discovered in mice, where AS-Uchl1 was found to have a post-transcriptional upregulating activity on its sense protein coding counterpart, Uchl1 mRNA [4]. Later studies confirmed and validated the expression of SINEUP in human cells [5, 6]. SINEUP molecular mechanism relies on its modular structure, composed of two fundamental domains: a Binding Domain (BD)—a region at the 5<sup>0</sup> of the lncRNA overlapping head to head to the 5<sup>0</sup> of the target mRNA—and an Effector Domain (ED)—constituted by an invSI-NEB2 repetitive element. The BD is crucial for target gene pairing and it confers molecular specificity, while the ED is the functional part of SINEUP required for loading the target transcript on polysomes and driving the translational increase [4].

The initial discovery was then supported by the crucial finding that the BD could be engineered in order to target a specific mRNA of interest, as first demonstrated with Green Fluorescent Protein (GFP) [4]. Additionally, miniSINEUP containing only the BD and a shorter version of the original ED were also proven to be effective [7]. This characteristic would enable to overcome the difficulties of long molecules delivery, especially for therapeutic purposes in which naked RNA molecules administration can be proposed. Recently, TranSINE Therapeutics Limited (Cambridge, UK) has been founded to translate the SINEUP technology into the clinics as therapeutics for haploinsufficiency.

All together, these findings qualified SINEUP as a flexible tool able to upregulate the protein production of virtually any mRNA target of interest, affecting solely the translational levels. As such, SINEUP molecules demonstrated to be a suitable tool for protein manufacturing, able to boost, for instance, the production of recombinant proteins and monoclonal antibodies in mammalian cells [8–10]. SINEUP molecules are also being studied from a therapeutic point of view, since their functional characteristics could confer advantages with respect to other gene therapy approaches. SINEUP molecules generally upregulate the endogenous protein of about two- to fivefold (almost within a physiologic range) and they are only effective in those districts where the target mRNA is physiologically expressed, therefore avoiding unspecific effects [10]. Finally, by targeting mRNA molecules, SINEUP does not introduce stable or unwanted changes in the host genome. Thus far, several synthetic SINEUP molecules were successfully designed and delivered as potential therapeutic molecules. For instance, synthetic SINEUP were designed and successfully used to increase the levels of disease-associated proteins in vitro, such as Parkinson's disease-associated DJ-1 in three human neuronal cell lines [7] and Glial-cell derived neurotrophic factor (GDNF) in mouse cell line [11]. Moreover, SINEUP have been used to rescue frataxin levels in a cellular model of Friedreich's Ataxia [12]. Concerning in vivo model systems, SINEUP could effectively rescue some phenotypes associated with microphthalmia with linear skin defects (MLS) syndrome in a medaka fish model of cox7B haploinsufficiency [13]. More recently, SINEUP targeting GDNF mRNA was tested in a neurochemical Parkinson's disease (PD) mouse model [11]. Interestingly, SINEUP-GDNF increased endogenous GDNF level for at least 6 months which lead to an enhancement of dopamine release in the striatum and an amelioration of motor behavior and neurodegeneration, without affecting body weight or food intake, common side effects of the ectopic expression of GDNF [11].

Here, we describe in detail the fundamental steps in order to design, clone, and deliver SINEUP molecular tools in the cellular system of interest.

#### 2 Materials

2.1 Design and Cloning of SINEUP

	- (a) For mAS Uchl1Δ5<sup>0</sup> : 5- CAGTGCTAGAGGAGGTCA GAAGAG-3
	- (b) Rev mAS Uchl1 fl: 5-CATAGGAGTGTTTCATT-3 Or primers from Zucchelli et al. [7]:
	- (a) FWD EcoRI invSINEB2: 5-TATAGAATTCCAGTGCTA GAGGAGG-3

#### 2.2 SINEUP Delivery into Cellular Model

	- 2. Poly-L-ornithine hydrobromide (20 μg/mL).
	- 3. Laminin (3 μg/mL).
	- 4. Dulbecco's Modified Eagle's Medium (DMEM).
	- 5. Ham's F-12 Nutrient (HAM F12).
	- 6. B27.
	- 7. Penicillin-Streptomycin solution.
	- 8. L-Glutamine.

#### 2.3 RNA and Protein Analysis 1. RadioImmunoPrecipitation Assay—RIPA—buffer. 2. Protease and Phosphatase inhibitors (PI and PhI, respectively). 3. Bicinchoninic acid (BCA) or Bradford protein assay kit. 4. Bovine serum albumin (BSA) standard curve. 5. Western blot apparatus. 6. Specific antibody for the target protein and a housekeeping protein. 7. Enhanced ChemiLuminescence (ECL) or equivalent assay. 8. Eurofins PCR primer design tool or equivalent (https:// eurofinsgenomics.eu/en/ecom/tools/pcr-primer-design/). 9. ThermoFisher primer analyzer tool or equivalent (https:// www.thermofisher.com/it/en/home/brands/thermo-scien tific/molecular-biology/molecular-biology-learning-center/ molecular-biology-resource-library/thermo-scientific-webtools/multiple-primer-analyzer.html). 10. DNAse. 11. DEPC-treated (nuclease-free) water. 12. Retro-transcriptase containing both oligo(dT) and Random


#### 3 Methods

Prepare all the solutions using analytical grade reagents with ultrapure water at room temperature, unless otherwise indicated. Solutions used with cells are filtered or sterilized at the beginning. Cells are handled under biological hoods while all the other reactions are performed on the bench. Follow safety instructions indicated by safety team in your institution.

#### 3.1 Binding Domain Design and Cloning


Fig. 1 Zenbu genome browser interrogation of transcript of interest showing TSS usage in selected model system. (a) Zenbu genome browser screenshot showing the genomic location of the gene of interest (in this example GJB2) in the human genome assembly 19 (hg19), green arrow identifies the Translation Initiation Site (TIS). RefSeq accession number for the transcript of interest (NM\_004004, in this case) is also visible. In the enlargement in (b) alternative TSS usage identified by FANTOM5 CAGE library (reported as purple bar plots) is shown. Both TSSs in the same orientation (reverse strand, purple bar plots) and in the opposite orientation (forward strand, green bar plots) of the gene are depicted. The purple box highlights the chosen region in which interrogates the browser about the expression of your target transcript. Main TSSs, TSS1, TSS2, and TSS3, are indicated by purple arrows. (c) Displays the tracks (experiments) in which the transcript is more expressed

(Fig. 1b); in the window below, the tracks expressing your transcript will be displayed (Fig. 1c). Among them, you can select the tissue/cells/organ of your interest.


Fig. 2 SINEUP modular structure: Binding domain (BD) design and cloning. (a) Representation of the modular structure of SINEUP and the design strategy to obtain BD oriented head to head to its target gene. BDs, in light purple, targeting the gene of interest (as an example, the CDS of EGFP is depicted in light green), untranslated region (UTR) in gray, Transcriptional Start Site (TSS) in black, and Transcriptional Initiation Site (TIS) in yellow, are shown. Different suggested lengths for BD are reported. SINEUP effector domain (ED) in light blue, with the antisense region, overlapping TIS is depicted in black-shadowed yellow. Adenine (A) of the TIS is set as 0, BDs lengths vary between 40 from the A and +32 nucleotides, 40 and +4 nucleotides, and 14 and +4 nucleotides, respectively. (b) Example of RNA folding prediction results using RNA Fold web server (http:/rna. tbi.univie.ac.at/cgi-bin/RNAWebSuite/RNAfold.cgi) with default parameters, here reported for EGFP transcript. The blue box displays an enlargement of the TIS (in yellow AUG) surrounding region, in which packed structures without big hairpin-loop are reported. Minimum free energy (MFE) prediction is used to create the output, with base-pair probability in form of a dot plot it depicted. Scale bar reports both base-pair and unpair probability for every base colored dot, with 1 highest probability that the base pair as well as 1 highest probability that the base unpair with the neighbors (c). (Schematic representation of pcDNA 3.1 vector structure obtained from Carrieri C. et al. 2012 [4], BD is depicted in light purple followed by ED in light blue are under CMV promoter)

> 8. Dissolve salt free quality primers in ultrapure water to obtain equal molarity for each primer to a final concentration of 10 μM.


Table 1

List of natural and synthetic SINEUP tested to upregulate protein production. Binding Domain (BD—A and B columns, the overlapping sequence with the sense spliced mRNA is reported) and Effector Domain (ED—C column) administered with different delivery methods (F columns) in diverse model systems (H and I columns) are listed. Position relatively to the targeted Methionine (A column) of the gene of interest (E column) are reported (AUG with A set as 0, M1 refers to first Methionine corresponding to TIS, whereas M76 refers to Methionine in position 76 in the amino acid chain)


(continued)


#### 72 Michele Arnoldi et al.

 [7]

 [16]

 [16]

 [16]

 [16]

 [16]

 [16]

 [16]

 [16]

 [16]

M1

Alu

 [16]

Reference

Table 1


#### Design and Delivery of SINEUP: A New Modular Tool to Increase Protein... 73


74 Michele Arnoldi et al.

Table 1 (continued)


#### Design and Delivery of SINEUP: A New Modular Tool to Increase Protein... 75

(continued)




19. By taking advantage of the restriction sites present in the BD and in the selected plasmid of interest, clone every BD upstream of the AS-Uchl1 ED. A shorter version of ED called miniSINEUP, which still comprehends the invSINEB2 element present in AS-Uchl1, should preferentially be used to obtain shorter SINEUP (see Note 7) in expression plasmid (e.g. pcDNA 3.1 vector from Clontech, Fig. 2c) using T4 DNA ligase (Fig. 2b). If necessary, inducible or tissue specific promoters can be used.

#### 3.2 Effector Domain Design and Cloning You can design the ED and insert in a different plasmid as needed. The ED comprehends an invSINEB2, free right Alu monomer (FRAM) or MIRb (Mammalian-wide Interspersed Repeat type b) transposable element sequence from natural SINEUP (see Note 8). Virtually any of those elements can present SINEUP activity, however, the element present in AS-Uchl1 was previously inserted in miniSINEUP vectors and it was the most widely characterized. For this reason, in this section, we will refer to invSINEB2 cloning as ED.


10. Analyze plasmids integrity on agarose gel and confirm again the presence of the insert and its correct orientation by restriction map and sequencing.

3.3 SINEUP Delivery into Cellular Model Different methods such as transfection, electroporation, or viral vectors transduction, can be chosen to deliver your plasmid into the cells of choice. The selected method should meet a criterion of high plasmid internalization efficiency (>60–70% positive cells). In fact, SINEUP molecules upregulate protein translation in a physiologic range (approximately two- to fivefold) so if few cells receive the plasmid this effect can be hidden or under-estimated. Here, we describe the method which best fits our expectations in hiNPCs [14]. For additional information on SINEUP delivery in animal and cellular models see Note 9.

> In this protocol we focused on our target cells of choice, hiNPCs, for which electroporation represents an optimal method of transfection for transient expression. For this purpose, we used Nucleofector™ Device (Lonza) and selected the recommended protocol for our specific cell line, as well as the advised program (A033) of the device. Dealing specifically with SINEUP, we recommend the following:


Fig. 3 Expected results when SINEUP positively increases target mRNA translation without altering its transcriptional level in human induced neural progenitor cells (hiNPCs). (a) Representative image of hiNPCs expressing EGFP after electroporation with control vector (SINEUP ) or with SINEUP targeting EGFP (SINEUP +). (b) Western Blot image reports target protein expression relative to a chosen reference protein in the presence (+) or absence () of SINEUP against the target mRNA, EGFP, in this example. (c) Bar graph representing fold change quantification of target protein from WB experiments in (b). (d) Bar graphs representing normalized expression level of target mRNA upon SINEUP administration ( control SINEUP, + SINEUP against target EGFP mRNA). SINEUP presence is depicted in (e). Transcripts expression was obtained from qPCR quantification and normalized with a chosen reference gene

#### 3.4 SINEUP Efficacy Assessment: Transcription and Translation Evaluation

Successful SINEUP function needs to be assessed by transcriptional and translational analysis, such as qPCR and WB, respectively. In particular, an increase in protein production between two- and fivefold changes is expected (Fig. 3b, c) without dysregulation of the target mRNA expression (Fig. 3d) when SINEUP is present (Fig. 3e).


#### 4 Notes


[17]. Among others, we suggest 40/+32, 40/+4, and 14/+4 BD regions as the most promising target to be chosen for the experiments (as reported in Fig. 2a).


experiment is positively responding to SINEUP administration. For this reason, we suggest testing the previously characterized SINEUP against GFP [4, 7], before moving to SINEUP targeting the gene of interest. To analyze the SINEUP efficacy on GFP translation, WB analysis as well as imaging analysis with semi-automatic detection methods can be performed [22]. Of importance, to underlie that protein translation is increased due to SINEUP mechanism, qPCR should be conducted and report stable expression of the target transcript.


#### References


cells. MAbs 10(5):730–737. https://doi.org/ 10.1080/19420862.2018.1463945


RNA with SINEB2 repeat. Nucl Acids Res 43(9):e58. https://doi.org/10.1093/nar/ gkv125


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 5

# How to Design U1 snRNA Molecules for Splicing Rescue

### Liliana Matos, Juliana I. Santos, Mª. Francisca Coutinho, and Sandra Alves

#### Abstract

Mutations affecting constitutive splice donor sites (5<sup>0</sup> ss) are among the most frequent genetic defects that disrupt the normal splicing process. Pre-mRNA splicing requires the correct identification of a number of cis-acting elements in an ordered fashion. By disrupting the complementarity of the 5<sup>0</sup> ss with the endogenous small nuclear RNA U1 (U1 snRNA), the key component of the spliceosomal U1 ribonucleoprotein, 50 ss mutations may result in exon skipping, intron retention or activation of cryptic splice sites. Engineered modification of the U1 snRNA seemed to be a logical method to overcome the effect of those mutations. In fact, over the last years, a number of in vitro studies on the use of those modified U1 snRNAs to correct a variety of splicing defects have demonstrated the feasibility of this approach. Furthermore, recent reports on its applicability in vivo are adding up to the principle that engineered modification of U1 snRNAs represents a valuable approach and prompting further studies to demonstrate the clinical translatability of this strategy.

Here, we outline the design and generation of U1 snRNAs with different degrees of complementarity to mutated 5<sup>0</sup> ss. Using the HGSNAT gene as an example, we describe the methods for a proper evaluation of their efficacy in vitro, taking advantage of our experience to share a number of tips on how to design U1 snRNA molecules for splicing rescue.

Key words U1 snRNA-based therapy, Splicing modulation, 5<sup>0</sup> ss mutations, Aberrant exon skipping, Modified U1 snRNA, Mucopolysaccharidosis IIIC

#### 1 Introduction

The U1 small nuclear ribonucleoprotein (U1 snRNP) is a key molecule involved in an early event of the splicing process. Like other snRNPs involved in the overall splicing regulation process, it contains a small RNA complexed with several proteins, namely seven Smith antigen (Sm) proteins and three U1-specific proteins (U1A, U1C, and U170K) [1]. U1 snRNA, the RNA component of the U1 snRNP is a 164 nucleotides-long molecule whose 5<sup>0</sup> end interacts by complementarity with the 5<sup>0</sup> splice donor site (5<sup>0</sup> ss). That interaction between the single stranded 5<sup>0</sup> tail of the U1 snRNA molecule and the moderately conserved stretch of nucleotides that constitutes the 50 ss (CAG/GURAGU, where R is a

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_5, © The Editor(s) (if applicable) and The Author(s) 2022

purine) marks the exon-intron boundary and initiates spliceosome assembly [2]. About 40%, 22%, and 5% of normal 50 ss contain two, three, or four mismatches towards the U1 snRNA, respectively [3, 4]. This variable degree of degeneration is among the major factors that significantly contribute to hinder a clear prediction of the effect of mutations flanking the canonical GU site. Furthermore, there is a number of additional elements, which may influence the splice site selection and need to be taken into account such as splicing silencer and enhancer motifs, the presence of alternative splice sites, secondary structures, and regulatory proteins [5]. Therefore, a straightforward prediction of the effect of mutations flanking the canonical GU site without a direct assessment of the mature mRNA produced can be quite challenging. Interestingly, however, it is also the variable degree of degeneration of 5<sup>0</sup> ss and the surprising heterogeneity existing among human spliceosomal snRNA, which allows for splicing correction using modified exogenous U1 snRNAs.

Overall, the rationale on the use of modified U1 snRNAs to correct splicing defects is as simple as it can be: as 5<sup>0</sup> ss mutations alter the 5<sup>0</sup> ss recognition by the endogenous U1 snRNA, exogenous U1 snRNAs may be engineered through complementary base pairing in order to correctly recognize the mutated allele and initiate spliceosome assembly, thus suppressing the mutation effect.

So far, the effects of modified U1 snRNAs have been tested in vitro in a number of cellular platforms from patient-derived cells to model cell lines overexpressing the splicing defects under study, and their potential to either fully or partially correct those mutations was demonstrated for a number of different diseases [5, 6]. Importantly, the application of this sort of modified U1 snRNAs in animal models has also been addressed in recent studies, with a few promising results reported to date [7–10] (see Note 1).

Globally, mutations affecting constitutive 5<sup>0</sup> ss represent roughly 8% of all known genetic disease-causing variants. Their pathogenicity derives from the reduced complementarity of the U1 snRNA to the 50 ss. 50 ss mutations mostly result in exon skipping but their effect over splicing may vary. Currently, there are a number of in silico tools that may help predict disease-causing effects, but cDNA analysis remains mandatory for a proper assessment of their consequence over splicing. For example, mutations affecting RNA splicing represent more than 20% of the mutant alleles in Mucopolysaccharidosis type IIIC (MPS IIIC; HGSNAT gene), a rare lysosomal storage disorder that causes severe neurodegeneration. Many of these mutations are located in the conserved splice donor or acceptor sites, while few are found in the nearby nucleotides. For three mutations that affect the donor site, we have previously developed different modified U1 snRNAs with compensatory changes that may allow for proper recognition of the mutated 50 ss, in an attempt to rescue the normal splicing process.

Fig. 1 Modified U1 snRNA therapeutic approach to correct the pathogenic effect of a 5<sup>0</sup> splice site mutation on the HGSNAT gene. (a) Schematic illustration of base pairing between the wild-type U1 (U1-WT) and the 5<sup>0</sup> ss of wild-type and mutant exon 2 of the HGSNAT gene. The mutation position in the 5<sup>0</sup> ss is marked in grey and it is in italics. The different U1 snRNAs used for the mutated 5<sup>0</sup> ss of HGSNAT (designated as U1-sup, for suppressor) are also shown. The U1 sequence modifications are illustrated in bold. (b) RT-PCR analysis of the endogenous splicing pattern of control and MPS IIIC patients derived fibroblasts after transfection with different U1 isoforms. The constitutive splicing of exon 2 of the HGSNAT gene was not altered in control fibroblasts after overexpression of U1-WT or any of the modified U1 constructs. In the MPS IIIC patients 1 (MPS IIIC P1) and 2 (MPS IIIC P2), bearing the homozygous mutation c.234+1G>A, only the fully adapted U1 (U1-sup4) resulted in partial correction of exon 2 skipping

For the c.234+1G>A mutation, a totally complementary U1 snRNA allowed for partial correction of exon 2 aberrant splicing in patients' fibroblasts (Fig. 1) [11]. Here, we take advantage of our experience on the development of modified U1 snRNAs to compensate for those HGSNAT mutations, to present a practical overview on how to design U1 snRNA molecules for splicing rescue.

In summary, we present an overview of the experimental design for in vitro testing the potential of modified U1 snRNA vectors to correct aberrant splicing caused by 50 ss mutations. Briefly, we show: (a) how to design in silico U1's with different degrees of complementarity to each mutated 50 ss by introducing a number of sequence changes, and (b) how the different U1 vectors harboring those alterations are obtained by site-directed mutagenesis of the original wild-type (WT) human U1 snRNA-harboring pG3U1 vector [12], a derivative of pHU1 [13]. We also describe how these molecules are transfected into patients' fibroblasts and how their effectiveness on splicing redirection can be assessed by posttransfection cDNA analysis and sequencing. Finally, we elaborate on the relevance of further addressing the treatment's effect at protein level.

#### 2 Materials

2.1 Generating Modified U1 snRNA Vectors Adapted to the 50 ss of Interest

	- 2. The sequence of the 50 ss of interest for splicing rescue can be found in Ensembl or other reference sequence databases (in this particular chapter we used the Homo sapiens HGSNAT gene sequence, ENSG00000165102).
	- 3. pG3U1 vector [12] a derivative of pHU1 [13] (see Note 2).
	- 4. Sense and antisense mutagenic primers.
	- 5. PCR mutagenesis kit.
	- 6. PCR thermocycler.
	- 7. Chemically Escherichia coli competent cells (Homemade or commercial; usually are included in the PCR mutagenesis kits).
	- 8. Water bath.

#### 2.2 In Vitro Therapeutic Evaluation of Modified U1 snRNA Vectors in Human Fibroblasts

2.2.1 Transfection of Modified U1 snRNA Vectors in Human Fibroblasts


#### 3 Methods

#### 3.1 Generating the Modified U1 snRNA Vectors

To design the primers for producing the desired modified human U1 snRNA vectors, it is first necessary to know the sequences of the 50 ss under study, both WT and mutant. Then, it is necessary to analyze the complementarity of those sequences with that of U1 snRNA. Next, several modified U1 snRNA vectors can be designed and constructed to have different complementarities to the target sequences (Fig. 2). To generate those constructs, the plasmid pG3U1 [12] (kindly provided by Prof. Dr. Bele´n Pe´rez) a derivative of pHU1 [13], containing the coding sequence of the human U1 can be used as template for site-directed mutagenesis PCR reactions (see Note 2). Depending on the number of mutations to insert in the U1 snRNA vector sequence, different mutagenic primer pairs need to be designed.

Fig. 2 Design and construction of modified U1 snRNA vectors. (a) Schematic representation of base pair interactions between the U1 snRNA and the wild-type and mutant 5<sup>0</sup> ss of HGSNAT exon 2, respectively. (b) Illustration of the strategy followed to increase the complementarity of U1 snRNA with the mutated 5<sup>0</sup> ss of HGSNAT gene. U1 complementarity was increased stepwise, and to try to compensate for the HGSNAT mutation at +1 position, four different U1-adaptations were designed [U1 sup1 (+1T); U1 sup2 (-1G +1T); U1 sup3 (-1G +4A); U1 sup4 (-1G +1T +4A)]. Upper case letters show exonic nucleotides, whereas the lower case letters denote intronic nucleotides. Base pairing is indicated by vertical lines and its loss by an <sup>X</sup>. The mutant nucleotide is highlighted in red and the changed nucleotides in the U1 sequence are illustrated in green 3.1.1 Engineering Modified U1 snRNA Vectors Adapted to the 5<sup>0</sup> ss of Interest


3.2 In Vitro Therapeutic Evaluation of Modified U1 snRNA Vectors in Human Fibroblasts

3.2.1 Modified U1 snRNA Vectors Transfection in Human Fibroblasts

containing ampicillin (100 μg/mL) to a sterilized flask(s) (see Note 11) and innoculate all the bacterial growth from the miniculture(s). Incubate the flask(s) in an orbital shaking incubator at 37 C and 220 rpm for 16–18 h. Using an endotoxinfree maxiprep plasmid DNA purification kit, maxiprep the plasmid(s) containing the modified U1 snRNA construct (s) and perform its sequencing analysis for validation.

Even though we must always find a balance between the best possible experimental design and the resources available, adequate controls may never be forgotten. Still, there is a minimum standard for cell culture experiments that must always be met if we want to draw strong conclusions out of them. Therefore, adequate controls to the variables under test should always be included (see Note 14).


$$N = \frac{\sum \mathfrak{n}}{8} \times 10^4$$

where N is the total number of cells per milliliter, n is the number of cells counted in each quadrant of the Neubauer chamber and the 104 factor allows for the correction of the total number of cells in 1 mL of cell suspension.

	- 2. For reverse transcription, use a cDNA synthesis kit following the manufacturer's protocol, and start with 1–2 μg of total RNA. The cDNA synthesis reaction can be stored at -20 C or used immediately for PCR amplification.
	- 3. Perform a PCR in standard conditions using a Taq polymerase supplemented with its buffer, dNTPs, gene-specific primers for a final concentration of 0.4 μM each (e.g. HGSNAT primers), 2 μL of cDNA, and RNase free water to a final volume of 50 μL.
	- 4. To evaluate the splicing rescue, analyze the amplification products through agarose gel electrophoresis in an agarose gel stained with 5 μL of ethidium bromide (see Note 17). Choose a DNA ladder according to the size of the amplified band. After separation, visualize the gel using an UV transilluminator. As an example, Fig. 1 shows the results of the partial correction of HGSNAT exon 2 splicing after expression of a modified U1 snRNA (totally complementary to the 50 ss of exon 2) in patients' fibroblasts.
	- 5. Assess the identity of the obtained band(s) by sequencing analysis (see Note 18). For this purpose, purify the PCR products directly with a PCR clean-up kit if there is only one amplified band or when multiple bands are present excise each band from the gel and purify them using a gel band

purification kit. Whatever the case, follow the indications present in the manufacturer's protocol.

6. Subject the purified bands to standard automated sequencing using gene-specific primers for the amplification (e.g. HGSNAT primers). Compare the obtained sequence (s) with the reference sequence of the gene of interest (retrieved from the Ensembl database) using the Clustal Omega bioinformatic tool (https://www.ebi.ac.uk/Tools/ msa/clustalo/), in order to analyze the effect of the modified U1 snRNA's in rescuing the normal splicing pattern.

While not included in this chapter, for it is case-specific, the effect of modified U1 snRNAs-treatment at protein level is mandatory whenever we want to proceed to in vivo studies in order to address the true therapeutic potential of a given U1 snRNA molecule.

Ideally, as soon as we get an RT-PCR pattern that confirms splicing correction to some extent, and that rescue is confirmed by band excision and Sanger sequencing, the overall effect of that rescue at protein level should also be checked. There is a variety of methods we can choose in order to address this issue, from the direct quantification of enzymatic activity (whenever the gene product under analysis has a catalytic activity) to that of the protein itself (through Western blot).

Usually, the method of choice depends on two major factors: the protein itself and the assays available in house to assess it. Virtually every method from Western blot to immunofluorescence may be informative and provide extra support to the conclusions drawn from the RT-PCR. Therefore, as a take-home message, we would recommend that, whenever designing U1 snRNA molecules for splicing rescue, the effect should be checked not only at cDNA level, but also at protein level.

#### 4 Notes


3.2.3 Assessment of the Effect of U1 snRNA-Induced Splicing Rescue at Protein Level

identification of the splicing defects, and in vitro rescue by U1 snRNA} was used, but the human U1 snRNA sequence can be cloned in other standard expression vector(s).

	- (a) both mutagenic primers must contain the desired mutation(s) and anneal to the same sequence on opposite strands of the plasmid;
	- (b) each primer should have between 25 and 45 bases in length with a melting temperature (Tm) of 78 C;
	- (c) the desired mutation(s) should be located in the middle of the primer (~12–15 nucleotides of the correct sequence on both sides);
	- (d) the primers should have a minimum GC content of 40% and should terminate in one or more C or G bases;
	- (e) the primers do not need to be 5<sup>0</sup> phosphorylated and purification may either be performed by liquid chromatography (HPLC) or by polyacrylamide gel electrophoresis (PAGE).

snRNA cassette from pG3U1 vector in a plasmid encoding GFP and monitor fluorescence and U1 expression simultaneously.


#### Acknowledgments

The authors would like to acknowledge Prof. Dr. Bele´n Pe´rez (Molecular Biology Department, Faculty of Sciences, University Autonoma of Madrid, Spain) for kindly provide the pG3U1 vector. In addition, the authors would like to acknowledge BioMedCentral, Part of Springer Nature for allowing the reproduction in this chapter of an adapted version of a figure originally published in Matos, L. et al. Therapeutic strategies based on modified U1 snRNAs and chaperones for Sanfilippo C splicing mutations. Orphanet J Rare Dis 9, 180 (2014). https://doi.org/10.1186/ s13023-014-0180-y. Copyright © 2014, Springer Nature. This work was partially supported by Fundac¸a˜o para a Cieˆncia e Tecnologia (FCT) IP (project: FCT/PTDC/BBBBMD/6301/2014) and by The National MPS Society (project: 2019DGH1642).

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 6

# Conjugation of Nucleic Acids and Drugs to Gold Nanoparticles

### Paula Mila´n-Rois, Ciro Rodriguez-Diaz, Milagros Castellanos, and A´ lvaro Somoza

#### Abstract

Gold nanoparticles (AuNPs) can be used as carriers for biomolecules or drugs in cell culture and animal models. Particularly, AuNPs ease their internalization into the cell and prevent their degradation. In addition, engineered AuNPs can be employed as sensors of a variety of biomarkers, where the electronic and optical properties of the AuNPs are exploited for a convenient, easy, and fast read out. However, in all these applications, a key step requires the conjugation of the different molecules to the nanoparticles. The most common approach exploits the great affinity of sulfur for gold. Herein, we summarize the methods used by our group for the conjugation of different molecules with AuNPs. The procedure is easy and takes around 2 days, where the reagents are slowly added, following an incubation at room temperature to ensure the complete conjugation. Finally, the unbound material is removed by centrifugation.

Key words Gold nanoparticles, Spherical nucleic acid, Functionalization, Oligonucleotides, Nanomedicine, Metal nanoparticles, Conjugation, Drug delivery, Sensors

#### 1 Introduction

Oligonucleotides and drugs face some challenges for their optimal delivery in cells and animal models. Particularly, oligonucleotides (e.g., antisense, gapmers, and siRNAs) usually present low stability and suffer from reduced cell internalization and selectivity [1, 2] and, for these reasons, transfection reagents such as lipofectamine are usually employed to improve delivery. On the other hand, drugs can be too hydrophobic and require solubilizing molecules (e.g., dimethylsulfoxide [DMSO], ethanol). However, these kinds of chemicals present critical restrictions such as cytotoxicity or limited loading. To overcome these drawbacks, delivery systems based on nanoparticles can be employed [3]. There are different types of nanoparticles such as liposomes, micelles, dendrimers, inorganic

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_6, © The Editor(s) (if applicable) and The Author(s) 2022

particles, carbon-based nanostructures, viral nanocarriers, polymeric, peptide or metallic nanoparticles, etc. [4–12]. Each vehicle presents different characteristics that can be exploited to address specific challenges related to the delivery of bioactive molecules.

Among the different systems, gold nanoparticles (AuNPs) present excellent properties for the delivery of oligonucleotides because of their low toxicity, cost, and particularly their ease of preparation and functionalization [13]. AuNPs can be synthesized in the laboratory through simple methods, such as the one described by Turkevich [14] and detailed in Subheading 3.1.

The properties of AuNPs can be tuned through their modification with oligonucleotides. When the nanoparticles are densely loaded with oligonucleotides, the resulting nanostructures are known as spherical nucleic acids (SNA) [15]. This kind of nanostructure presents interesting features, such as high internalization in a wide variety of cells and low toxicity. Therefore, these derivatives can be employed for multiple applications, such as drug delivery systems, gene therapy and regulation, or molecular diagnosis [16, 17].

Regarding the vehiculization of therapeutics, AuNPs can be used for the delivery of hydrophobic drugs such as paclitaxel, doxorubicin, or AZD8055 without affecting their effectiveness [18, 19]. On the other hand, AuNPs functionalized with oligonucleotides (e.g., siRNAs, gapmers) could be used as a substitute for transfection reagents in different applications involving gene regulation, or even immunomodulatory processes, for the treatment of diseases such as cancer, sepsis, skin disorders, diabetes, etc. [16, 20– 22].

In the case of diagnostics, it is worth mentioning that fast and accurate point-of-care diagnostic systems are critical in personalized medicine. In particular, nucleic acid detection is of great importance for the diagnosis and treatment of many diseases caused by genetic mutations, infectious agents, or other physiologically abnormal circumstances. Conventional methods such as RT-PCR offer high accuracy and sensitivity; however, these methods are not suitable for routine diagnosis because they are time-consuming and need highly trained personnel and expensive equipment. One development that seems to simplify the nucleic acid detection and we study in the lab is the use of SNA based on a single-stranded oligonucleotide with a unique stem-loop structure (Molecular Beacon, MB) [23, 24].

This chapter describes how to conjugate drugs or oligonucleotides to AuNPs, which can be further used as delivery systems of therapeutics and sensors.

To attach any compound to AuNPs, the high affinity of thiol groups to gold could be exploited. Thus, the molecules (e.g., oligonucleotides, drugs) should be functionalized with linkers containing sulfur-based moieties, such as thiols or dithiolanes [25], which are commented in this chapter.

AuNPs conjugation requires a few simple steps of addition, incubation, and washes. The method might change slightly depending on the linker employed for the conjugation, which can be designed to control the release or stability of the cargo. In general, the use of dithiolane provides more robust structures and can be achieved in few hours, whereas the use of thiols implies more than 1 day. For the reader's convenience, we have included the preparation of the dithiolane linkers used in our group. The approach can be used for the conjugation of drugs, polymers (e.g., polyethylene glycol [PEG]), or the preparation of oligonucleotides in a DNA synthesizer using a tailored solid support, usually based on controlled pore glass (CPG).

#### 2 Materials




#### 3 Methods



$$\text{concentration} = \frac{A}{\varepsilon \times l} \tag{1}$$

where A is the absorbance at 520 nm, l is the optical path in cm, and ε extinction coefficient in M<sup>11</sup> cm<sup>1</sup> .

3.2 Dithiolane-Based The preparation of the dithiolane-based derivatives of drugs (3) and PEG (2) is summarized in Fig. 1 and described in the following instructions. In the case of oligonucleotides, the required solid support (CPG) containing a dithiolane moiety for the preparation of oligonucleotides is also described (7).


Linkers Synthesis

3.2.1 Compound 1 [2,5- Dioxopyrrolidin-1-yl(R)-5- (1,2-Dithiolan-3-yl) Pentanoate]

Fig. 1 Schematic representation of the synthesis of dithiolane-modified products: PEG, drug, and CPG


3.2.5 Compound <sup>5</sup>: N-(1- (bis(4-Methoxyphenyl) (Phenyl)Methoxy)-3- Hydroxybutan-2-yl)-5-(1,2- Dithiolan-3-yl) Pentanamide

3.2.6 Compound 6: 4- ((3-(5-(1,2-Dithiolan-3-yl) Pentanamido)-4-(bis(4- Methoxyphenyl)(Phenyl) Methoxy)Butan-2-yl)oxy)- 4-Oxobutanoic Acid

3.2.7 Compound 7: 4- ((3-(5-(1,2-Dithiolan-3-yl) Pentanamido)-4-(bis(4- Methoxyphenyl)(Phenyl) Methoxy)Butan-2-yl)oxy)- 4-Oxobutanamide CPG

3.3 AuNP Functionalization with Thiol-Modified Oligonucleotides


Oligonucleotides can be easily attached to AuNPs using a thiolbased linker, which is commercially available, and most oligonucleotide providers offer this modification. However, the thiol group should be deprotected, as detailed below, before incubating the oligonucleotides with AuNPs (Fig. 2).

1. Incubate the oligonucleotide with TCEP (see Note 2) using a 100excess relative to the oligonucleotide's thiol (see Note 3)

Fig. 2 Schematic representation of: (a) Deprotection of oligonucleotides bearing a thiol moiety. (b) Functionalization of AuNPs with thiol-modified oligonucleotides

for 2 h at room temperature and moderate agitation on a minishaker (e.g., for deprotect 500 μL of an oligonucleotide solution at 20 μM (i.e., 20 pmol/μL) use 2 μL of TCEP at 0.5 M) (see Note 4).


unbound oligonucleotide from the solution (see Subheading 3.4) (see Note 7).


Note 5). Vortex the solution quickly after each addition of NaCl solution and incubate the sample for at least 10 min on a mini-shaker between each addition.


3.5.2 AuNP Functionalization with Dithiolane-Modified Drugs In this case, the drugs have to be modified with a dithiolane-based linker (Fig. 4), and the AuNPs should be stabilized with oligonucleotides or PEG containing a sulfide-based linker. In this case, for 1 mL of 13 2 nm AuNP (12 nM) add at least 2000 pmol of stabilizing agent (e.g., PEG, oligonucleotide) and then the required amount of the modified drug for a total of 10,000 pmol (stabilizing agent + drug) in the solution.


#### 4 Notes


Fig. 3 Schematic representation of AuNPs functionalization with dithiolane-modified oligonucleotides

Fig. 4 Schematic representation of AuNPs functionalization with a dithiolane-modified drug using oligonucleotides or PEG as stabilizers


30 min at 13.2 rpm. Collect the supernatant and measure it as described in Subheading 3.3 and 3.4.

8. Drug quantification could also be done by comparing the drug supernatant absorbance or fluorescence with a proper standard linear calibration curve of the drug [19] or using analytical chromatography (HPLC).

#### References


331–345. https://doi.org/10.1007/s00775- 018-1542-z


therapeutics. Nanoscale 6:7436–7442. https://doi.org/10.1039/C4NR00019F


with hydrophobic molecular beacon-like structures. Chem Commun 50(23):3018–3020. https://doi.org/10.1039/c3cc47862a


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Determination of Optimum Ratio of Cationic Polymers and Small Interfering RNA with Agarose Gel Retardation Assay

#### Omer Aydin, Dilek Kanarya, Ummugulsum Yilmaz, and Cansu € Umran Tunc¸

#### Abstract

Nanomaterials have aroused attention in the recent years for their high potential for gene delivery applications. Most of the nanoformulations used in gene delivery are positively charged to carry negatively charged oligonucleotides. However, excessive positively charged carriers are cytotoxic. Therefore, the complexed oligonucleotide/nanoparticles should be well-examined before the application. In that manner, agarose gel electrophoresis, which is a basic method utilized for separation, identification, and purification of nucleic acid molecules because of its poriferous nature, is one of the strategies to determine the most efficient complexation rate. When the electric field is applied, RNA fragments can migrate through anode due to the negatively charged phosphate backbone. Because RNA has a uniform mass/charge ratio, RNA molecules run in agarose gel proportional according to their size and molecular weight. In this chapter, the determination of complexation efficiency between cationic polymer carriers and small interfering RNA (siRNA) cargos by using agarose gel electrophoresis is described. siRNA/cationic polymer carrier complexes are placed in an electric field and the charged molecules move through the counter-charged electrodes due to the phenomenon of electrostatic attraction. Nucleic acid cargos are loaded to cationic carriers via the electrostatic interaction between positively charged amine groups (N) of the carrier and negatively charged phosphate groups (P) of RNA. The N/P ratio determines the loading efficiency of the cationic polymer carrier. In here, the determination of N/P ratio, where the most efficient complexation occurs, by exposure to the electric field with a gel retardation assay is explained.

Key words Small interfering ribonucleic acid (siRNA), Agarose gel retardation assay, siRNA/cationic polymer carrier complex, Nanoparticles, N/P ratio, Gene delivery

#### 1 Introduction

Regulation of a specific gene has been used for the treatment of a wide range of diseases such as cardiovascular diseases [1], neurodegenerative diseases [2], and cancer [3]. RNA Interference (RNAi) has become a powerful tool for gene silencing studies due to its advantageous properties such as high specificity, effectiveness, a minimum amount of side effect, and easy preparation [4]. RNAi mechanism was first determined by Andrew Z. Fire, Craig

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_7, © The Editor(s) (if applicable) and The Author(s) 2022

C. Mello, and their colleagues [5]. As a result of their studies, they received the Nobel Prize in Physiology or Medicine in 2006. Interfering RNAs have the ability to silence target genes in cells [6]. At this silencing process, 18–31 nucleotides length small RNA molecules are introduced into cells and induce a sequence-specific gene silencing at the post-transcriptional level by blocking mRNAs containing a matched sequence.

siRNA is the most commonly used interfering RNA and has shown high potential as a therapeutic RNA for gene-based treatments [7]. It regulates the expression of various genes by binding to mRNAs in the cell cytoplasm and causing degradation of their mRNA target. The siRNA is double stranded in nature and is about 22 nucleotides in length. Its precursor is initially recognized by Dicer RNase and then incorporated with the RNA-induced silencing complex (RISC). The siRNA-RISC complex can bind the targeting region of the mRNA and lead to a sequence-specific cleavage with endonuclease Argonaute-2 (AGO2), thereby reducing the expression of the targeted protein [8].

Although siRNA has had particular interest in research, there are some limitations. The major limitations of siRNAs-based therapeutics are their rapid degradation by serum nucleases, poor cellular uptake due to the negatively charged backbone, rapid renal clearance following systemic delivery, off-target effects, and induction of immune responses [9]. In addition, even after siRNA is released from the endosome without being exposed to the lysosome and released into the cell cytoplasm, gene silencing of the siRNA may not be immediately observed [10]. Thereby, before the silencing therapeutic effect of siRNA begins, there is always an induction period due to the intracellular half-life of the target protein. The silencing effect of the given siRNA decreases over time owing to the natural degradation of the siRNA molecules in the cell. Moreover, the therapeutic effect persists for a limited time in rapidly dividing cells such as cancer cells, due to the continuous dilution of siRNA in the replication.

Bare siRNA molecules have poor cellular internalization and they need a carrier to enter cells, where their mechanism of action occurs. The major challenge in the delivery of nucleic acids is the availability of a suitable carrier for transferring siRNA to target cells. To do that, there are two main approaches such as viral and non-viral vectors [11]. The suitable vectors should provide a high degree of transfection for a long period without causing systemic toxicity and immunogenicity [12]. Despite the high transfection activity of viral vectors, possible damage to host genes, immune system stimulation, and infection potential limit its application for gene therapy [13].

To overcome these limitations different types of delivery systems have been designed such as lipid [14, 15], polymer [16], peptide [17, 18], dendrimer [19–21], and micelle [22] based vehicles. An ideal siRNA delivery system should be non-toxic, safe, and effective. Thus, many studies have focused on the development of non-viral vectors with minimal toxicity. Furthermore, the carrier systems should assure entrance of siRNA cargos to the cell cytoplasm without being interrupted by biological barriers such as serum, cell membrane, and endosome/lysosome [23]. In detail, the cell entry of siRNA/cationic polymer carriers is mostly facilitated by the mechanism known as "endocytosis" [10]. In particular, siRNA/cationic polymer carriers should be able to have endosomal escape. Otherwise, siRNA could be degraded in the acidic and enzymatic milieu of endosome/lysosome [24].

Cationic polymers and lipids have frequently been employed in research due to their advantages in gene delivery such as biodegradability, low cytotoxicity, structure variety, and easy scale-up production [25]. Therapeutic nucleic acid cargoes are loaded into the carrier systems mostly through the positive–negative charge interactions between positive charges of carrier and negative charges of phosphate groups in RNA. However, cationic polymer carriers with excess of positive charge may cause toxicity. Cationic carriers cause considerable disruption of cellular membrane integrity because of the negatively charged constitution of cell membrane [26]. Moreover, cationic nanocarriers induce cell necrosis due to the positive charge [27]. They also cause mitochondrial and lysosomal damage and formation of a high number of autophagosomes [28]. In order to overcome this problem, smart carriers have been developed not to cause cell membrane hydrolysis and necrosis so that they can deliver the therapeutic agents to the target site [29]. Although the benefits of nanocarriers in drug delivery have attracted much attention and great efforts have been made to investigate better cationic carriers, toxicity has always been the main problem of cationic carrier applications [30]. Because of this toxicity issue, the number of positive charges of polymer carriers should be kept low. In that case, the required therapeutic concentration of siRNA could not be achieved. Thus, N/P ratio has a great importance for gene delivery. Consequently, it is essential to keep the N/P ratio low, which indicates the complexation efficiency of the anionic therapeutic agents and the cationic carriers, in order to prevent cytotoxicity from excess amount of cationic polymer carriers. As a result, the main aim for optimum N/P ratio is to carry the most efficient number of siRNA with minimum number of cationic polymer carriers.

To identify the optimum ratio of N and P, the gel retardation assay commonly used for nucleic acid separation could be chosen for determination of N/P ratio [31]. This technique is frequently utilized for the determination of DNA or RNA fragments based on their molecular weight [32]. The phosphate backbone of the DNA or RNA is negatively charged, and therefore RNA fragments migrate to the positively charged anode when placed in the electric field for separation, identification, and purification [33].

Agarose which, is a pure linear polymer obtained from seaweed, is frequently used for gel electrophoresis [34]. The polymer is boiled to dissolve in a buffer solution and polymerized in gel form by hydrogen bonding when left to cool at room temperature. There is no other component such as catalysts required than agarose. Therefore, preparing agarose gel is simple and fast. The advantages of agarose gel electrophoresis are being non-toxic gel medium, rapid, and easy to cast of gels and providing well separation of high molecular weight nucleic acids [35]. In addition, the samples can be recovered from the gel by melting or digesting the gel with any agarose enzyme or by treating a chaotropic salt [36].

The movement of molecules in an agarose gel depends on their size, charge, the type of electrophoresis buffer, and the pore size of the gel. In this method, siRNA is forced to migrate through an extremely cross-linked agarose base in response to an electric field. In the solution, the phosphate groups of the siRNA are negatively charged so the siRNA molecule migrates to the positive pole (Fig. 1).

This technique is also being used for determination of the complexation efficiency of siRNA/cationic polymer carriers. By adding a positively charged polymer to the siRNA, the overall charge of the complex is neutralized. Because of decreasing the negative charge density of siRNA complex, the movement of siRNA/cationic polymer carriers in the gel is getting difficult. If the complexation does not happen completely, for example, there are free forms of siRNA with siRNA/cationic polymer carriers, the free siRNAs travel far away than the complexation forms [37].

All in all, the movement of a siRNA/cationic polymer carrier complexation through a gel depends on (a) size of the complexation structure, (b) agarose concentration, (c) type of agarose, (d) applied voltage, (e) presence of staining dye, and (f) electrophoresis buffer type [38]. After running the samples in a suitable dye-containing gel, the complexed and free siRNA can be visualized under UV light.

#### 2 Materials


Fig. 1 Agarose gel retardation assay to evaluate siRNA binding efficiency of 15 kDa cationic polymer complexation with siRNA at different N/P (amine/ phosphate) ratios. The gel retardation assay is set as the number of carriers is increasing, siRNA is kept constant. The orange-colored rectangle marking in the figure shows the optimal N/P ratio (2/1) which is siRNA completely complexed with the carriers


Prepare solutions with distilled water and nuclease-free water. Store all reagents (except siRNA and loading dye) at room temperature.

	- 2. Preparation of stock solution 5.0 l 50 TAE buffer (pH 8.0): Weight 1.21 g of Tris base, 67.24 g of EDTA, and draw 285.95 mL of acetic acid and dissolve all of them in 5.0 L distilled water, carefully. Perform this step on the magnetic stirrer and adjust pH 8.0 with hydrochloric acid (HCl) (see Note 2). Store the solution at room temperature.


4. Preparation of 1% agarose gel: Weight 1.0 g agarose powder, add 100.0 mL of electrophoresis buffer. Agarose powder is dissolved in the electrophoresis buffer to the desired concentration (see Note 4).

#### 1. Preparation of polymer solution: Add 1.0 mg of polymer in 1.0 mL of nuclease-free water. Store at 4 C.

2. Stock siRNA solution: Dissolve siRNA in nuclease-free water at a concentration of 50 μmol. Afterwards, allocate 800.0 μL of stock siRNA solution into microcentrifuge tubes (see Note 5) and store at 20 C.

#### 3 Methods

2.2 Polymer and siRNA



Table 1 The siRNA/cationic polymer carrier complexation for different N/P ratios

Fig. 2 Brief summary of the creation stages of gel retardation assay



#### 4 Notes


3.3 Loading siRNA/ Cationic Polymer Carrier Complex into 1% Agarose Gel


#### Acknowledgments

This work was supported by Scientific and Technological Research Council of Turkey (TUBITAK)-2515 COST Program Grant number: 118Z952 and Research Fund of Erciyes University (Project number: MAP-2020-9692).

#### References


resistance. Eur J Pharm Biopharm 136:18–28. https://doi.org/10.1016/j.ejpb.2019.01.006


25(2):237–253. https://doi.org/10.1038/cr. 2015.9


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Generation of Protein-Phosphorodiamidate Morpholino Oligomer Conjugates for Efficient Cellular Delivery via Anthrax Protective Antigen

### Valentina Palacio-Castan˜ eda, Roland Brock, and Wouter P. R. Verdurmen

#### Abstract

Phosphorodiamidate morpholino oligomers (PMOs) offer great promise as therapeutic agents for translation blocking or splice modulation due to their high stability and affinity for target sequences. However, in spite of their neutral charge as compared to natural oligonucleotides or phosphorothioate analogs, they still show little permeability for cellular membranes, highlighting the need for effective cytosolic delivery strategies. In addition, the implementation of strategies for efficient cellular targeting is highly desirable to minimize side effects and maximize the drug dose at its site of action. Anthrax toxin is a three-protein toxin of which the pore-forming protein anthrax protective antigen (PA) can be redirected to a receptor of choice and lethal factor (LF), one of the two substrate proteins, can be coupled to various cargoes for efficient cytosolic cargo delivery. In this protocol, we describe the steps to produce the proteins and protein conjugates required for cytosolic delivery of PMOs through the cation-selective pore generated by anthrax protective antigen. The method relies on the introduction of a unique cysteine at the C-terminal end of a truncated LF (aa 1–254), high-yield expression of the (truncated) toxin proteins in E. coli, functionalization of a PMO with a maleimide group and coupling of the maleimide-functionalized PMO to the unique cysteine on LF by maleimide-thiol conjugation chemistry. Through co-administration of PA with LF-PMO conjugates, an efficient cytosolic delivery of PMOs can be obtained.

Key words Antisense, Anthrax toxin, Protective antigen, Phosphorodiamidate morpholino oligomers, DNA analog, Drug delivery, Cellular internalization, Bioconjugate chemistry

#### 1 Introduction

Phosphorodiamidate morpholino oligomers (PMOs) are uncharged DNA analogs with therapeutic potential due to their ability to specifically bind to target sites on RNA. By steric inhibition of translation initiation complexes, PMOs can block translation. Alternatively, by targeting sites associated with splicing of pre-mRNAs, PMOs can mediate splice modulation and thereby correct the consequences of splicing mutations at the pre-mRNA level, for instance those in inherited retinal dystrophies [1]. PMOs

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_8, © The Editor(s) (if applicable) and The Author(s) 2022

have several qualities that are excellent for therapeutic development, including nuclease-resistance, long-term activity, low toxicity, and high specificity [2, 3]. However, a major challenge remains, which is achieving an efficient cellular delivery, particularly in vivo. PMOs are neutral molecules that because of their size are impermeable to cellular membranes. Delivery approaches that have been developed up to now include scraping of cells, particle-based approaches, and cell-penetrating peptide (CPP)-based delivery [3–5]. Cell scraping cannot be translated to in vivo applications and particle-based approaches suffer from delivery challenges in vivo such as poor tissue penetration and liver enrichment [6]. CPP-mediated delivery has demonstrated potential, but still does not target specific cell-surface receptors, indicating the need for a novel approach.

Recently, several groups have demonstrated that anthrax toxin, a sophisticated protein-based molecular machine that has evolved to efficiently deliver toxic catalytic proteins into the cytosol, can be employed for the functional delivery of various types of cargoes, including antisense oligonucleotides (AON) [7, 8]. The full anthrax toxin consists of three proteins: a pore-forming protein, called protective antigen (PA), that generates cation-selective pores and two enzyme components [9], called lethal factor (LF) and edema factor (EF). LF and EF in turn consist of two domains: the first domain binds the protein pore and the second domain is the enzymatically active domain and is thus responsible for the actual toxicity. For the delivery of PMOs via this mechanism, only PA and the PA pore-binding domain of LF are needed as protein components. For LF, this means that the enzymatic (toxicity-causing) domain of LF is replaced with a PMO.

In this chapter, we describe the preparation of the components needed to mediate cytosolic delivery of PMOs by the anthrax toxin translocation mechanism (Fig. 1a). The individual protein components, PA and LF (1–254) are produced in high quantity in soluble form in the cytosol of E. coli. LF (1–254) is by itself cysteine-free, so through the introduction of a unique C-terminal cysteine, a sitespecific conjugation via maleimide-thiol conjugation chemistry can be achieved. Maleimide-thiol conjugation chemistry is useful for coupling biologically active molecules because it is fast, highly selective and it can be done in physiological buffers at 37 C or at 4 C [10].

To enable the conjugation of the PMO to the protein, PMOs containing a primary amine at the 3<sup>0</sup> end are functionalized with a maleimide moiety through a bifunctional linker containing an NHS-ester and a maleimide group separated by a cyclohexane spacer. After coupling maleimide-functionalized PMOs with LF, uncoupled PMO is removed via dialysis, producing LF-PMO conjugates that can be delivered to the cytosol via anthrax protective antigen (Fig. 1b).

Fig. 1 Cytosolic delivery of PMO using the anthrax toxin mechanism. (a) Schematic representation of the cytosolic delivery of the LF-PMO conjugate via anthrax protective antigen. Numbers indicate the distinct steps in the delivery process. (b) Schematic representation of the coupling of a PMO to LF-cys. LF lethal factor, PA protective antigen, PMO phosphorodiamidate morpholino oligomer, SMCC succinimidyl 4-(N-maleimidomethyl)cyclohexane-1-carboxylate, TCEP tris(2-carboxyethyl)phosphine

#### 2 Materials

All buffers should be prepared with double-distilled water to ensure highly pure buffers. It is not necessary to work under sterile conditions. However, to perform experiments in the absence of antibiotics, make sure that the final conjugates are filter-sterilized before use.

#### 2.1 Protein Expression


#### 2.2 Protein Purification


#### 2.3 Protein-PMO Conjugation


#### 3 Methods


heat or formation of foam (see Note 8).

Fig. 2 TEV cleavage of MBP-LF and time dependency of coupling efficiency of LF to PMO. (a) SDS-PAGE gel showing LF fused to MBP before TEV cleavage (red arrow) and LF after cleavage and purification via reverse IMAC (orange arrow). (b) SDS-PAGE gel illustrating the effect of incubation time on the conjugation efficiency of coupling LF to the PMO. Red arrow indicates 86 kDa band corresponding to MBP-LF before coupling. Green arrow shows a band with increased molecular weight at approximately 92 kDa corresponding to the coupled fraction (MW of PMO: 6.8 kDa). No marked differences are seen between the incubation time and the coupling efficiency, indicating that 4 h incubation is sufficient to achieve 50% coupling. Proteins on gels were visualized by stain-free imaging technology (Bio-Rad). IMAC immobilized metal ion affinity chromatography, LF lethal factor, MBP maltose-binding protein, PMO phosphorodiamidate morpholino oligomer, TEV tobacco etch virus


3.3 Purification of PA by Size-Exclusion Chromatography (SEC) After purification of PA via IMAC, we typically see some co-purification of degraded fragments, which can be removed via SEC (see Note 11). We generally purify our PA or PA fusion constructs via SEC using an A¨ kta pure chromatography system equipped with an S200 column (see Note 12), which yields pure and active proteins [11, 12].

> 1. Concentrate the sample to a volume of less than 500 μL, which is the maximal volume that can be loaded on an S200 column. Make sure that the protein solution is sufficiently concentrated so that in the maximally 500 μL several milligrams of proteins can be loaded (ideally 2–5 mg). For lower amounts, the separation of individual peaks can become problematic. Filtration of the protein using a low protein-binding syringe filter (0.22 μm pore size) before loading is recommended since protein

aggregates may clog the column. Alternatively, centrifugation of the sample at high speed (~20,000 g) for 5 min and loading the supernatant can be done.

	- 2. Calculate the amount needed of the SMCC linker for the PMO coupling, taking into account that a 20-fold molar excess of the linker is needed for the reaction. Weigh the desired amount of the SMCC linker and dissolve in anhydrous DMF (e.g. 1.5 mg SMCC in 100 μL of DMF to make a stock solution of 45 mM). Freeze aliquots at 20 C and only thaw briefly for use. The NHS moiety of the linker is very reactive and may hydrolyze already through the presence of trace amounts of water.
	- 3. Mix the PMO with a 20-fold molar excess of the SMCC linker with minimal dilution (e.g. 900 μM PMO and 18 mM SMCC linker) and incubate for 2 h at 4 C (see Note 14).
	- 4. Quench unreacted linker by adding lysine to a final concentration of 100 mM using a 1 M lysine solution.

#### 3.4 Functionalizing PMO with a Maleimide Moiety

	- 2. Dialyze overnight to remove the unreacted PMO. Exchange dialysis buffer the next morning and dialyze for two more hours. Upon complete removal of the excess uncoupled PMO, the concentration of the PMO-conjugate can be calculated using the absorption coefficient of the PMO at 265 nm, while correcting for the absorption of the protein at 265 nm (see Note 16). The conjugates can be snap-frozen in liquid nitrogen and stored at 80 C.

#### 4 Notes


3.5 Coupling of Anthrax Lethal Factor to Maleimide-Functionalized PMO

K2HPO4. Adjust volume to 1 L. Mixing the separately autoclaved solutions can be done at the day of the experiment.


#### Acknowledgments

This work was supported by funding from Landelijke Stichting voor Blinden en Slechtzienden, grant number: UitZicht 2017-14.

#### References


properties. Antisense Nucleic Acid Drug Dev 7(3):187–195

3. Shabanpoor F, McClorey G, Saleh AF et al (2015) Bi-specific splice-switching PMO oligonucleotides conjugated via a single peptide active in a mouse model of Duchenne muscular dystrophy. Nucleic Acids Res 43(1):29–39


antigen. Biochem Biophys Res Commun 376(1):200–205


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part III

In Vitro Model Systems

# Chapter 9

# Development and Use of Cellular Systems to Assess and Correct Splicing Defects

### Nuria Sua´rez-Herrera, Tomasz Z. Tomkiewicz, Alejandro Garanto, and Rob W. J. Collin

#### Abstract

A significant proportion of mutations underlying genetic disorders affect pre-mRNA splicing, generally causing partial or total skipping of exons, and/or inclusion of pseudoexons. These changes often lead to the formation of aberrant transcripts that can induce nonsense-mediated decay, and a subsequent lack of functional protein. For some genetic disorders, including inherited retinal diseases (IRDs), reproducing splicing dynamics in vitro is a challenge due to the specific environment provided by, e.g. the retinal tissue, cells of which cannot be easily obtained and/or cultured. Here, we describe how to engineer splicing vectors, validate the reliability and reproducibility of alternative cellular systems, assess pre-mRNA splicing defects involved in IRD, and finally correct those by using antisense oligonucleotide-based strategies.

Key words ABCA4, Antisense oligonucleotide, Exon skipping, Genetic therapy, Inherited retinal diseases, Maxigene, Midigene, Pre-mRNA, Pseudoexon, Splicing modulation, Splicing vectors

#### 1 Introduction

Technologies such as next generation sequencing (NGS) expanded the discovery of genetic variants from coding regions to the entire genome. As a consequence of high-throughput data analysis, it is crucial to be able to correctly identify and distinguish diseasecausing variants from single nucleotide polymorphisms (SNPs). This is especially relevant for intronic variants, as many of them have an unknown functional significance. In the field of inherited retinal diseases (IRDs), a common autosomal recessive condition known as Stargardt disease (STDG1) [1] lacks the bi-allelic molecular diagnosis in 30% of cases, i.e. the second variant cannot be identified in the coding regions of adenosine triphosphate (ATP) binding cassette type A4 (ABCA4) gene [2, 3].

Nuria Sua´rez-Herrera and Tomasz Z. Tomkiewicz contributed equally to this work.

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_9, © The Editor(s) (if applicable) and The Author(s) 2022

According to the Human Gene Mutation Database, mis-splicing mutations have been estimated to represent 8.6% of the total mutations underlying inherited diseases (23,868/ 275,716) [4]. In vitro functional assays can provide insight into the underlying mechanisms behind aberrant splicing and identify the mutations that interfere with this process. Currently, the ideal model to assess and correct mis-splicing mutations in STGD1 are iPSC-derived retina-like cells (such as photoreceptor precursor cells (PPCs) or retinal organoids) from a patient harboring the genetic variants of interest, as these represent the right cell type(s) with the proper genetic context. In parallel, great efforts have been made to develop a more cost-effective and less time-consuming strategy to reach the same goal, by trying to mimic the pathological situation in a reliable and controlled manner. Engineering and use of multiexon splice vectors have been proven to be extremely effective [2, 5–7] in gaining insight into IRDs and, more specifically, STDG1. In general, splice vectors or midigenes refer to a specific type of vector containing a large genomic region that allows to study the splicing processes between the included exons. An even longer genomic content allows for inclusion of long-range cis-acting elements to more accurately reflect the dynamics of splicing [8]. These "artificial" genomic vectors were shown to be very valuable when it comes to ABCA4, as the entire 128-kb gene has been successfully spanned in a set of midigenes representing an alternative to the impossibility of cloning such a large genomic region in a single vector [6].

The suitability of midigenes for some cell lines reduced the complexity of the study of pathologic deep-intronic variants (among other types of mutations) and their effect on splicing. In addition, validated midigenes harboring splicing variants represent a system for reliable and relatively quick identification of potential therapeutic molecules such as antisense oligonucleotides (AONs). AON-based therapies represent a very effective approach to target mis-splicing mutations. AONs are chemically modified RNA molecules that have the ability of modulating splicing by binding to premRNA and interfering with the spliceosome [9]. They are used in the field of IRDs [10], as well as in other genetic diseases as the purpose of AONs is not limited to splice-switch function only [11]. Generating a reliable artificial splicing system is of major importance when it comes to the development of a potential therapeutic molecule. For the early investigation of possible causative variant affecting pre-mRNA splicing in retinal genes and subsequent assessment of AON potency, midigene technology is a suitable approach that has been shown to produce reliable results [6, 12].

Following transfection of the vector into HEK293T cells, AONs are co-transfected, and the splicing correction can be assessed at RNA level. Subsequently, the AONs that are identified as most potent can be tested in more advanced and precise cell models such as iPSC-derived retina-like cells.

Despite the proven efficiency of the HEK293T cells, they are not derived from ocular tissue, which allows to speculate whether the splicing dynamics enforced by HEK293T represents that of the retina. As an alternative, with the aim to better represent the retinalike splicing dynamics in in vitro splice assays, we also describe the use of retinoblastoma WERI-Rb-1 cells. These cells are also suitable for midigene transfection and in some cases demonstrate splicing dynamics that are more similar to that of the retina when compared to HEK293T cells.

In this chapter, we describe how to design multi-exon splice vectors followed by appropriate validation and correction of mis-splicing mutations by performing in vitro studies in cellular systems.

#### 2 Materials

2.1 Design of Midigene and Maxigene Splice Vector


1. Generated midigene and maxigene vectors (see Subheadings 3.1.1 and 3.2.2).

2. Primers to introduce the desired mutation, in this chapter we use the c.859-506G > C mutation in the ABCA4 midigene as an example (the variant is in bold and underlined):

Forward primer 5<sup>0</sup> - CTGTGATTTGTTGTTGTTGTTG TTGTTGTTTT G AGACGGAGTAT -TGCTCAG-3<sup>0</sup> and reverse primer 5<sup>0</sup> - GACACTAAACAACAACAACAACAACAA CAA-AACTCTGCCTCATAACGAGTC-3<sup>0</sup> [3].


2.2 Site-Directed Mutagenesis


2.3 Culture Conditions and Cell Lines

	- 2. WERI-Rb-1 cells (ATCC® HTB-169™). Culture medium: DMEM 15% FCS medium (DMEM medium supplemented with 15% FCS, 100 U/mL of penicillin, 100 μg/mL streptomycin and 10 mL of 1 M HEPES).
	- 3. T75 flasks to culture cell lines.
	- 4. 0.25% trypsin solution for cell dissociation.
	- 5. 1-PBS.
	- 2. Midigene vector, as an example we used ABCA4 BA7 midigene [6].
	- 3. AON stock: resuspend the lyophilized AONs at final concentration of 0.1–1 mM in 1-PBS previously autoclaved twice.
	- 4. 6-well plates and 24-well plates.
	- 5. OptiMEM and transfection reagents (i.e., FuGene® or Lipofectamine®).
	- 6. 0.25% trypsin solution for cell dissociation.
	- 7. 1-PBS.

#### 2.5 RT-PCR 1. Commercially available RNA isolation kit.

	- (a) Region of interest.
		- <sup>l</sup> ABCA4 exon 7 forward: 5<sup>0</sup> - TCTGAGATCTTGGG GAGGAA-30 .
		- <sup>l</sup> ABCA4 exon 8 reverse: 5<sup>0</sup> - TGGAGTCAATCCCCA GAAAG-30 .
	- (b) Actin loading control (see Note 2).
	- <sup>l</sup> RHO exon 5 forward: 5<sup>0</sup> - ATCTGCTGCGGCAA GAAC-30 .
	- <sup>l</sup> RHO exon 5 reverse: 5<sup>0</sup> -AGGTGTAGGGGATGGGA GAC-30 .

#### 3 Methods

Retina-specific genes are not readily expressed outside the ocular tissue. The inability to express or poorly express the genes of interest in non-ocular tissues makes it difficult to study variants affecting pre-mRNA splicing. Generation of PPCs from patientderived reprogramed fibroblasts is an alternative to this, but is timeconsuming and expensive. 3.1 Design of Midigene Splice Vectors 3.1.1 Gateway Cloning In here, we describe the generation and use of pCI-NEO-RHO Gateway-adapted in-house vector (see Fig. 1). 1. Identify the gene and sequence of interest in genomic databases such as Ensembl Genome Browser or UCSC [13, 14] and then identify the region of interest. In this case, the region of interest is a deep-intronic variant causing guanine to cytosine substitution at position c.859-506 of the ABCA4 gene. The base substitution strenghtes a cryptic deep-intronic splice acceptor side and causes generation of a 56-nts long pseudoexon between exon 7 and 8. 2. Design primers suitable for Gateway® BP cloning (see Note 3). 3. Use the BAC clone, CH17-325O16 (insert g.94,434, 639–94,670,492), containing the entire ABCA4 gene. Isolate the BAC DNA using commercially available midiprep kit and use it as a PCR template to generatet the midigenes. Prepare the PCR reaction with 0.5 μM of each forward and reverse primer, 0.2 mM dNTPs, Phusion High-Fidelity DNA polymerase, 1- Phusion GC buffer, 3% DMSO, and 2.5 ng of BAC DNA in a total of 50 μL. Run the PCR program where the initial denaturation is at 98 C for 30 s; 15–20 cycles of denaturation at 98 C for 10 s each, annealing at 58 C and

Fig. 1 Schematic representation of wild-type and mutant ABCA4 midigene engineering. The simplified protocol for Gateway® system cloning and site-directed mutagenesis are shown on the left and right section, respectively. SDM: site-directed mutagenesis

extension at 72 C for at least 1 min per kb of insert, with a final extension at 72 C for 15 min [6].


3.1.2 Side-Directed Mutagenesis



In this section, we describe the side-directed mutagenesis protocol for midigene constructs:


3.2 Design of Maxigene Splice Vectors There are some variants that may need larger genomic environment in order to correctly assess them. This could be the case for mutations whose effect is only detected when other splice regulatory motifs are present, generally located in introns. As a consequence, larger genomic context should be included and based on our experience, it is difficult to obtain a splicing vector of this size because of several reasons. The first one is the probability of inducing single nucleotide changes during the amplification of the genomic region of interest. Long-range PCR and high-fidelity DNA Taq polymerases may prevent this from happening, although these chances increase when amplifying >10 kb fragments. Recombination efficiency between the fragment and the donor vector is also affected, as well as the efficiency of site-directed mutagenesis on the entry clone. Both cases are highly associated with the size of the vector. To overcome these limitations, we propose to use the already engineered midigenes completely covering the whole gene in order to generate a maxigene, which combines the genomic context of more than one midigene. In this section, we show an example of the mentioned cloning strategy.

3.2.1 In Silico Design of Maxigene Strategy 1. Select the two midigenes that include the introns and exons of interest.

3.2.2 Cloning of Maxigene Vectors


Fig. 2 Schematic representation of wild-type and mutant maxigene engineering. Cloning steps and sitedirected mutagenesis are shown on the left and right section, respectively. This example of maxigene strategy is based on the alternative procedures explained in Subheading 3.2. SDM: site-directed mutagenesis


3.2.3 Site-Directed Mutagenesis To obtain the mutant maxigene, you can either use midigene 1 or 2 containing the mutation of interest and follow the same protocol indicated in the previous section or perform the site-directed mutagenesis on the wild-type maxigene. In order to perform the second option, we propose the following strategy as an example for a maxigene (see Fig. 2):


#### 3.3 In Vitro Evaluation of Splice Vectors in Cell Lines

3.3.1 Transfection in

HEK293T

The constructed midi/maxigenes need to be validated and assessed in vitro. In the functional studies of IRDs, the cell line of choice is usually HEK293T. These cells are easy to transfect and do not express retina-specific genes. Therefore, the expression of retinaspecific gene delivered with the vector is not interfered by the endogenous expression of such gene. This advantage is counterbalanced by fact that the HEK293T cells have different properties compared to retina cells. As a consequence, the splicing dynamics are often but not always the same. As an alternative to better represent the retina-like splicing dynamics, we also describe the use of WERI-Rb-1 cells, which are retinoblastoma cells. These cells are still suitable for midigene transfection and sometimes mimic retina-specific splicing patterns better in comparison with HEK293T cells (see Fig. 3).

	- 2. Once the cells are attached, transfect 1.2 μg of plasmid in a 6-well plate. In this example we used FuGENE (ratio is 3:1 FuGENE:plasmid). To make transfection mix, to 200 μL of OptiMEM medium, add 3.6 μL of FuGENE. To the transfection mix add 1.2 μg of plasmid and incubate at RT for 15–20 min.
	- 3. Dispense the transfection mix on top of the corresponding well (see Note 12).
	- 4. Incubate the plate at 37 C for 24 (see Subheading 3.4.1) or 48 h (see Note 13).
	- 5. After the incubation, gently rinse the cells with pre-warmed 1- PBS and then detach the cells using 500 μL trypsin. Collect the detached cells into a 1.5 mL Eppendorf using P1000 pipette.
	- 6. Centrifuge the Eppendorf tubes for 5 min at 1000 g to pellet the cells, remove the supernatant and wash the cells again in 1- PBS for 5 min at 1000 g.
	- 7. Discard the supernatant and:
		- (a) freeze the cell pellets at 80 C (storage) after 48 h incubation and proceed with further analysis at another moment or,
		- (b) proceed with the RNA isolation (validation of the midigene expression and splicing) after 48 h incubation.
	- 1. Dispense 1 to 2 - 10<sup>6</sup> cells to 1.5 mL Eppendorf tube in a total volume of 500 μL of DMEM 15% FCS.
	- 2. Transfect 1.2 μg of plasmid. In this example, we used FuGENE (ratio is 3:1 FuGENE:plasmid). To make transfection mix

3.3.2 Transfection in WERI-Rb-1

Fig. 3 Experimental strategy to assess splicing defects in HEK293T or WERI-Rb-1 cell lines. The cells are first transfected with a wild-type (WT) or mutant (MUT) ABCA4 midigene and, following a 48-h incubation, splicing is validated by RT-PCR as outlined in Subheading 3.4.3 The gel picture shows an approximate read-out of the expected ABCA4 transcripts. MQ negative control of the PCR reaction, EMP empty transfection mix (endogenous expression of the selected genes within the cell line used)

200 μL of OptiMEM medium with 3.6 μL of FuGENE. And to this tube, add 1.2 μg of plasmid and incubate at RT for 15–20 min.

	- (a) freeze the cell pellets at 80 C (storage) after 48 h incubation and proceed with further analysis at another moment or,
	- (b) proceed immediately with the RNA isolation (validation of the midigene expression and splicing) after 48 h incubation.

3.3.3 Validation of Splicing Events by Reverse Transcriptase PCR (RT-PCR)

Before starting to use midigenes as an artificial system, it is necessary to check the effect of the variant at pre-mRNA level, and therefore, the RT-PCR is used as a validation method for splicing events.

1. Design forward and reverse primers to flank the region of interest. If the cell line used presents endogenous expression of your target gene, design one of the primers in the artificial exon of the midigene (RHO) (see Note 14).


In this part of the chapter, we describe the experimental design to correctly compare a set of AON sequences by using midigenes as a model system and how to determine the efficacy of aberrant splicing correction. The utilization of photoreceptor precursor cells and/or retinal organoids is a recommended step in the final stages of AON potency validation in in vitro studies.

3.4.1 Midigene Vector and AON Co-transfection Seeding 0.5 - 106 cells/well in 6-well plate provides enough cells to seed 6 wells on the 24-well plate. It is recommended to perform early calculations into how many wells will be required for the splice correction assay as this will influence the number of wells on the 6-well plate required for midigene transfection. If 6 or less wells on the 24-well plate are needed, one 6-well plate seeded with 0.5 - 106 HEK293T cells and transfected with midigene is enough.

When setting up splice assay with AONs, do not forget about the minimum required controls. These include non-transfected control or empty transfection mix (EMP), cells transfected with

3.4 Correcting Splicing Defects in Artificial Systems

Fig. 4 Analysis of AON-mediated splicing correction in midigene-based splice assays. (a) Experimental strategy to correct splicing defects by co-transfection of wild-type (WT) or mutant (MUT) ABCA4 midigenes with AONs in cell lines as described in Subheading 3.4.1. (b) Representation of splicing read-out on agarose gel and further semi-quantitative analysis. The gel picture represents an approximation of expected ABCA4 transcripts, further semi-quantitative analysis and graphs are based on this estimation. MQ: negative control of the PCR reaction, EMP: empty transfection mix (endogenous expression of the selected genes within the cell line used), NT: non-treated (transfected with midigene but no AONs), SON: scrambled or sense oligonucleotide

wild-type (WT) midigene or mutant (MUT) midigene but not transfected with AONs (NT), and cells transfected with wild-type or mutant midigene and co-transfected with the corresponding scrambled or sense oligonucleotide (SON). The controls cells have to be seeded together with test cells and exposed to the exact same treatment conditions, which include medium change and transfer from 6-well plate to 24-well plate.


Image J

100 μM stock concentration. Incubate the transfection reaction for 15–20 min at RT. After incubation add enough medium to make 1 mL.

	- (a) freeze the cell pellets at 80 C (storage) and proceed with further analysis at another moment or,
	- (b) proceed immediately with the RNA isolation and the cDNA synthesis following manufacturer's instructions.

3.4.2 Quantification of Splicing Redirection with Measuring efficiency of an AON-based strategy represents the last step in order to discard or select a potential therapeutic molecule for further clinical studies. There are several methods available to calculate this efficiency and, in this chapter, we focus on a semiquantification strategy for RT-PCR readouts (see Fig. 4b).


% of Transcript <sup>¼</sup> Area Transcript Total Area -100


3.4.3 How to Know If an AON Is Effective? Once you assess the aberrant transcript rescue of the different AONs that are being tested, it is possible to classify them in several groups depending on their splicing redirection efficiency (see Note 24). In previous studies [15], we classified AONs in the following five groups in order to analyze their properties in a straightforward way (see Fig. 4b): highly effective (>75% correction), effective (between 75% and 50% correction), moderately effective (between 50% and 25% correction), poorly effective (between 25% and 0% correction), and noneffective (when correction is not detected).

However, there are some limitations as this protocol is based on a semi-quantitative strategy, such as the interference of heteroduplexes and midigene artifacts. Even though we presented different ways of overcoming these limitations, there is the possibility of using other quantitative strategies [16]. As an example, Fragment Analyzer, TapeStation or digital droplet PCR can be implemented in order to get more accurate splicing readouts (see Note 25).

#### 4 Notes


incubation with phosphatase is needed at least in one of the digested midigenes and screening of colonies afterwards might be more extensive. In our case, phosphatase incubation was performed in digestion product from midigene 2 to avoid re-ligation with itself.


#### Acknowledgments

This work was supported by European Union's Horizon 2020— Marie Sklodowska-Curie Actions grant no. 813490 (to R.W.J.C.) and Retina UK Foundation grant no. GR596 (to R.W.J.C.), Algemene Nederlandse Vereniging ter Voorkoming van Blindheid, Stichting Blinden-Penning, Landelijke Stichting voor Blinden en Slechtzienden, Stichting Oogfonds Nederland, Stichting Macula Degeneratie Fonds, and Stichting Retina Nederland Fonds (who contributed through UitZicht 2015-31 and 2018-21), together with the Rotterdamse Stichting Blindenbelangen, Stichting Blindenhulp, Stichting tot Verbetering van het Lot der Blinden, Stichting voor Ooglijders, and Stichting Dowilvo (to A.G. and R.W.J. C.). This work was also supported by the Foundation Fighting Blindness USA, grant no. PPA-0517-0717-RAD (to A.G. and R. W.J.C.). The funding organizations had no role in the design or conduct of this research. They provided unrestricted grants.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Modeling Splicing Variants Amenable to Antisense Therapy by Use of CRISPR-Cas9-Based Gene Editing in HepG2 Cells

### Arı´stides Lo´ pez-Ma´rquez, Ainhoa Martı´nez-Pizarro, Bele´n Pe´rez, Eva Richard, and Lourdes R. Desviat

#### Abstract

The field of splice modulating RNA therapy has gained new momentum with FDA approved antisensebased drugs for several rare diseases. In vitro splicing assays with minigenes or patient-derived cells are commonly employed for initial preclinical testing of antisense oligonucleotides aiming to modulate splicing. However, minigenes do not include the full genomic context of the exons under study and patients' samples are not always available, especially if the gene is expressed solely in certain tissues (e.g. liver or brain). This is the case for specific inherited metabolic diseases such as phenylketonuria (PKU) caused by mutations in the liver-expressed PAH gene.

Herein we describe the generation of mutation-specific hepatic cellular models of PKU using CRISPR/ Cas9 system, which is a versatile and easy-to-use gene editing tool. We describe in detail the selection of the appropriate cell line, guidelines for design of RNA guides and donor templates, transfection procedures and growth and selection of single-cell colonies with the desired variant, which should result in the accurate recapitulation of the splicing defect.

Key words Splicing, Gene editing, CRISPR/Cas9, HepG2, Inherited metabolic diseases, Phenylketonuria, Cellular models

#### 1 Introduction

Splicing defects account for up to one-third of human diseasecausing variants, according to the current estimates [1–3]. Constitutive splicing relies on the recognition of consensus splicing sequences (5<sup>0</sup> splice site, 3<sup>0</sup> splice site, branch point, and polypyrimidine tract) by spliceosomal components, as well as of other less conserved regulatory elements, referred to as exonic or intronic splicing enhancers or silencers (ESE, ISE, ESS, or ISS), that modulate spliceosome recruitment [4]. These cis-regulatory elements are recognized by trans-acting factors including the serine/argininerich domain-containing (SR) protein and heterogeneous nuclear

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_10, © The Editor(s) (if applicable) and The Author(s) 2022

ribonucleoprotein (hnRNP) families that may act co-ordinately to accurately regulate exon inclusion.

Pathogenic splicing variants disrupt conserved splice sites or regulatory elements or cause aberrant splicing by creating/activating alternative splice sites or by promoting the aberrant inclusion of intronic pseudoexons [4]. Splicing can be modulated therapeutically using antisense approaches, and to date, the clinically approved splice-switching antisense oligonucleotides (SSO) for spinal muscular atrophy, Duchenne muscular atrophy and for an individual patient with a rare, fatal neurodegenerative disease [5–7], represent landmarks in the field, opening new avenues for treatment of patients with defects amenable to splice-mediated correction.

The first requirement for the accurate design and testing of antisense splice correction therapy is the availability of relevant experimental models in which to dissect the underlying molecular mechanisms of pathogenic variants and to test candidate molecules. In this sense, the development of clustered-regulatory interspaced short palindromic repeats (CRISPR)-CRISPR associated nuclease (Cas) genome editing has paved the way to the rapid and easy generation of new and improved cell/animal models of disease. This has facilitated the understanding of the specific pathogenic effect and has allowed efficient testing of targeted therapies, including allele-specific repair for splicing mutations, in tissue types with native expression levels [8–13]. Based on a naturally employed bacterial defense mechanism [14, 15], CRISPR/Cas9 technology was developed as a programmable system of genetic editing that commonly uses the Cas9 nuclease from Streptococcus pyogenes and a RNA duplex comprised of a sequence-specific CRISPR RNA (crRNA) and a generic trans-activating CRISPR RNA (tracrRNA) that directs the nuclease to a cut site point, three base pairs upstream of the protospacer adjacent motif or PAM. The PAM is a three-nucleotide motif essential for the nuclease to recognize its DNA target which in the case of Cas9 is NGG. The crRNA and the tracrRNA can be delivered individually or linked in a single RNA molecule. These elements can be delivered to cells as plasmids or as a ribonucleoprotein (RNP) complex [16].

Once Cas9 nuclease cuts the DNA introducing a double stranded break (DSB), the cell can repair this through two different mechanisms: non-homologous end-joining (NHEJ) which usually results in small insertions or deletions, useful for the generation of gene knockouts, or homology driven repair (HDR), used to introduce specific changes via a DNA template with homology arms to our target locus and containing the sequence or point mutation desired [16].

In our laboratory we have used CRISPR/Cas9 technology to introduce splicing mutations causing inherited metabolic diseases (IMD) in cellular and animal models. IMD are monogenic diseases characterized by dysregulation of the metabolic networks that underlie development and homeostasis [17]. They belong to the category of rare diseases due to their low individual prevalence and are generally enzyme deficiencies of autosomal recessive inheritance, characterized by the toxic accumulation of precursors and of their derivatives or by lack of downstream metabolites. Several of the most frequent and well characterized IMD, e.g. organic acidemias and amino acid disorders, are of major hepatic expression and, as in other genetic diseases, 13–25% of all disease-causing variants interfere with mRNA splicing (HGMD statistics, Professional Release 2019.3). These data warrant further investigation of the therapeutical potential of SSOs in these diseases and the generation of liver specific cellular models for these studies.

The generation of a cell model using CRISPR/Cas9 system can be done in a huge variety of cell lines. In this chapter we describe the protocol for efficient introduction of a specific splicing variant in the PAH gene, coding for phenylalanine hydroxylase, and responsible for the well characterized disease phenylketonuria (PKU, MIM#261600), inherited in autosomal recessive fashion. Human PAH is exclusively expressed in liver, so in this protocol we use hepatoma cell line HepG2 seeking to attain edition in both alleles (homozygous phenotype). We explain how to select for the appropriate cell line in each particular case, describe the design of RNA guides and donor templates, transfection procedures, growth of single-cell colonies, selection and testing to confirm genomic edition, and accurate recapitulation of the splicing defect (Fig. 1). Appropriate controls to be included in each step are explained, as well as the necessary precautions to be taken especially for intronic splicing variants. We use as example the CRISPR/Cas9-mediated introduction of the recently characterized PAH intronic variant, c.1199 + 20G > C, that causes exon skipping due to disruption of a splicing regulatory element [18]. This variant creates a PshAI restriction site, which is used to screen for gene edition in the transfected cells.

#### 2 Materials

	- 2. Humid CO2 incubator.
	- 3. Centrifuge.
	- 4. Phase-contrast microscope.
	- 5. Hemocytometer–double chamber with Neubauer rulings.
	- 6. Manual Counter.
	- 7. Consumables: Tissue culture plates, filtered tips, falcon tubes, Eppendorf tubes.

Fig. 1 Outline of the gene editing experimental protocol. (This image was created using BioRender)




#### 2.8 Web Resources 1. Sequences and genomes: https://www.ensembl.org/.


#### 3 Methods


Fig. 2 Schematic representation of the PAH gene region surrounding the c.1199 + 20G > C mutation, showing the sequence of the ssDNA template which will include the desired change (green line with red box) and the crRNA guides used (purple arrows), indicating the corresponding PAM sequences (gray rectangles) and the Cas9 nuclease cut sites 3 nucleotides upstream of PAM (blue arrows)

	- 2. Prepare the RNA Duplex at a final concentration of 1 μM by mixing the tracrRNA and the crRNA in equimolar concentrations in Nuclease-Free Duplex Buffer (add 1 μL of each crRNA and tracRNA-ATTO550 to 98 μL of buffer) (see Note 20).
	- 3. Heat at 95 C for 5 min.
	- 4. Cool to room temperature (25 C).

#### 3.4 Preparation of the Ribonucleoprotein Complex (RNP)


3.5 Reverse Transfection of RNP and DNA Donor Template



3.7 Generation of the Single-Cell Colonies


3.8 Genomic DNA Extraction and RFLP Analysis

176 Arı´stides Lo´ pez-Ma´rquez et al.

Fig. 3 RFLP analysis to monitor for gene edition. The wild-type and mutant sequences are shown in panel <sup>a</sup> and panel <sup>b</sup> is a representative gel showing RFLP analysis of single-cell colonies. Top bands correspond to the amplified PCR products and lower-sized bands correspond to the products obtained by digestion with PshAI enzyme due to the introduction of a restriction site with the point mutation c.1199 + 20G > C. <sup>C</sup> undigested control. Colonies 1, 3, 4, 6, 8, 9, 11–15 are positive and heterozygous (one allele edited)

3.9 Sequencing Analysis of Candidate Clones and Off-Targets Analysis


#### 3.10 RNA Isolation 1. Once a correctly edited clone has been identified, expand the culture to obtain enough cells for RNA isolation.


#### 4 Notes


#### 3.11 RT-PCR and Sequencing Analysis to Confirm the Splicing Defect

Reagent (Thermofisher) has been the method and the reagent chosen is this protocol.


The election of one method or another will depend on the cell line, so it is advisable to test this before generating the colonies with the edited cells. In our hands, for example, HepG2 cells exhibited high mortality after sorting and plating in 96-well plates, so we selected option c. For some cell lines the use of conditioned medium (filtered culture medium collected from control cells) can aid the growth in the form of a colony derived from a single cell. The time of growth and appearance of single-cell colonies will depend on the type of cells you are working with. With HepG2 cells, colonies emerged and reached the correct size after circa 20 days.

6. It is advisable to have the region sequenced before starting the editing experiment to identify single-nucleotide polymorphisms in the specific cell line used which may affect the design of RNA guides and DNA templates, as well as result in erroneous interpretation of the sequencing analysis of the edited clones (concluding there has been an extra change introduced during DNA repair after Cas9 reaction when it was already present in the sequence prior to editing).


does not increase the homologous recombination success rate. However, for longer edits (e.g. insertion/deletion of several nucleotides), the length of the homology arms must also be increased.


only depend on the quality of the guide, but also on the transfection method, the locus we are editing, the cell type, etc.


well of the 24-well plate into two wells of a 12-well plate. It is important to keep accurate record of each duplicate, since one of them will be used to extract DNA for analysis, and the other will be used to freeze the colony and, in case it is the one selected, expand it for further characterization and use.


conclude that off-targets, although it is important to sequence and validate them, are not the biggest problem. However, we frequently found extra changes in the area near the edited nucleotide (on-target). In this sense, these errors have been the main problem and the cause of having had to discard many clones before finding the final positive one.


#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 11

# In Vitro Models for the Evaluation of Antisense Oligonucleotides in Skin

### Jeroen Bremer and Peter C. van den Akker

#### Abstract

The genodermatosis dystrophic epidermolysis bullosa (DEB) is caused by mutations in the COL7A1 gene which encodes type VII collagen (C7). In the cutaneous basement membrane zone, C7 secures attachment of the epidermal basal keratinocyte to the papillary dermis by means of anchoring fibril formation. The complete absence of these anchoring fibrils leads to severe blistering of skin and mucosa upon the slightest friction and early mortality. To date, although preclinical advances toward therapy are promising, treatment for the disease is merely symptomatic. Therefore, research into novel therapeutics is warranted.

Antisense oligonucleotide (ASO)-mediated exon skipping is such a therapy. Clinical examination of naturally occurring exon skipping suggested that this mechanism could most likely benefit the most severely affected patients. The severe form of DEB is caused by biallelic null mutations. Exon skipping aims to bind an ASO to the mutated exon of the pre-mRNA in the cell nucleus. Thereby, the ASO inhibits the recognition of the mutated exon by the splicing machinery, and as a result, the mutated exon is spliced out from the mRNA with its surrounding introns, i.e., it is skipped. Here, we describe in vitro methods to evaluate ASO-mediated exon skipping in a preclinical setting.

Key words Epidermolysis bullosa, Therapy, Exon skipping, Antisense RNA, Fibroblasts, Keratinocytes, Splice modulating

#### 1 Introduction

In this chapter, we describe the evaluation of ASO-mediated exon skipping as a therapeutic approach for DEB in a preclinical in vitro setting. DEB is caused by mutations in the COL7A1 gene which encodes type VII collagen (C7) [1]. DEB is a rare disease affecting 1–9 in every one million births, worldwide. The disease is characterized by severe blistering of skin and mucosae. DEB can be inherited both dominantly and recessively, and the severity of the disease strongly depends on the quantity and functionality of the C7 protein present at the cutaneous basement membrane zone. The most severe recessive form of DEB (RDEB-gen sev) is caused by biallelic null mutations and the complete absence of C7 in the skin. Previously, we have shown that for RDEB-gen sev caused by

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_11, © The Editor(s) (if applicable) and The Author(s) 2022

biallelic null mutations, exon skipping is anticipated to be clinically beneficial [2].

Exon skipping relies on specifically designed ASOs that bind to the pre-mRNA in the cell nucleus. When bound, these ASOs inhibit the recognition of the mutated exon by the splicing machinery through steric hindrance [3]. As a result, the mutated exon is spliced out (skipped) of the mRNA together with its surrounding introns. If the skipped exon is in frame, the reading frame of the transcript is maintained and produces a slightly shorter but functional protein [4].

Exon skipping affects the pre-mRNA; therefore, it is essential for the ASO to pass the cell membrane and the nuclear envelope. The commonly used 2<sup>0</sup> -O-methyl phosphorothioate (2OMePS) and 2<sup>0</sup> -methoxyethyl phosphorothioate (2MOE) ASOs are negatively charged and therefore not able to easily pass the cell membrane in cell cultures. Therefore, active transfer across the cell membrane is essential. Cationic lipid transfection is such a way of active transfer and widely used to achieve efficient uptake by in vitro cultured cells. Widely studied cells of the skin are dermal fibroblasts and epidermal keratinocytes. Here, we describe the in vitro evaluation of distribution and activity of ASOs in cultured fibroblasts and keratinocytes, as C7 is expressed by both the cell types. However, cationic lipid transfection of fibroblasts and keratinocytes can be used to evaluate the activity of antisense RNA for many diseases, as they express many proteins.

#### 2 Materials

	- 2. Dispase II (2.4 U/mL).
	- 3. Penicillin/Streptomycin (100 U/mL and 100 μg/mL, respectively).
	- 4. Saline solution: 0.9% NaCl in dH2O sterilized through 0.22-μ m filter.
	- 5. Antisense oligonucleotides: 50 μM stock solution in dH2O (final concentration depends on experimental setup).
	- 6. Fibroblast medium: DMEM 4.5 g/L glucose, L-glutamate, 10% fetal bovine serum (FBS), 1penicillin/streptomycin.
	- 7. Phosphate-buffered saline (PBS).
	- 8. Fibroblast transfection agent: Polyethylenimine (PEI) 1 mg/ mL.
	- 9. Keratinocyte medium: CellnTec Prime (CnT-PR).
	- 10. HEPES-buffered saline solution (HBSS).

#### 3 Methods

3.1 Isolation and Culture of Epidermal Keratinocytes and Dermal Fibroblasts

Full-thickness skin biopsies (4–6 mm) or larger skin tissue (1–2 cm) are used to isolate cells.



Fig. 1 Microscopy image of cells transfected with fluorescently labeled AON. Left: Fibroblasts transfected with a fluorescently labeled (green) AON using polyethylenimine. Right: Keratinocytes transfected with the same fluorescently labeled AON. A transfection efficiency of more than 95% is observed in both fibroblasts and keratinocytes, as shown by the green signal in the nuclei


#### 4 Notes


#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# In Vitro Delivery of PMOs in Myoblasts by Electroporation

### Remko Goossens and Annemieke Aartsma-Rus

#### Abstract

Antisense oligonucleotides (AONs) are small synthetic molecules of therapeutic interest for a variety of human disease. Their ability to bind mRNA and affect its splicing gives AONs potential use for exon skipping therapies aimed at restoring the dystrophin transcript reading frame for Duchenne muscular dystrophy (DMD) patients. The neutrally charged phosphorodiamidate morpholino oligomers (PMOs) are a stable and relatively nontoxic AON modification. To assess exon skipping efficiency in vitro, it is important to deliver them to target cells. Here, we describe a method for the delivery of PMOs to myoblasts by electroporation. The described protocol for the Amaxa 4D X unit nucleofector system allows efficient processing of 16 samples in one nucleocuvette strip, aiding in high-throughput PMO efficacy screens.

Key words AON, PMO, Electroporation, Nucleofection, Myocytes, Duchenne muscular dystrophy, DMD

#### 1 Introduction

Antisense oligonucleotides (AONs) are versatile, powerful tools for the potential treatment of a variety of diseases. AONs are short synthetic oligonucleotides consisting of modified DNA or RNA nucleic acid analogs. AONs can be exploited in multiple ways, including the modulation of splicing. Here, AONs bind to the unspliced mRNA and mask splice sites or exonic splice enhancer or silencer sites, resulting in an exon being ex- or included in the mature mRNA. Examples of such AONs are eteplirsen, golodirsen, viltolarsen, and casimersen to treat Duchenne muscular dystrophy (DMD) patients with eligible mutations, and nusinersen to treat spinal muscular atrophy (SMA). For DMD, antisense-mediated exon skipping is used to restore the reading frame of the dystrophin (DMD) transcript, allowing the production of an internally deleted partially functional dystrophin protein [1]. This approach is mutation specific. Currently, for Duchenne, four AONs have been approved by the Food and Drug Administration (FDA, USA),

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_12, © The Editor(s) (if applicable) and The Author(s) 2022

which induce skipping of exon 51 (eteplirsen), exon 53 (golodirsen and viltolarsen) or exon 45 (casimersen).

Various different AON chemistries have been developed, such as 20 -O-methyl phosphorothioate (2<sup>0</sup> O-MePS), 20 -O-methoxyethyl phosphorothioate (20 -MOE-PS), and phosphorodiamidate morpholino oligomer (PMO) [2]. These different chemistries have unique chemical properties and were developed to enhance stability, solubility, and cellular uptake of AONs [2]. AONs are sometimes covalently conjugated to other molecules, in an attempt to improve their uptake by target tissues after systemic delivery [3]. For treatment of SMA, intrathecal injection of the AON into the cerebrospinal fluid (CSF) leads to efficient uptake by neurons and other cells in the nervous system with a long half-life [4]. However, in DMD all of the >700 different skeletal muscles are affected, and as such systemic delivery of AONs is required, which currently involves weekly intravenous infusions [5].

DMD consists of 79 exons, and there is a wide variety of unique patient mutations [6, 7], with a mutation hotspot spanning exon 45 through 53 [6, 8]. Approximately 55% of total DMD mutations causative for Duchenne would be eligible for some form of exon skipping therapy [7]. While skipping certain exons is applicable to larger groups of patients, it is crucial to skip also additional exons, which individually apply to small groups of patients, to increase the general applicability of this approach to as many patients as possible. To optimally design AONs for most of the DMD exons, it is important to have the ability for high-throughput screening of AON exon skipping efficacy. To perform reliable initial testing of AONs in vitro, it is essential to establish a reproducible, efficient means of delivery to a target cell. This can be achieved using electroporation of immortalized muscle cells [9]. Using immortalized cells has the advantage of theoretically unlimited proliferation, so large amounts of cells with homogenous characteristics can be generated. Primary cell sources are finite, and each new donor will have to be validated for reproducibility, which hampers screening potential when a large number of different AONs are to be tested simultaneously. Furthermore, it is our experience that the capacity of primary cultures to differentiate into myotubes declines with advanced passages.

Unlike 20 O-MePS AONs and dsDNA (e.g., plasmids), PMO AONs are neutrally charged, impeding the delivery by cationic lipid transfection systems. An alternative method for delivery of PMOs to mammalian cells is electroporation [9]. Electroporation relies on the formation of pores in the cell membrane by the application of an electric pulse through the transfection medium, mediating delivery of the particle of interest. The pore-forming pulse either serves to simultaneously deliver a charged molecule or is followed by a dedicated secondary delivery pulse. While electroporation efficiency is aided by active mobilization of charged molecules of interest into the cells by the electric current applied, the pore formation itself is already able to allow passive entry of inert molecules such as PMOs, albeit potentially at a lower efficiency. In fact, efficiency of in vitro PMO electroporation is relatively high when compared to other methods such as gymnosis and calciumenriched medium (CEM) [10].

Classic electroporation is performed with cells in suspension, requiring dissociation of adherent cells from the culture vessel prior to the procedure. For studying DMD exon skipping, a cell line which expresses full-length DMD (Dp427m) is required. Alternatively, if available, a patient-derived cell line with a specific mutation can be used where skipping of a specific exon restores dystrophin production. However, as DMD expression in proliferating myoblasts is very low, myoblasts need to differentiate after electroporation with the Amaxa 4D X unit to form mature myotubes, expressing higher levels of DMD. Novel electroporation techniques also allow for electroporation of adherent cell layers, which can be advantageous when working with cells that grow slowly or have limited proliferation capacity upon differentiation, such as mature myotubes or neuronal cells. These methods, just like other electroporation techniques, require thorough characterization and optimization of conditions for maximum efficiency and are not included in this chapter.

In this chapter, we provide a protocol for the delivery of PMOs to immortalised myoblasts by electroporation with the Lonza Amaxa 4D-nucleofector X unit, and their subsequent differentiation to DMD expressing myotubes. The procedure for further sample processing and analysis of the skipped transcript by endpoint reverse transcription polymerase chain reaction (RT-PCR) and quantitative PCR (RT-qPCR) are also outlined.

#### 2 Materials

	- 2. Culture medium (proliferation): For myoblasts either: F10 Nutrient mix (nutmix) medium supplemented with 20% fetal bovine serum (FBS), 1% PenStrep, 10 ng/mL rhFGF and 1 mM dexamethasone, or Skeletal Muscle Cell Growth Medium (SMCGM) supplemented with 15% FBS and 50 μg/ mL gentamicin (see Note 2).
	- 3. Culture medium (resuspension): F10 Nutmix + 20% FBS + 1% PenStrep.
	- 4. Culture medium (differentiation): DMEM (4.5 g/L glucose) + 2% FBS or 2% knockout serum replacement (KOSR) + 50 μg/mL Gentamicin or 1% PenStrep (see Note 2).

#### 2.2 Electroporation 1. Lonza 4D nucleofector core unit.

	- 2. Chloroform.
	- 3. 2-Propanol.
	- 4. 70% Ethanol (EtOH).
	- 5. RNase-free water (DEPC treated).
	- 2. dNTP mix (10 mM each).
	- 3. 5reverse transcriptase (RT) reaction buffer.
	- 4. Reverse transcriptase enzyme.
	- 5. RNase inhibitor.
	- 6. RNase-free water (DEPC treated).

#### 2.5 RT-PCR Analysis of Skipping Efficiency 1. cDNA generated from >1 μg total RNA by random hexamer primers (Subheading 3.3, step 5).


#### 2.6 RT-qPCR of Skipping Efficiency 1. cDNA generated from >1 μg total RNA by random hexamer primers (Subheading 3.3, step 5).


#### 3 Methods

RNA isolated from the nucleofected myotube cultures is used to generate cDNA, which can subsequently be used for RT-PCR or RT-qPCR analysis of exon skipping efficiency. To purify a sufficient amount of total RNA from myotube cultures, we usually transfer cells to six-well plates after nucleofection. Smaller culture vessel might be suitable, or more optimal, for other purposes. We use up to 1 - 10<sup>6</sup> myoblast cells per 20 μL nucleofection reaction, requiring 1.6 - 10<sup>7</sup> cells in total for a full 16-well cuvette strip. A T182 culture vessel of high confluence contains about 2 - 107 myoblast cells. A graphical overview of the procedure is outlined in Fig. 1. All steps described in Subheading 3.1, steps 1–17 should be performed under aseptic conditions. Subheading 3.1, steps 18 through Subheading 3.2, step 11 should be performed using suitable personal protection according to local regulations.

	- 1. Preferentially subculture myoblasts 1 day prior to nucleofection to ensure proper viability and growth phase of cells. Cells from confluent cultures might exhibit a decreased efficiency or survival.
	- 2. Label wells and add media to the culture vessels used to transfer cells post-nucleofection, e.g., 3 mL proliferation media per well of a 6-well plate. Place plates in a 37 C CO2 incubator to warm and equilibrate the media.
	- 3. On the day of the nucleofection experiment, wash the cells with dPBS and trypsinize the cells. Use 2 mL trypsin 0.05% for a

Fig. 1 Schematic workflow of nucleofecting PMO in myoblast cells

T182 and carefully tilt flask to cover all cells. Place cells with trypsin in a 37 C incubator.

	- 2. Mix thoroughly by shaking for 30 s.
	- 3. Incubate on ice for 5 min.
	- 4. Centrifuge samples for 15 min at 16,000 RCF at 4 C.
	- 5. Carefully transfer the upper aqueous phase to a new microcentrifuge tube, without disturbing the organic- and interphase.
	- 6. Add an equal volume of 2-propanol to transferred aqueous phase.
	- 7. Mix well and incubate for >30 min on ice, then centrifuge for 15 min at 16,000 RCF at 4 C.
	- 8. Discard supernatant and wash pellet with 1 mL 70% EtOH.
	- 9. Centrifuge for 5 min at 16,000 RCF at 4 C.
	- 10. Discard supernatant and air-dry pellet.
	- 11. Dissolve the pellet in RNase-free water (e.g., 25 μL).
	- 12. Determine RNA concentration using a Nanodrop ND-1000.
	- 13. Store RNA at <sup>80</sup> C (see Note 19).
	- (a) X μL RNA for a total of 1 μg.
	- (b) 1 μL dNTP mix (10 mM each).
	- (c) 1 μL Random hexamer primers (N6) (20 ng/μL).
	- (d) X μL RNase-free water up to 12.5 μL.
	- 2. Incubate reaction for 5 min at 65 C. Chill on ice for 2 min.
	- 3. To each tube add:
		- (a) 4 μL 5-RT-reaction buffer.
		- (b) 1 μL Reverse transcriptase enzyme.
		- (c) 0.5 μL RNase inhibitor.
	- 4. Incubate at 42 C for 1 h.
	- 5. Incubate at 85 C for 5 min.
	- 6. Dilute the 20 μL cDNA reaction to a final volume of 100 μL with ultrapure MQ.
	- 7. Store cDNA at 20 C.

#### 3.4 RT-PCR Analysis 1. Set up RT-PCR (25 μL reactions) according to the follow set-up per reaction (see Note 21):


Fig. 2 Example of the result of RT-PCR analysis after successful nucleofection of a DMD exon 51 targeting PMO in KM155 myotubes using various programs of the 4D-nucleofector. (a) Agarose gel electrophoresis of endpoint RT-PCR products using primers specific for DMD exon 47 through 52. Skipping of exon 51 will lead to the production of a smaller PCR product as indicated. Different lanes consist of different experimental combination of nucleofection buffers and programs. (b) Approximate quantification of agarose gel shown in A by FIJI analysis. Ratios of Exon 51 skipped product intensities over the regular (exon 51 containing) product are plotted. The average of the negative controls (no PMO/no pulse samples (orange bars)) is shown as a dotted line. (c) RT-qPCR analysis of DMD exon 51 skipping in KM155 myotubes with 6 nucleofection programs and 5 nucleofection buffers. Exon skipping was determined with a primer set only amplifying the DMD transcript without exon 51 (Exon 50-52F + Exon 52R). A primer set specific for DMD exon 49 through 50 shows all DMD transcript, and MYH3 is used as a marker for myogenic differentiation. Cells resuspended in nucleofection buffer not subjected to an electric pulse were used as a negative control. In our hands, exon skipping efficiency was highest using a combination of program CM-137 and buffer P1

	- (a) 1: 5 min, 95 C—initial melt.
	- (b) 2: 30 s, 95 C—melt.
	- (c) 3: 30 s, 60 C—annealing (change for specific primers).
	- (d) 4: 40 s, 72 C—extension (~30 s per kb).
	- (e) 5: Go to step 2, 34 additional times (35 cycles total).
	- (f) 6: 5 min, 72 C—final extension.

1. For RT-qPCR analysis, measure the gene of interest and at least one reference gene. Always measure a technical triplicate of each cDNA-primer combination. Set up RT-qPCR according to the following set-up per reaction (we use 8 μL reactions in a 384-well plate):

	- (a) 1: 5 min, 95 C—initial melt.
	- (b) 2: 10 s, 95 C—melt.
	- (c) 3: 30 s, 60 C—Annealing and extension (change temperature for specific primers).
	- (d) 4: Read plate, go to step 2, 39 additional times (40 cycles total).
	- (e) 5: Melt curve analysis.

3.5 RT-qPCR Analysis

4. A suitable primer pair for the skipped product will yield little to no signal in the un-nucleofected control samples and will have increased abundance when the exon was successfully skipped.

#### 4 Notes


incorrectly. Therefore, while useful for estimating relative intensity, many applications will require more sensitive methods to reliably quantify exon skipping efficiency. Furthermore, detection of DNA in agarose gels is facilitated by the amount of EtBr intercalating with the DNA, resulting in signal when exposed to UV light. As shorter PCR products bind less EtBr, their intensity is inherently underestimated when measured in analysis software.

23. Prior to analyzing PCR products on an Agilent 2100 bioanalyzer, it is advised to always run some of the sample on an EtBr agarose gel to confirm successful PCR amplification.

#### References


Guergueltcheva V, Chan S, Korngut L, Campbell C, Dai Y, Wang J, Barisˇic´ N, Brabec P, Lahdetie J, Walter MC, Schreiber-Katz O, Karcagi V, Garami M, Viswanathan V, Bayat F, Buccella F, Kimura E, Koeks Z, van den Bergen JC, Rodrigues M, Roxburgh R, Lusakowska A, Kostera-Pruszczyk A, Zimowski J, Santos R, Neagu E, Artemieva S, Rasic VM, Vojinovic D, Posada M, Bloetzer C, Jeannet PY, Joncourt F, Dı´az-Manera J, Gallardo E, Karaduman AA, Topalog˘lu H, El Sherif R, Stringer A, Shatillo AV, Martin AS, Peay HL, Bellgard MI, Kirschner J, Flanigan KM, Straub V, Bushby K, Verschuuren J, Aartsma-Rus A, Be´roud C, Lochmu¨ller H (2015) The TREAT-NMD DMD global database: analysis of more than 7,000 Duchenne muscular dystrophy mutations. Hum Mutat 36(4):395–402. https://doi.org/10.1002/ humu.22758


effect of oligonucleotides. Nucleic Acids Res 43(19):e128. https://doi.org/10.1093/nar/ gkv626


Lee J, Touznik A, Mamchaoui K, Aoki Y, Takeda S, Nagaraju K, Mouly V, Maruyama R, Duddy W, Yokota T (2017) Quantitative antisense screening and optimization for exon 51 skipping in Duchenne muscular dystrophy. Mol Ther 25(11):2561–2572. https://doi.org/10.1016/j.ymthe.2017. 07.014

13. Spitali P, Heemskerk H, Vossen RHAM, Ferlini A, den Dunnen JT, t Hoen PAC, Aartsma-Rus A (2010) Accurate quantification of dystrophin mRNA and exon skipping levels in duchenne muscular dystrophy. Lab Investig 90(9):1396–1402. https://doi.org/10.1038/ labinvest.2010.98

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Rapid Determination of MBNL1 Protein Levels by Quantitative Dot Blot for the Evaluation of Antisense Oligonucleotides in Myotonic Dystrophy Myoblasts

### Nerea Moreno, Irene Gonza´ lez-Martı´nez, Rube´n Artero, and Estefanı´a Cerro-Herreros

#### Abstract

Western blot assays are not adequate for high-throughput screening of protein expression because it is an expensive and time-consuming technique. Here we demonstrate that quantitative dot blots in plate format are a better option to determine the absolute contents of a given protein in less than 48 h. The method was optimized for the detection of the Muscleblind-like 1 protein in patient-derived myoblasts treated with a collection of more than 100 experimental oligonucleotides.

Key words Myotonic dystrophy, Oligonucleotides, Quantitative dot blot, Muscleblind-like protein 1, DM1 myoblasts

#### 1 Introduction

Myotonic dystrophy type 1 (DM1) is a degenerative genetic disease that is classified as rare because it affects less than 1 in 2000 people (1/3000 to 1/8000; [1]). DM1 originates from an expansion of the CTG trinucleotide repeat in the 3<sup>0</sup> -untranslated region (UTR) of the DMPK gene that, upon transcription, forms CUG hairpins that behave as toxic RNAs. CUG expansion RNA aberrantly binds and sequesters essential developmental proteins of the Muscleblind-like (MBNL) family, which are key regulators of alternative splicing. The depletion in MBNL protein function causes alterations in RNA metabolism that originate defined symptoms of the disease [2]. Studies in animal models have shown that the increase of MBNL in a genetic background of DM improves the

Nerea Moreno and Irene Gonza´lez-Martı´nez contributed equally to this work.

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_13, © The Editor(s) (if applicable) and The Author(s) 2022

pathological phenotypes and that the overexpression of MBNL1 in control mice is well tolerated [3].

Patients suffer from myotonia and muscle atrophy and weakness, which, in advanced stages of the disease, lead to respiratory distress and early death. Currently, there is no effective treatment for DM1, and management of symptoms is the only option to preserve the quality of life of people living with DM1. In its most common form, the onset of symptoms occurs during adolescence, and affected people have a significantly shortened lifespan of 48–55 years. Because of its prevalence and the severity of the clinical manifestations, finding a cure for DM1 is a social and medical need [4, 5].

The development of effective high-throughput tools in drug discovery research has increased the demand for complementary high capacity immunoblot methods in which to assess the consequences of drug candidates at protein level. One example is the need to quickly evaluate the levels of MBNL1 protein in patientderived myoblasts [6] treated with hundreds of oligonucleotide variants to block repressive miRNAs miR-23b and miR-218, as a means to boost endogenous levels and compensate sequestration by CUG expansions in mutant DMPK [7]. To this end, we have generated a diversity (>100) of highly modified antisense oligonucleotides (AONs) to block miR-23b and miR-218. Examples of these modifications are the substitution of natural ribose rings by locked nucleic acid (LNAs), C2<sup>0</sup> hydroxyl substitutions with a methoxy (2<sup>0</sup> OMe) or methoxyethyl (2<sup>0</sup> -MOE), or the use of phosphorothioates to link two nucleotides, instead of natural phosphodiester bonds, to improve stability in vivo as they make these ASOs resistant to intracellular and extracellular nucleases. AON can also be made electrostatically neutral by using phosphorodiamidate morpholino oligomers (PMO) and can be conjugated to a cholesterol moiety to improve the diffusion through cell membranes and cell uptake [8, 9].

For this purpose, we propose the use of quantitative dot blot (QDB) analysis as an alternative to Western blot. For the development of the QDB assay, we have modified two previously published protocols [10, 11]. Dot blot was developed to simplify the process of Western blot analysis (Fig. 1) when the antibody is very specific for the detection of the protein of interest, and there is no need to determine its molecular weight, for example, when screening the effects of several molecules on the expression of a single protein. Specifically, QDB transforms traditional immunoblots into quantitative assays and allows expression analysis of a certain protein in your samples in 96-well format, being more efficient and faster than a Western blot.

Fig. 1 Illustration of the entire QDB process for evaluation of antisense oligonucleotides in DM1 myoblast

#### 2 Materials


cocktail. Storage in 4 C.



Fig. 2 A photograph of a QDB plate upside down. The figure shows the different components of the plate and how to load the sample



#### Acknowledgments

The project leading to these results has received funding from "la Caixa" Banking Foundation under the project code HR17-00268 (TATAMI project to R.A.). I.G.-M. was funded by the Precipita Project titled "Desarrollo de una terapia innovadora contra la distrofia mioto´nica"; E.C.-H. was supported by the post-doctoral fellowship APOSTD/2019/142; and N.M.-C was supported by the pre-doctoral fellowship PRE2019-090622. Part of the equipment employed in this work has been funded by Generalitat Valenciana and cofinanced with ERDF funds (OP ERDF of Comunitat Valenciana 2014-2020).

#### References


RNA, sequestration of muscleblind proteins and deregulated alternative splicing in neurons. Hum Mol Genet 13(24):3079–3088. https:// doi.org/10.1093/hmg/ddh327


myotonic dystrophy muscle cell lines to assess therapeutic compounds. Dis Model Mech 10(4):487–497. https://doi.org/10.1242/ dmm.027367


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Evaluation of Exon Skipping and Dystrophin Restoration in In Vitro Models of Duchenne Muscular Dystrophy

### Andrea Lo´ pez-Martı´nez, Patricia Soblechero-Martı´n, and Virginia Arechavala-Gomeza

#### Abstract

Several exon skipping antisense oligonucleotides (eteplirsen, golodirsen, viltolarsen, and casimersen) have been approved for the treatment of Duchenne muscular dystrophy, but many more are in development targeting an array of different DMD exons. Preclinical screening of the new oligonucleotide sequences is routinely performed using patient-derived cell cultures, and evaluation of their efficacy may be performed at RNA and/or protein level. While several methods to assess exon skipping and dystrophin expression in cell culture have been developed, the choice of methodology often depends on the availability of specific research equipment.

In this chapter, we describe and indicate the relevant bibliography of all the methods that may be used in this evaluation and describe in detail the protocols routinely followed at our institution, one to evaluate the efficacy of skipping at RNA level (nested PCR) and the other the restoration of protein expression (myoblot), which provide good results using equipment largely available to most research laboratories.

Key words Dystrophin, Antisense oligonucleotide, Duchenne muscular dystrophy, Eteplirsen, Myoblot, Exon skipping

#### 1 Introduction

Duchenne muscular dystrophy (DMD) is an X-linked inherited neuromuscular disorder caused by mutations in the dystrophin gene (DMD) and characterized by rapid progression of muscle weakness and loss of ambulation in early adolescence. Most DMD mutations are deletions that disrupt the open reading frame (ORF), preventing the synthesis of dystrophin protein. On the other hand, Becker's muscular dystrophy (BMD) is a milder neuromuscular disorder caused by in-frame mutations in the same gene. In this case, mutations do not disrupt the ORF and a shorter but semifunctional form of dystrophin is produced resulting in a milder form of disease [1, 2].

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_14, © The Editor(s) (if applicable) and The Author(s) 2022

The reading-frame rule, exemplified in Becker patients, is the basis of the concept of exon skipping by antisense oligonucleotides (AONs) as a possible therapy for DMD. In this case, AONs are designed to attach to specific RNA sequences and disrupt the binding of the spliceosome, which in turn causes the skipping of a particular exon and the restoration of the expression of a truncated, but semi-functional, dystrophin [3]. This therapeutic approach could benefit around 83% of DMD patients, but due to the vast array of different DMD deletions and mutations causing DMD, to treat each subset of DMD patients, different exons would need to be targeted by specific AONs [4, 5]. Eteplirsen, golodirsen, viltolarsen, and casimersen are AONs that facilitate the skipping of DMD exons 51, 53, 53, and 45 respectively, and have been approved by the FDA (2016, 2019, 2020, and 2021) [6– 10]. These drugs would only be applicable to 13% (eteplirsen) and 8% of DMD patients (golodirsen, viltolarsen, and casimersen). New AONs, skipping other exons, are being designed for the remaining deletions that may benefit from this approach [11–14].

The preclinical development of new exon-skipping AONs relies on the use of cell cultures from patients harboring specific deletions. It is common practice to design a panel of candidate AONs and evaluate their efficacy in culture before advancing to preclinical testing, and many methods are followed to evaluate them, at both RNA and protein levels, as represented in Fig. 1.

1.1 Evaluating Exon Skipping at RNA Level When evaluating exon skipping at RNA level, several methods, both quantitative and semi-quantitative, are routinely used. A recent publication compared five different quantification techniques: quantitative real-time PCR (qPCR) [18], digital droplet PCR (ddPCR) [23], single PCR assessed with an Agilent bioanalyzer, and nested PCR with agarose gel image analysis [15– 17]. They concluded that ddPCR was the most precise and less dispersed quantitative method for exon-skipping detection, while the other techniques may overestimate exon skipping and present high data variation.

The ddPCR protocol is based on the mix of the DNA sample with an oil–water media to divide the template in several thousand droplets. Each droplet contains none or one single template strand that would provide individual amplification reactions and absolute quantification. It is a highly reproducible and efficient technique as it does not depend on the PCR efficiency and may be adapted to quantify exon skipping using specific probes. Nevertheless, the technique requires highly specific equipment not available in most laboratories [23].

Quantitative real-time PCR for DMD transcripts uses custom probes to specifically detect skipped transcripts that are lately amplified for transcript quantification. Data are normalized with endogenous transcript controls but the requirement of a pre-amplification

Fig. 1 Methods that may be used to evaluate exon skipping in cell culture. Currently, the most accurate methods are digital droplet PCR (ddPCR) and capillary western immunoassay (Wes), but the equipment required to perform these experiments is not present in the majority of research laboratories. Advantages (green) and disadvantages (red) of each method are presented in this figure. Main bibliographical references for these methods are: RT-PCR followed by gel densitometry analysis [15, 16], single-round PCR followed by bioanalyzer [16, 17], qPCR [18], ddPCR (13), capillary western immunoassay (Wes) [19], Western blotting [20], myoblots [21]. Several attempts to compare these methods have been published by a consortium to evaluate dystrophin protein quantification methods [20] and another to compare methods to quantify exon skipping [17]. A recent meeting to find consensus among stakeholders on all dystrophin protein quantification methods described was also recently published [22]. Created with Biorender.com

> step and specific probes for each of the deletions tested may decrease linearity and increase costs [18].

> Most methods are semi-quantitative at best and provide an indication of efficacy that should be evaluated further by assessing the downstream consequences of exon skipping (dystrophin restoration). Our approach combines the study of exon skipping at RNA level by a rather simple nested PCR plus gel densitometry analysis method, with a quantitative evaluation of dystrophin protein by myoblots.

1.2 Evaluating Dystrophin Restoration in Cell Culture Dystrophin protein quantification can be challenging due to its large size (3685 amino acids) and expression pattern, and it has been an extremely hot topic when evaluating the results of dystrophin restoration clinical trials [24]. A recent meeting of stakeholders interested in dystrophin quantification methodology highlighted the problems derived from the variability of dystrophin

Fig. 2 Workflow of an exon skipping evaluation experiment. RNA evaluation: a 6-well plate is seeded and transfected with two different AON concentrations. Cells are collected after 48 h for RNA extraction, reverse transcription (RT), and nested PCR analysis by densitometry. Dystrophin protein quantification: a 96-well plate is seeded, transfected, and allowed to differentiate for a week. Plates are fixed before being analyzed by myoblot. Created with Biorender.com

levels between different muscles and individuals as well as the lack of standardized controls [22]. When analyzing preclinical in vitro experiments, other points need to be considered: patient-derived cell cultures are difficult to expand and differentiate, and dystrophin expression is only detectable when cells are differentiated into myotubes. Western blotting analysis from cultured cells requires very large amounts of protein lysate that does not produce good quantitative results while using a lot of sample.

A novel advance in protein quantification is the capillary Western immunoassay (Wes), based on the use of capillary as separation modules for protein isolation followed by dystrophin detection by an immunoassay that allows accurate quantification. The Wes system is said to detect low protein concentrations, in a range of 0.125–1.25 μg [19, 22]. However, this equipment is not yet commonly found in most laboratories.

We detail in this chapter the quantitative method developed at our department: myoblots, a method based on the in-cell western blotting optimized for the quantification of muscle proteins in cell culture [21]. Myoblots are performed in 96-well plates seeded with patient-derived cells, which are allowed to differentiate. Signal is normalized by cell number, and the differentiation status of the cultures is assessed as a quality control of the experiment.

A summary of our strategy to evaluate exon skipping drugs in cell culture is shown in Fig. 2. In the example used in this chapter, we evaluated two concentrations of an AON skipping exon 51 on immortalized cell cultures [25] from a patient harboring a mutation that causes the deletion of exon 52 from the CNMD Biobank (London, UK).

#### 2 Materials


9. 5 PCR Loading Buffer.



Specific primer 55R was used in the RT reaction. Two different sets of primers targeting regions between exons 48F-54R (first PCR) and 49F-53R (second PCR) were used for the nested-PCR method

10. HyperLadder 100 bp.

11. Gel extraction kit to isolate PCR fragments for sequencing.

#### 2.3 Myoblots 1. Access to an Odyssey CLx Scan (LI-COR Biosciences).

	- (a) MF20 antibody (Developmental Studies Hybridoma Bank (DSHB)).
	- (b) Mandys1, kindly provided by Prof. G Morris, The MDA Monoclonal Antibody Resource.
	- (c) Mandys106, kindly provided by Prof. G Morris, The MDA Monoclonal Antibody Resource.
	- (d) Dys1 (Leica Biosystems).
	- (a) Goat Anti-Mouse IgG H&L (Biotin, Abcam).
	- (b) IRDye 800CW Streptavidin (LI-COR Biosciences).
	- (c) IRDye 800CW Goat anti-Mouse IgG (LI-COR Biosciences).
	- (d) CellTag™ 700 Stain from (LI-COR Biosciences).


incubator.

Fig. 3 Examples of plate distribution for exon skipping evaluation at protein level. (a) The four experimental conditions are distributed as follows in the 96-well plate: 2 columns for control (Lipofectamine only), 2 columns for DM (differentiation medium only), 4 columns for 100 nM AON, and 4 columns for 300 nM AON, both transfected with Lipofectamine. (b) The four experimental conditions are distributed as follows in the 96-well plate: 1 row for control (Lipofectamine only), 1 row for DM (differentiation medium only), 3 rows for 100 nM AON, and 3 rows for 300 nM AON, both transfected with Lipofectamine. (c) Reduction of the edge effect by filling edge wells with 100 <sup>μ</sup>l of preferred buffer, but not cells. Then, the four experimental conditions are distributed as follows in the 96-well plate: 1 row for control (Lipofectamine only), 1 row for DM (differentiation medium only), 2 rows for 100 nM AON, and 2 rows for 300 nM AON, both transfected with Lipofectamine. Each experimental condition will be probed with different antibodies in the 800-nm channel (green): the majority of wells will be probed with a dystrophin antibody mix, some with an MF20 antibody (experimental quality control to assess differentiation), and the remaining wells will have no primary antibody to eliminate background. In the 700-nm channel (red), every well will be probed with a cell number stain (CellTag™ 700 Stain) to allow for cell number normalization. Created with Biorender.com


may be stored in PBS at 4 C until analysis (see Subheading 3.3).

#### 3.2 Nested PCR and Gel Image Analysis Method 1. After RNA extraction, RNA concentrations are measured with Nanodrop and 1 μg of template RNA is used for reverse transcription (RT).

3.2.1 Reverse Transcription

	- 2. Add 3 μl of cDNA (direct RT product) to the PCR mastermix.
	- 3. Run the first PCR as follows: 94 C for 5 min, 30 cycles of 94 C for 40 s, 60 C for 40 s, and 72 C for 1 min and 20 s, and finally, 72 C for 7 min.
	- 4. Three microliters of the first PCR product are used as template in the second PCR, which has the same PCR mastermix plus 2 μl of each of the second pair of primers at 10 μM (49F and 53R in the example) (see Note 6).
	- 5. The second PCR (or nested) should be run as follows: 94 C for 5 min, 35 cycles of 94 C for 40 s, 60 C for 40 s, and 72 C for 1 min and 20 s, and finally, 72 C for 7 min.
	- 6. Run a 2% agarose gel with the whole PCR product (50 μl) for 1 h at 100 mV.
	- 7. Place gel on the transilluminator of the gel documentation system and acquire the image. A representative image is shown in Fig. 4b. This figure shows the skipped and non-skipped product highlighting the bands selected for quantification.
	- 8. Cut the relevant PCR bands for DNA extraction and verification by Sanger sequencing.



Fig. 4 Nested PCR results. (a) Skipped and non-skipped product schematic representations. Primers 48F and 54R were used in the first PCR and primers 49F and 53R in the second. Skipped product size is reduced due to the lack of exon 51 and 52 while the non-skipped product is just lacking exon 52. Created with Biorender.com. (b) Agarose gel image from AON transfection. Fifty microliter of PCR product were examined in a 2% agarose gel stained with SYBRSafe. Image was captured in a Gel Doc™ EZ Imager (BIORAD). Red boxes indicate the quantified bands. The described relative percentage of exon skipping for 100 nM AON was 95.41% and for 300 nM AON was 53.62%


Exon skipping% <sup>¼</sup> ð Þ normalized peak area skipped fragment ð Þþ normalized peak area skipped fragment ð Þ normalized area non skipped fragment <sup>100</sup>



#### 3.3.2 Myoblot Analysis 1. In the Image Studio™ Software from your computer, select "In Cell Western analysis" and start the acquisition.


Fig. 5 In cell Western analysis. (a) Readings performed by the Odyssey Scan at 700 nm (Red, CellTag™ 700 Stain) and 800 nm (Green, Dystrophin, MF20 and background) provide a single intensity value for each well at each channel. To normalize each value, the corresponding background average value (no primary antibody) is subtracted from each value and later divided by its corresponding one for the CellTag™ 700 Stain signal (700 nm value). Created with Biorender.com. (b) Example of omitted wells by antibody specks (marked with arrows) and its corresponding eliminated value. (c) Bar graph of data: average SEM of the two AON concentrations against control (lipofectamine only) and analysis by two-way ANOVA with multiple comparisons (\*\*\*\*<sup>p</sup> <sup>&</sup>lt; 0.0001)


protein level. As seen in the example we used, they show concordant results supports AON suitability for dystrophin restoration in preclinical studies.

#### 4 Notes


#### Acknowledgments

This work was supported by funding from Health Institute Carlos III (ISCIII, Spain) and the European Regional Development Fund (ERDF/FEDER), "A way of making Europe" (grants CP12/ 03057 and PI15/00333), Basque Government (grants 2016111029), and Duchenne Parent Project Spain (grant 05/2016). V.A.-G. holds a Miguel Servet Fellowship from the ISCIII (CPII17/00004), part-funded by ERDF/FEDER. P. S-M holds a Rio Hortega Fellowship from ISCIII (CM19/00104), part funded by ERDF/FEDER. P. S-M holds a Rio Hortega Fellowship from ISCIII (CM19/00104) V.A.-G. also acknowledges funding from Ikerbasque (Basque Foundation for Science). The authors declare that they have no conflict of interest.

#### References


Dystrophin quantification and clinical correlations in Becker muscular dystrophy: implications for clinical trials. Brain 134(pt 12): 3547–3559. https://doi.org/10.1093/ brain/awr291


of antisense-mediated exon skipping for Duchenne muscular dystrophy mutations. Hum Mutat 30(3):293–299. https://doi.org/10. 1002/humu.20918


20(2):102–110. https://doi.org/10.1016/j. nmd.2009.10.013


13(4):e0195850. https://doi.org/10.1371/ journal.pone.0195850


organisation meeting on dystrophin quantification methodology. J Neuromuscular Dis 6 (1): 147–159. doi:https://doi.org/10.3233/ JND-180357


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Generation of Human iPSC-Derived Myotubes to Investigate RNA-Based Therapies In Vitro

### Pablo Herrero-Hernandez, Atze J. Bergsma, and W. W. M. Pim Pijnappel

#### Abstract

Alternative pre-mRNA splicing can be cell-type specific and results in the generation of different protein isoforms from a single gene. Deregulation of canonical pre-mRNA splicing by disease-associated variants can result in genetic disorders. Antisense oligonucleotides (AONs) offer an attractive solution to modulate endogenous gene expression through alteration of pre-mRNA splicing events. Relevant in vitro models are crucial for appropriate evaluation of splicing modifying drugs. In this chapter, we describe how to investigate the splicing modulating activity of AONs in an in vitro skeletal muscle model, applied to Pompe disease. We also provide a detailed description of methods to visualize and analyze gene expression in differentiated skeletal muscle cells for the analysis of muscle differentiation and splicing outcome. The methodology described here is relevant to develop treatment options using AONs for other genetic muscle diseases as well, including Duchenne muscular dystrophy, myotonic dystrophy, and facioscapulohumeral muscular dystrophy.

Key words Splicing, Human iPSC, Skeletal muscle, Antisense oligonucleotides, In vitro models

#### 1 Introduction

Pre-mRNA splicing is a highly conserved process in eukaryotes that plays a role in pre-mRNA processing. Alternative splicing can diversify gene function to produce isoforms with specific functions in distinct cell types [1, 2]. Genetic variations can lead to defects in pre-mRNA splicing that cause human disease [3]. Modulation of pre-mRNA splicing can be directed to correct aberrant splicing, to skip protein coding variants, to restore the reading frame, or to prevent expression of toxic gene products. This is possible by targeting antisense oligonucleotides (AONs) toward canonical splice sites or to cis-acting regulatory elements such as cryptic splice sites or splicing silencers/enhancers [4]. Alternative splicing in skeletal muscle is abundant and essential for muscle development and function [5]. Deregulation of pre-mRNA splicing in skeletal muscle is known to be the underlying cause of multiple human

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_15, © The Editor(s) (if applicable) and The Author(s) 2022

myopathies [5]. Suitable in vitro and in vivo models are crucial to investigate novel splicing modulating drugs in target cells and tissues.

In vitro human skeletal muscle models can be obtained directly from muscle biopsies, or these can be generated by (trans-)differentiation of primary fibroblasts, pluripotent stem cells, or non-muscle cells with myogenic capacity like pericytes and mesoangioblasts [6–8]. Several protocols have been described to generate muscle progenitor cells (MPCs) derived from human patientderived induced pluripotent stem cells (hiPSCs) using directed differentiation methods for disease modeling [9–11].

Here we describe how purified, expandable hiPSC-derived MPCs, generated using a transgene-free procedure [11] can be differentiated into multinucleated myotubes to test the modulating activity of AONs. These methods can be used to analyze splicing correction in vitro to develop RNA-based therapies for muscle disorders. We have used this strategy to test AONs for Pompe disease [12] and describe the methodology here in detail.

#### 2 Materials

All cell culture work needs to be performed under sterile conditions in safety cabinets. All cell lines should be tested for mycoplasma following the manufacturer instructions (Lonza; LT07-318). Cell lines are cultured at 5% CO2 and 37 C in humidified incubators.

2.1 Skeletal Muscle Progenitor Cell Culture 1. Human MPC lines (see Note 1). 2. DMEM 4.5 g/L Glucose. 3. Fetal Bovine Serum. 4. Penicillin/Streptomycin/Glutamine 100 (p/s/g). 5. Fibroblast Growth Factor 2 (FGF2) (see Note 2). 6. Sterile cell culture grade Bovine Serum Albumin (7.5% BSA). 7. TrypLE™ Express Enzyme (1), phenol red. 8. Phosphate Buffered Saline (DPBS). 9. Extracellular Matrix gel from Engelbreth (ECM; 1). 10. DMEM:F12. 11. Insulin/Transferrin/Selenium 100. 12. DMSO. 13. Freezing containers. 2.2 Cell Culture Media 1. Proliferation medium: DMEM 4.5 g/L Glucose, supplemented with 10% FBS, 1 Pen/Strep, and 100 ng/ml FGF2 (added directly to plate/well).


2.5 RNA Isolation, cDNA Synthesis, and Quantitative RT-PCR (RT-qPCR)


#### Table 1 Primers used for RT-qPCR


#### 3 Methods


	- 1. Resuspend the PMO AONs in RNAse-free MilliQ at a concentration of 1 mM.
	- 2. Add 4.5 μl of Endoporter reagent per ml of medium directly to the cells and mix by gentle shaking (see Note 7).
	- 3. Add the desired amount of PMO AONs to the cells and mix by gentle shaking.
	- 4. Transfect AONs 1 day prior differentiation (day 1). Cells should be 60–80% confluent.
	- 5. Switch to differentiation medium (day 0).
	- 6. Leave cells to differentiate for 4 days and either collect protein or RNA or fix cells for immunofluorescence.
	- 1. For immunofluorescence analysis of patient-derived myotubes, prepare cells using 48-well plates.
	- 2. Wash cells once in PBS.
	- 3. Fix cells using 4% PFA in PBS for 10 min at RT, remove and add PBS. Cells can be stored at 4 C before proceeding.
	- 4. Wash twice in PBS for 2 min each.
	- 5. Incubate for 10 min with 0.3% Triton-X100 in PBS for permeabilization.
	- 6. Incubate for 30 min with 3% BSA, 0.1% Tween in PBS for blocking.
	- 7. Repeat washing step 4.
	- 8. Incubate with primary antibodies for 1 h at RT in 0.1% BSA, 0.1% Tween in PBS (see Note 8).
	- 9. Repeat washing step 4.
	- 10. Incubate with secondary antibodies for 45 min at RT in 0.1% BSA, 0.1% Tween in PBS.
	- 11. Repeat washing step 4.
	- 12. If biotinylated antibodies were used, incubate with tertiary for 30 min at RT in 0.1% BSA, 0.1% Tween in PBS.
	- 13. Repeat washing step 4.

3.2.2 Immunofluorescence

3.2 Delivery and Efficacy of Antisense Oligonucleotides in Patient-Derived Myotubes

3.2.1 Transfection

Fig. 1 Wide field images of differentiating MPCs. Representative images of the differentiation of MPCs over 4 days. Scale bar 100 <sup>μ</sup>m

Fig. 2 Immunofluorescence images of 4 days differentiated MPCs. MPCs were stained with MYH1E (red), MYOGENIN (green), and the nuclei with Hoechst (blue). Arrowheads indicate nuclei present in multinucleated myotubes. Scale bar 100 <sup>μ</sup>m


#### 4 Notes


Endoporter used is independent of the concentration of PMO AONs. Other backbones might require different delivery reagents.


#### Acknowledgments

We thank Erik van der Wal for the critical reading and the revisions of this manuscript. This work was funded by Tex net, the Prinses Beatrix Spierfonds/Stichting Spieren voor Spieren (grant W. OR13-21), the Sophia Children's Hospital Foundation (SSWO) (grant S-687 and S17-32), and Metakids (grant 2016-063).

#### References


phenotypes among Duchenne muscular dystrophy patient-specific myoblasts derived using a human iPSC-based model. Cell Rep 15(10):2301–2312. https://doi.org/10. 1016/j.celrep.2016.05.016


van Ijcken WFJ, Cheung TH, van der Ploeg AT, Schaaf GJ, Pijnappel W (2018) Largescale expansion of human iPSC-derived skeletal muscle cells for disease modeling and cellbased therapeutic strategies. Stem cell Rep 10(6):1975–1990. https://doi.org/10.1016/ j.stemcr.2018.04.002

12. van der Wal E, Bergsma AJ, van Gestel TJM, In 't Groen SLM, Zaehres H, Arauzo-Bravo MJ, Scholer HR, van der Ploeg AT, Pijnappel W (2017) GAA deficiency in Pompe disease is alleviated by exon inclusion in iPSC-derived skeletal muscle cells. Mol Ther Nucleic Acids 7:101–115. https://doi.org/10.1016/j. omtn.2017.03.002

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 16

# Eye on a Dish Models to Evaluate Splicing Modulation

### Kwan-Leong Hau, Amelia Lane, Rosellina Guarascio, and Michael E. Cheetham

#### Abstract

Inherited retinal dystrophies, such as Leber congenital amaurosis, Stargardt disease, and retinitis pigmentosa, are characterized by photoreceptor dysfunction and death and currently have few treatment options. Recent technological advances in induced pluripotent stem cell (iPSC) technology and differentiation methods mean that human photoreceptors can now be studied in vitro. For example, retinal organoids provide a platform to study the development of the human retina and mechanisms of diseases in the dish, as well as being a potential source for cell transplantation. Here, we describe differentiation protocols for 3D cultures that produce retinal organoids containing photoreceptors with rudimentary outer segments. These protocols can be used as a model to understand retinal disease mechanisms and test potential therapies, including antisense oligonucleotides (AONs) to alter gene expression or RNA processing. This "retina in a dish" model is well suited for use with AONs, as the organoids recapitulate patient mutations in the correct genomic and cellular context, to test potential efficacy and examine off-target effects on the translational path to the clinic.

Keywords Retinal organoids, Induced pluripotent stem cells, Differentiation, 3D culture, Retinal degeneration, Photoreceptor, Retina in a dish

#### 1 Introduction

The dysfunction and death of photoreceptor cells are associated with inherited retinal diseases (IRDs), which are a major cause of blindness. The lack of effective treatment to prevent loss of photoreceptors means these diseases are currently irreversible. Recent progress in the differentiation of stem cells to retinal cells has enabled the generation of functional retinal organoids in vitro or a "retina in a dish" [1–4]. By recapitulating the retina from patientderived induced pluripotent stem cells (iPSC), retinal organoids offer a platform for developing therapeutic treatments and modeling patient disease [5, 6].

A dynamic and complex microenvironment is involved in eye development, including direct and indirect cell–cell interaction and specific signaling regulation in different stages of development

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_16, © The Editor(s) (if applicable) and The Author(s) 2022

[7]. Because of this complex microenvironment, retinal organoids have the potential to develop a more mature retina than photoreceptors differentiated in 2D conditions only. Several studies have shown that with defined culture conditions, embryonic stem cells (ESC), and iPSC can be differentiated into retinal organoids in a 3D environment, producing a laminated retina that mimics the in vivo human retina [2, 3, 8]. In addition to recapitulating the structure of native eye development, rudimentary disorganized outer segments can be observed in photoreceptors from retinal organoids.

In this chapter, we describe three different methods to differentiate iPSC to retinal organoids in 3D. Retinal organoids generated from these protocols are well laminated with photoreceptors in their outer layer and develop rudimentary outer segments. Importantly, they also recapitulate photoreceptor mRNA processing and the exquisite pattern of alternative splicing they present [9– 11]. This makes retinal organoids ideal for studying aberrant splicing events associated with patient variants in several forms of IRDs [9, 12]. Furthermore, they can then be applied to the development of antisense oligonucleotides (AONs) as potential treatments [9, 13].




	- 2. Neural induction Medium (NIM): Advanced DMEM/F12, 1% N2 supplement, 1% NEAA, 1% Glutamax, and 1% Pen/Strep (see Note 1).
	- 3. Retinal Differentiation Media (RDM): DMEM/F12 (3:1), 1% Pen/Strep, 1% NEAA, and 2% B27 (see Note 1).
	- 4. Neural Retina Maturation Medium 1 (RMM1): DMEM/F12 (3:1), 1% Pen/Strep, 2% B27, 10% FBS, 100 μM Taurine, 1% NEAA, and 1% Glutamax (see Note 1).
	- 5. Neural Retina Maturation Medium 2 (RMM2): DMEM/F12 (3:1), 1% Pen/Strep, 2% B27 (without vitamin A), 1% N2, 10% FBS, 100 μM Taurine, 1% NEAA, and 1% Glutamax (see Note 1).
	- 2. PBS.
	- 3. Micropestle.

#### 3 Methods

Protocol

iPSC are maintained with Essential 8 Flex (E8F) in Geltrex coated 6-well plates (see Note 2). Once they reach 70% confluence, iPSC are treated with 500 μl cell dissociation buffer for 2 min at 37 C in the incubator. After the incubation, remove the cell dissociation buffer and add 1 ml of E8F into a well. Use 1 ml tip scraping the well to collect iPSC in small clumps. Cell clumps are collected and transferred into a new Geltrex-coated plate with 1 ml tip. Medium is changed every other day, and iPSC can be double-fed with 4 ml E8F to cover the weekend (see Note 3).


4. On Day 7, transfer EB from three wells to six wells of Geltrexcoated 6-well plate in NIM, 4 ml in each well. Gently mix the

Fig. 1 EB suspension protocol. Top row: Schematic diagram of EB suspension protocol steps and media. Lower row: Representative organoids at different stages of differentiation are shown. Visible lamination can be observed at approximately Day 35 and good organoids can maintain the lamination and mature during differentiation to form an outer nuclear layer of photoreceptors with inner segment and outer segment (which can be seen by the "brush border," arrowhead). Scale bar is 250 <sup>μ</sup>m. The cartoon images are made with BioRender

Fig. 2 EB adherent protocol. Top row: Schematic diagram of EB adherent protocol steps and media. Lower row: Representative images at different stages of differentiation are shown. EB formation in suspension is followed by attachment to Geltrex-coated wells and formation of NR. Picked NR successfully mature through the differentiation form an outer nuclear layer of photoreceptors with inner segment and outer segment (which can be seen by the "brush border," arrowhead). Scale bar is 250 <sup>μ</sup>m. The cartoon images are made with BioRender

medium in the wells to let the EB equally distributed in the wells (Fig. 2) (see Note 2).


#### 3.3 Non-EB Adherent Protocol This protocol is adapted from the method initially described by Ali and colleagues [8].


#### 3.4 AON Treatment of Organoids 1. Mature organoids are generated from protocols described in the above sections.

2. Dilute AONs into working concentration (e.g., 0.1–10 μM) with culture medium, depending on the methods (see Note 13).

Fig. 3 Non-EB adherent protocol. Top row: Schematic diagram of non-EB adherent protocol steps and media. Lower row: Representative images at different stages of differentiation are shown. NR are formed in NIM medium, and picked NR cultured in suspension going through differentiation mature to form organoids with an outer nuclear layer of photoreceptors with inner segment and outer segment projecting outwards (which can be seen by the "brush border," arrowhead). Scale bar is 250 <sup>μ</sup>m. The cartoon images are made with BioRender


AONs. The assays used for downstream analyses are dependent on the specific questions being asked. Routine analyses would usually involve RT-PCR and qPCR, but the organoids are also amenable to RNAseq, single-cell sorting, next-generation sequencing, or longrange sequencing. This can provide a unique insight into human photoreceptor splicing and its manipulation for discovery science or therapeutic benefit.

#### 4 Notes


#### Acknowledgments

This work is supported by Wellcome Trust, Fight for Sight, Foundation Fighting Blindness, Retina UK, Moorfields Eye Charity and NC3Rs. We would like to thank the other members of the Cheetham, Hardcastle, and van der Spuy groups past and present for their support, encouragement and help in iPSC and organoid maintenance. We would also like to thank Anai Gonzalez-Cordero for advice on the non-EB adherent protocol.

#### References


West EL, Pearson RA, Ali RR (2017) Recapitulation of human retinal development from human pluripotent stem cells generates transplantable populations of cone photoreceptors. Stem Cell Rep 9(3):820–837. https://doi. org/10.1016/j.stemcr.2017.07.022


Cheetham ME (2018) Splice-modulating oligonucleotide QR-110 restores CEP290 mRNA and function in human c.2991+1655A>G LCA10 models. Mol Ther Nucleic Acids 12:730–740. https://doi.org/ 10.1016/j.omtn.2018.07.010

14. Capowski EE, Samimi K, Mayerl SJ, Phillips MJ, Pinilla I, Howden SE, Saha J, Jansen AD, Edwards KL, Jager LD, Barlow K, Valiauga R, Erlichman Z, Hagstrom A, Sinha D, Sluch VM, Chamling X, Zack DJ, Skala MC, Gamm DM (2019) Reproducibility and staging of 3D human retinal organoids across multiple pluripotent stem cell lines. Development 146(1). https://doi.org/10.1242/dev.171686

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 17

# Establishment of In Vitro Brain Models for AON Delivery

#### Elena Daoutsali and Ronald A. M. Buijsen

#### Abstract

Progress in stem cell biology has made it possible to generate human-induced pluripotent stem cells (hiPSC) that can be differentiated into complex, three-dimensional structures, where the cells are spatially organized. To study brain development, Lancaster and colleagues developed an hiPSC-derived threedimensional organoid culture system, termed cerebral organoids, that develop various discrete, although interdependent, brain regions. Here we describe in detail the generation of cerebral organoids using a modified version of the culture protocol.

Key words Cerebral organoid, Disease modeling, Induced pluripotent stem cells

#### 1 Introduction

Many brain disorders are hereditary diseases with a known genetic cause, which allowed scientists to generate animal models to study disease progression, understand disease mechanisms, and perform therapeutic intervention studies [1, 2]. However, (1) mice are different from humans, and it is difficult to translate results from animal experiments into clinical application; (2) the genetic cause of many diseases is not yet known; (3) many disease-causing genes are mainly expressed in the cells that are affected; (4) for many of them, there are no (humanized-)mouse models available; (5) there is governmental and public pressure to advance the development of alternative model systems to replace animal studies. This emphasizes the need for patient-derived disease models that bridge the translational gap between animal models and human clinical trials. Progress in stem cell biology has made it possible to generate human induced pluripotent stem cells (hiPSCs) [3] that can be differentiated into the important cell types of the brain, neurons, and astrocytes [4, 5]. The disadvantage of these 2D models is that they are descriptive at a cellular level, but they fail to adequately provide the details that could be derived from a more complex, three-dimensional structure, where the cells are spatially organized

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_17, © The Editor(s) (if applicable) and The Author(s) 2022

[6]. In 2013, Lancaster and colleagues developed a hiPSC-derived three-dimensional organoid culture system, termed cerebral organoids, that develop various discrete, although interdependent, brain regions [7]. These organoids recapitulate many features of human cortical development, including a progenitor zone organization with abundant outer radial glial stem cells [8].

Here we describe the generation of cerebral organoids using a modified version of the Lancaster protocol [7, 9]. In short, feederfree cultured hiPSCs were dissociated and replated in neural induction medium in a non-adherent cell culture plate, and differentiated for 100 days (Fig. 1). Cryosections of these organoids can be used for immunofluorescence studies. Organoids can be used for many different purposes including disease modeling, studying disease mechanisms, or analyzing therapeutic interventions (using for example antisense oligonucleotides) at any given time point.

#### 2 Materials


3. Bioreactor or shaker.


Fig. 1 Cerebral organoids during the various stages of organoid culturing. Organoids are cultured using a modified version of the Lancaster protocol. After 5 days of neuroectodermal differentiation, the neurospheres are embedded in Matrigel and cultured in the neurosphere medium in a 6-well plate for 5 days. For cerebral organoid maturation, the embedded neurospheres are transferred into a spinner flask and can be used for downstream applications if needed


#### 3 Methods



#### 3.2 Neurosphere Embedding 1. Use an 1-ml micropipette with a wide orifice pipet tip to place the neurospheres on a silicone organoid embedding sheet (see Notes 6 and 7).



Fig. 2 Immunofluorescent staining of a cortical plate structure. Cortical plate structure in cerebral organoids stained with DAPI (blue), the neural progenitor marker PAX6 (green), and the neural marker TUBB3 (red). The scale bar represents 100 <sup>μ</sup>m


#### 4 Notes


#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part IV

In Vivo Model Systems

# Considerations for Generating Humanized Mouse Models to Test Efficacy of Antisense Oligonucleotides

# Irene Va´ zquez-Domı´nguez and Alejandro Garanto

#### Abstract

Over the last decades, animal models have become increasingly important in therapeutic drug development and assessment. The use of these models, mainly mice and rats, allow evaluating drugs in the real-organism environment and context. However, several molecular therapeutic approaches are sequence-dependent, and therefore, the humanization of such models is required to assess the efficacy. The generation of genetically modified humanized mouse models is often an expensive and laborious process that may not always recapitulate the human molecular and/or physiological phenotype. In this chapter, we summarize basic aspects to consider before designing and generating humanized models, especially when they are aimed to test antisense-based therapies.

Keywords Humanized models, Model systems, Mouse model generation, Splicing defects, Antisense oligonucleotides, In vivo drug testing

#### 1 Introduction

Even though in 2020 the Food and Drug Administration (FDA) approved 53 new drugs (their second biggest approval number ever) [1], the mean annual number of drug approvals was 41 between 2010 and 2018 (approximately 15% of all evaluated drugs until 2019) [2]. It could be argued that this limited success is mainly caused by the insufficient recapitulation of the physiological and pathological of human disease in the preclinical phase. Studying human diseases is often difficult due to the restrictions and ethical concerns regarding the manipulation of human samples or tissues and the use of animal models allows the evaluation of drugs within an entire organism. However, these models are different species, with a different DNA sequence and different behavior, that may not recapitulate the complete human phenotype. Among all animal models, mice and rats are the most frequently used models in early preclinical stages. This is due to their relatively small size, ease of handling and maintenance, short reproductive

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_18, © The Editor(s) (if applicable) and The Author(s) 2022

cycle, and the similarities with regard to the genomic and physiological properties compared to humans, as well as the easy genetic manipulation that can be conducted, especially in mice [3].

Antisense oligonucleotide (AON)-based therapies are sequence-specific approaches aimed to either interfere with the splicing mechanism or regulate gene expression, protein translation, or RNA/protein binding [4]. All these processes are highly regulated in cells through different mechanisms such as epigenetics and interactions between mRNA and proteins, which can differ between species [5]. Unfortunately, if the sequence of interest is not completely conserved, the molecule designed to target the human sequence cannot be assessed in the animal model. In this case, the humanization of the models offers the possibility to (partially) replace mouse genes by human counterparts, insert a human copy of the gene in the mouse genome, as well as harbor human tissues [6, 7]. This allows to have the human sequence in an in vivo model. However, before generating such a model, it is crucial to first evaluate if the effect of the mutation of interest is also recapitulated in mouse [8]. In this chapter, we discuss the most important considerations to need to be taken into account before generating a genetically modified humanized mouse model to assess the therapeutic effect of AON molecules.

#### 2 Human and Mouse: Species Are Similar But Not Equal

During the last decades, mouse models have served as valuable organisms for investigating human biology and disease as mouse and human are very similar to each other at the cellular as well as the biochemical level. Furthermore, most of the human cellular pathways are conserved in mouse, at the genetic and molecular level [5]. Despite these similarities, mice and humans can also differ in several aspects at multiple levels. For instance, it is described that around 1% of the human genes do not have an ortholog in mouse [9], which means that not all genes are present in both genomes. Furthermore, some data indicate that there is a higher variability of gene expression between the same tissue of two different species, compared to two different tissues within the same species [5]. These findings support the hypothesis that regulatory pathways, like epigenetics, can be responsible for the interspecies differences [5]. Within epigenetics, histone modifications, such as methylation and acetylation of the promoter region, play a crucial role in expression, leading to most of the differences between both species [10] (Fig. 1a). In addition, differences at the individual gene level give rise to different protein isoforms. Finally, the conservation rate of amino acids generates some differences in protein function or substrate recognition [9] (Fig. 1b). These differences can also cause different response to drug administration [11]. All

Fig. 1 Summary of some aspects that can differ between human and mouse. (a) Epigenetics. Organs in which the gene of interest is expressed are represented in color while those organs in which the gene is not expressed because of epigenetics regulation (such as methylation, me3) are in gray. As the pattern of gene expression between both organisms (inside the blue circle) is different, the functional role of the gene in both may also be different. (b) Amino acid conservation rate. As an example, part of the human and mouse Histone 1 amino acid sequences is represented. Discrepancies between amino acids are indicated with different symbols (: or .) while conserved regions are marked with an asterisk (\*). (c) Mutation effect. Not always the same mutation has the same effect on both species. In humans, the mutation can underlay a pathological defect (left) that is not recapitulated in mice (right). (d) Splicing. The recapitulation of the normal splicing after these discrepancies can make that a defective gene or pathological mutation in humans may have a completely different outcome when the orthologous gene is mutated or truncated in mice [12– 14] (Fig. 1c).

AONs are often used to modulate splicing and correct aberrant splicing. It has been reported that the consensus splice site sequences in mouse and human are highly conserved and comparable even though some small changes in these patterns have been described [15]. That could mean that a human splicing defect may not be completely reproduced in mouse. Some differences between both species at the splicing level have been already described (Fig. 1d). One example is the variant c.315-48T>C in the FECH gene, a modifier mutation for erythropoietic protoporphyria [14]. This mutation generates an aberrant splice acceptor site that results in an intron inclusion of 63 nucleotides upstream of exon 4. However, the generated mouse model not only presents this mis-splicing effect but also a strong skipping of the partially humanized exon 3 [14]. This example illustrates how a hybrid gene (or humanized mouse gene) can have undesirable effects on splicing. Another example is the low recognition of the pseudoexon insertion caused by a deep-intronic mutation in the CEP290 gene (c.2991+1655A>G) in the humanized mouse model carrying this specific variant [13]. Further studies showed that the recognition of the pseudoexon seemed to be specific of primates, and it was less efficient in other species such as pig, dog, mouse, or Drosophila. In addition, as shown in FECH, additional splicing events not detected in humans were found in the humanized Cep290 model [16]. Based on all these studies, we recommend performing several in vitro validation steps before generating a humanized mouse model (Fig. 2).

#### 3 Checkpoints Before Generating a Humanized Mouse Model

3.1 Literature Research An important aspect is to look first in literature or in mouse databases (such as the Mouse Genome Informatics (MGI, http://www. informatics.jax.org/) and the International Mouse Phenotyping Consortium (IMPC, https://www.mousephenotype.org/)) whether any genetically modified animal model for the gene of interest has already been obtained. With this information, one can avoid the generation of a model already available. In addition, if a humanized model is aimed to study disease, the availability of a

Fig. 1 (continued) humanization is indicated on the left. Mouse regions are indicated in purple and human regions in yellow. On the right, the presence of the splicing-related mutation of interest, located in an intron sequence, causes the insertion of a pseudoexon (PS) in the mRNA transcript. In this situation the splicing defect is recapitulated after the humanization

Fig. 2 Schematic representation of the different checkpoints for generating a humanized mouse model

knockout/mutant model will contribute to predict the possible phenotype of the potential new humanized model. For instance, mutations in ABCA4 generate a non-functional protein involved in the visual cycle that leads to retinal degeneration and subsequent loss of vision, causing Stargardt disease in humans [17–19]. However, the Abca4/ mouse model shows a mild late onset phenotype, despite the fact that no ABCA4 protein is encoded [20]. That means that the defect caused by the absence of this protein is not completely comparable to the human phenotype. Thus, depending on the purpose of study, a humanized mouse model may not be suitable to evaluate antisense therapies for a particular gene.

3.2 Comparison of the Gene Sequence If the model is not available, the first question that needs to be answered is whether the gene of interest is present in the mouse genome. The probability is high, since only 1% of human genes do not have a mouse orthologue [9]. In addition, it is important to check the structure of the gene, to ensure it is similar and no relevant exons for the study or protein function are missing. To do so, databases such as Genome Browser (https://genome.ucsc. edu) or Ensembl (https://www.ensembl.org) can be useful. If the gene name does not coincide, a blast (https://blast.ncbi.nlm.nih. gov) at DNA or protein level can be done to search for similar genes or proteins in other species.

3.3 Assessment of Gene Expression in Mouse Although mouse and human are similar at cellular and molecular levels, some discrepancies between both species have been already reported, including variation in expression levels in different tissues. Therefore, assessing that the gene of interest is expressed in the proper murine tissue (or at least the tissue of study) is crucial. For instance, it is already known that one of the most important pathways in cancer and apoptosis, the phosphoinositide 3-kinase (PI3K) signaling cascade [21], shows high differences in the expression of its two key genes, mTOR and AKT2, between both species [22]. Thus, mouse is not the best model to conduct oncological studies that aim to investigate this pathway in vivo. In addition, it has been published that muscle, liver, and neuronal cells show a strong similarity of gene expression profiles between both species while other tissues such as testis, lung, and pancreas showed more differences due to the evolution [22]. Therefore, it is important to check expression levels in the tissue(s) of interest in mouse, to ensure that the generation of the humanized model will be useful for future experiments.

#### 3.4 Humanization Feasibility In general, there are two main ways to introduce the human gene into the mouse genome:


In the first option, the complexity lays on where to exactly insert the gene and how to regulate its expression. The second option requires a more in-depth study on the recombination possibilities, the conservation rate between human and mouse, and the resulting humanized gene and protein. For that, first it is important to identify the mouse exons that correspond to the human ones. It is also important to check whether they encode for the same part of the protein and how conserved it is. This can be done with the previously mentioned freely available databases. Finally, the overall procedure needs to be technically feasible, by either homologous recombination in embryonic stem cells or genome editing techniques using a donor template. These steps need to be discussed with experts on these techniques either at other academic institutes or companies specialized in generating animal models.

3.5 In Vitro Validation As indicated previously, AONs can be used for splicing modulation. Thus, it is important to study whether the splicing machinery in mouse will allow proper:


One way to perform this validation is by using artificial systems (e.g., mini-/midigenes) harboring the human region of interest either with the "wild-type" sequence or the mutated one [16]. In that way, it can be assessed whether they behave in a mouse-specific environment as expected based on observations in humans. For that, conventional mouse-derived cell lines can be a good start [23]. The data obtained from these studies are crucial to assess if the mouse might be a good animal for the generation of a humanized model.

3.5.1 In Vitro Splicing Assays Considerations The easiest way for a first screening is to develop human mini-/ midigenes with and without the mutation [24] and deliver them to conventional mouse cell lines easy to transfect, such as iMCD3, B16-F10 or N2A. However, depending on the tissue of interest, this information might not be completely accurate. For instance, in human cells, we and others have found differences in pseudoexon recognition, depending on how similar the molecular background was compared to the one of the cell/tissue of interest [24–26], probably due to tissue-specific splicing regulatory elements [27]. Therefore, it is important to choose an appropriate cell line. For instance, if the gene is implicated in neurological disorders affecting the brain, the mouse cell line N2A (derived from mouse brain) might be a better option than using B16-F10, derived from mouse skin or iMCD3, derived from kidney [28]. It is always good to test in several different cell lines: if the splicing pattern observed in humans is well recapitulated in multiple cell lines, most probably the humanized model will also show the same splicing patterns. In case the splicing pattern is not recapitulated at all, we recommend to stop with the generation of the humanized mouse model and try to find alternative models.

> Next step, if the splicing defect is recapitulated, is to determine whether the AONs of interest previously designed and tested in vitro human models can also rescue the splicing in the murine molecular background. With that, it is possible to evaluate if those molecules will also be successful in the mouse molecular background. The generation and validation of mini-/midigenes and AON rescue can be done in 3–4 weeks as previously reported

[24, 29] and can contribute significantly to reduce the number of animals by performing initial screenings in vitro [24].

If all the aforementioned points are met successfully (Fig. 2), the chances of success are higher. However, it can still happen that even though the molecular defect is recapitulated, the model does not show a similar phenotype. Unfortunately, this is difficult to predict in vitro. In addition, in order to discern whether the humanization of the model may influence the potential phenotype, especially in those cases where a mutation has been introduced, it is strongly advisable to generate two humanized models, one with only the human "wild-type" region and another one with the mutated sequence.

#### 4 Current Examples of Humanized Mouse Models

Different approaches have been used to generate humanized mouse models, ranging from introducing the entire gene or part of it or replacing specific parts the gene by the human counterpart. Some of these models have also been used to test AON-based therapies for different types of disease.

4.1 Insertion of the Entire Human Gene One of the approaches frequently used is the introduction of the entire human gene into the mouse genome. This method also takes into account the endogenous expression of the corresponding mouse gene and whether it can affect the development of the desired phenotype. One example of this is the humanized Tg32 FcRn mouse, which harbors the complete human FCGRT gene (around 11 kb) plus 5<sup>0</sup> and 3<sup>0</sup> untranslated regions (around 10 kb). This transgene is introduced into C57BL/6J oocytes with a null mutation in the FcRn mouse gene, which is involved in IgG and serum albumin turnover. As a result, only the human FcRn is expressed under the control of its natural human promoter. Besides introducing the "wild-type" sequence in the mouse genome, Anderson and colleagues also generated a defective allele [30]. In this, a neomycin-resistance cassette was inserted, replacing the human "wild-type" located between the promoter end and the beginning on exon 2. This study revealed that the FcRn+/ model recapitulated the "wild-type" condition indicating that only one allele is sufficient to regulate the IgG levels and transport [31–33]. However, the defective mouse model (FcRn/) successfully recapitulated the defective neonatal transport phenotype [30]. This model has already been used in drug preclinical studies [34]. Most of these studies focused on the evaluation and the pharmacokinetics and pharmacodynamics of human IgG [35], but it may also be useful for other therapeutic approaches, such as AONs [36].

Retinitis pigmentosa (RP) is the most frequent inherited retinal disease, for which no therapy is currently available. In this disorder, most of the studies have been focused on the mutation p.P23H of rhodopsin (RHO), a recurrent mutation causing autosomal dominant RP [37, 38]. To study this mutation, several animal models have been developed, including the generation of humanized models in either mouse [39] or rats [37, 40]. In all of them, the transgene segregated in an autosomal dominant allele manner and the lines were maintained within the murine wild-type rhodopsin background (Rho+/+). However, they differed in the expression levels of the murine and human rhodopsin mRNA. These studies revealed that a higher number of copies of the mutant allele induces a more severe RP phenotype, but also that overexpression of the wild-type allele has a detrimental effect, causing similar RP abnormalities [39, 41–43]. Some of these models have been used to successfully test AON-based therapies aiming to degrade the mutated transcript [44]. These and other results led to a recently started clinical trial (NCT04123626, http://www.clinicaltrials. gov).

AON-based therapies have had a major development in neuromuscular disorders [45]. Several mouse models recapitulating muscular dystrophy phenotypic traits have been generated. For instance, the mdx model, which harbors a nonsense mutation in exon 23 of the mouse Dmd gene causing loss of the functional protein expression, presents a moderate-severe phenotype with an early onset of skeletal muscle degeneration and impairment in muscle functions [46]. However, this model can only be targeted with mouse-specific AONs to the mutation in exon 23 [47–49]. To solve this limitation, the hDMD/mdx mouse model was generated by crossing the mdx model with a model carrying the entire human DMD gene (hDMD), allowing the expression of the "wild-type" human dystrophin in the mdx model [50]. However, the expression of hDMD hampered the development of the dystrophic phenotype caused by the mutation present in the mouse Dmd gene. In 2018, the group of Prof. Aartsma-Rus overcame these limitations by generating a new humanized mouse model (del52hDMD/mdx mice). This model carries both human and murine DMD genes with pathogenic mutations previously described. As a consequence of the nonsense mutation in exon 23 in the mouse Dmd gene and the deletion of exon 52 in the hDMD gene, none of the genes produce functional dystrophin protein [51]. The absence of dystrophin expression caused a more severe muscular dystrophy phenotype than the one observed in other models. As a result, this model was suitable to test human-specific AONs, and thereby improve AON-based therapies for Duchene muscular dystrophy patients by enabling studies at the mRNA, protein and functional level.

#### 4.2 Replacing Part of the Mouse Gene by the Human Gene

Another way to generate a humanized mouse model is by replacing a specific region in the mouse genome by the human counterpart. One example is the humanized mouse model generated to study the mutation c.2991+1655A>G in the CEP290 gene. This mutation is the most recurrent mutation underlying Leber congenital amaurosis, accounting up to 15% of all cases in some populations [25, 52]. Two mouse models were generated in which the mouse sequence from exon 25 to exon 26 was replaced by the corresponding human exon 26, intron 26 (with or without harboring the mutation of interest), and exon 27 [13]. In the model Cep290lca/lca, human intron 26 harbors the mutation of interest while the model Cep290hum/hum does not. In both the models, Cep290 expression levels and regular splicing of the gene were maintained when compared to "wild-type" mice. However, the splicing defect was barely present in the Cep290lca/lca model, the retina being the only tissue where the exon was recognized at detectable levels, but by far not enough to lead to the human phenotype [25]. Subsequent studies, revealed that the pseudoexon recognition strongly correlates with evolutionary distance, being highly recognized in primates, and hardly recognized in lower species such as rodents and fly [16].

#### 5 Conclusions

In summary, the suggestions given in this chapter highlight the importance of performing an accurate study to increase the chances of success when generating a humanized mouse model, e.g., to assess AON-based molecules. Despite similarities, human and mouse are different species and therefore differences are expected at all levels. When aiming to generate a humanized model, it is crucial to determine first if the gene of interest is present in the genome, has a similar structure, and if it is expressed in the same tissue(s) as in humans. Subsequently, when working with splicing defects, it is important to validate that these will be recognized in the murine molecular background, which can be easily done using "artificial systems" in vitro. If everything is conserved, the technical part is also important, is it feasible to humanize the model? If so, the chances of success are high. However, in some cases, it is not feasible, or a mouse model is not an option. In those cases, other models can be explored such as zebrafish or cellular models based on organ-on-a-chip technology. Following these suggestions may increase the success of the generation of humanized models and reduce the number of animals used in research. However, although all these in silico and in vitro tests can help to predict the recapitulation of a molecular defect in a humanized model, it is important to keep in mind that such models may not recapitulate the entire human phenotype.

#### Acknowledgments

We thank Dr. Rob W.J. Collin for the critical reading of this chapter. The group is financially supported by the Foundation Fighting blindness (PPA-0517-0717-RAD to A.G.), the Curing Retinal Blindness Foundation (to A.G.) as well as the Landelijke Stichting voor Blinden en Slechtzienden, Stichting Oogfonds Nederland (who contributed through UitZicht 2019-17), together with the Rotterdamse Stichting Blindenbelangen, Stichting Blindenhulp, and Stichting Dowilvo (to A.G). The funding organizations had no role in the design or conduct of this research. They provided unrestricted grants. All the figures were made with BioRender.

#### References


Genome Res 15(1):111–119. https://doi. org/10.1101/gr.3108805


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Generation of Humanized Zebrafish Models for the In Vivo Assessment of Antisense Oligonucleotide-Based Splice Modulation Therapies

### Renske Schellens, Erik de Vrieze, Ralph Slijkerman, Hannie Kremer, and Erwin van Wijk

#### Abstract

Antisense oligonucleotide (AON)-based splice modulation is the most widely used therapeutic approach to redirect precursor messenger RNA (pre-mRNA) splicing. To study the functional effect of human mutations affecting pre-mRNA splicing for which AON-based splice redirection would be a potential therapeutic option, humanized knock-in animal models are pivotal. A major limitation of using humanized animal models for this purpose is the reported poor recognition of human splice sites by the splicing machineries of other species. To overcome this problem, we provide a detailed guideline for the generation of functional humanized knock-in zebrafish models to assess the effect of mutation-induced aberrant splicing and subsequent AON-based splice modulation therapy.

Keywords Pre-mRNA splicing, Species-specific minigene splice assay, Antisense oligonucleotides, Inherited retinal dystrophies, Usher syndrome, Zebrafish

#### 1 Introduction

Precursor messenger RNA (pre-mRNA) splicing is a tightly regulated and complicated process. The spliceosome, a multicomponent protein complex, removes introns from the pre-mRNA by recognizing and joining each splice donor site, located at the 50 end of the intron, to its corresponding splice acceptor site, located at the 3<sup>0</sup> end of the intron. The remaining exons are subsequently fused together to form mature mRNA. However, genetic variants in both introns and exons can affect the process of splicing and result in a range of pathogenic phenotypes.

For inherited retinal dystrophies (IRDs), it has been estimated that ~20% of all identified mutations affect pre-mRNA splicing [1]. Numerous of those mutations have already been described in literature [2–4], of which the recurrent deep-intronic

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_19, © The Editor(s) (if applicable) and The Author(s) 2022

c.7595-2144A > G mutation in USH2A intron 40 is an example [5]. This mutation creates a strong splice donor site resulting in the incorporation of a pseudoexon (PE40) into all USH2A transcripts and as a consequence, the premature termination of usherin translation. Loss of usherin function in man results in either combined hearing and vision loss (Usher syndrome) or non-syndromic vision loss (retinitis pigmentosa).

Currently, the most widely used approach to interfere with premRNA splicing is antisense oligonucleotide (AON)-based splice modulation. AONs are small single-stranded RNA molecules designed complementary to their target pre-mRNA. Upon binding to splicing regulatory elements within the pre-mRNA transcript, AONs are capable of correcting aberrant pre-mRNA splicing, inducing the skipping of (pseudo)exons or promoting the inclusion of native exons [6].

Humanized animal models, in which a specific part of the species' genomic DNA is replaced by the orthologous human sequence, are pivotal to study the functional effect of aberrant splicing and AON-based splice correction therapy. Multiple vertebrate model organisms, each having their pros and cons, are currently being used for IRD-associated translational research purposes. Among those, the zebrafish represents a fast and inexpensive model which has proven its strength in studying IRDs. Zebrafish are easy to genetically manipulate, have a retinal structure comparable to humans, and for most IRDs (including Usher syndrome) robust phenotypes can already be detected at 5 days post fertilization (dpf) [7].

A major difficulty of using humanized knock-in animal models for the purpose of studying aberrant splicing and evaluating AON-based splice correction is the reported poor recognition of human splice sites by the splicing machineries of other species [8, 9]. As such, thorough in silico bio-informatic analyses and in vitro cell-based splice assays are essential for the development of a humanized animal model that properly recognizes the splice sites of the introduced human sequence. In this chapter, we will discuss the step-by-step procedure to generate functional humanized zebrafish models, including optimization in cross-species splice-site recognition, to assess the effect of mutation-induced aberrant splicing and subsequent AON-based splice redirection therapies.

#### 2 Materials

2.1 In Silico Splice Site Analysis 1. A computer with internet access and a web browser.




3. Taq polymerase mix.


Fig. 1 Optimization of human USH2A PE40 splice acceptor site for improved recognition by the zebrafish splicing machinery. (a) The nucleotide distribution of the splice donor site found in man (Hs) and zebrafish (Dr). The PE40 splice donor sequence is presented under the splice donor sites with the c.7595-2144A <sup>&</sup>gt; G mutation, indicated by an asterisk. (b) Similar as in (a), the PE40 splice acceptor sequence is presented under the consensus splice acceptor sites as used in human (Hs) and zebrafish (Dr). Based on comparisons, the M1 site was selected for optimization, predicted to result in a stronger PE40 splice acceptor site. The predicted strength of the splice acceptor site is indicated in brackets. (c) A zebrafish-specific minigene splice assay containing the human USH2A PE40 with flanking sequences was generated. The plasmids, either containing the c.7595-2144A <sup>&</sup>gt; G mutation (mut) or not (wt), further contained M1 (indicated by vertical arrow). (d) The effect of the introduction of M1 on recognition of human USH2A PE40 after expression in zebrafish cells (Zendo-1) determined by RT-PCR. untr.: untransfected cells, PCR(): negative template PCR control. (Reproduced from Slijkerman et al. [9], with permission from Mary Ann Liebert, Inc.)

> 6. Extract plasmid DNA from the bacterial cultures using a Plasmid DNA extraction kit.


3.3 Validation of the Optimized Splice Site Using an Engineered Zebrafish-Specific Minigene Splice Assay <sup>234</sup> <sup>g</sup> for 5 min. Isolate total RNA using a RNA isolation kit, according to manufacturer's instructions.


3.4 Guide RNA Design for CRISPR/ Cas9 Injection

3.5 Generation of Donor Template for CRISPR/Cas9-Induced Homology Directed DNA Repair

	- 2. Prepare a microinjection plate by casting a 1% agarose gel using a plastic mold that produces six 1.5-mm-wide trenches (see Note 26).
	- 3. Prepare the injection mixture by combining Cas9 protein (800 ng/μl), sgRNA (100 ng/μl), potassium chloride (0.3 M), phenol red (0.1%), and donor template DNA (25 pg) (see Note 27). Incubate injection mixture at 37 -C for 5 min prior to injections (see Note 28).
	- 4. Perform injections by using a Pneumatic Picopump pv280 microinjector, a pipette holder, a foot pedal, and a stereoscope or similar (see Note 29) (see Note 30). Load the injection needle with injection mixture and calibrate the needle by

3.6 Generation of the Humanized Zebrafish Line

adjusting the pressure and time settings of the microinjector until the injection volume is 1 nl (see Note 31).


#### 3.7 Genotyping After Fin Clipping 1. Prepare for each adult fish one PCR tube containing 75 μl of lysis buffer and one single box containing 1 l of fresh water.


which the reaction is cooled down to 16 -C until further processing.

	- 3. Dilute your synthesized cDNA 2–10 (depending on gene expression levels) and use it as a template for PCR amplification. Typically, one PCR (20 μl) contains dNTPs (final concentration of 0.2 mM for each nucleotide), forward and reverse primers (final concentration of 0.2 mM for each primer), 1 μl 2–10 diluted cDNA, 0.25 μl (0.5 units) DNA polymerase, and 4 <sup>μ</sup>l 5 reaction buffer (see Note 6). Cycling conditions are: 98 -C for 2 min, followed by 35 cycles with 98 -C for 15 s, "Tm" -C for 20 s, and 72 -C for 45 s. The "Tm" temperature depends on the primers used and should be optimized on beforehand. The reaction is finalized by a 5-min incubation step at 72 -C after which the reaction is cooled down to 16 -C until further processing.
	- 4. Visualize human (pseudo)exon inclusion by analyzing the amplified PCR products on an agarose gel. Figure 2a shows an example of increased in vivo recognition of the (pseudo) exon after optimization of the splice site.

3.8 Visualization of Human (Pseudo)Exon Inclusion in Zebrafish

3.9 Quantification of Human (Pseudo)Exon Inclusion Using Quantitative RT-PCR (RT-qPCR)

Fig. 2 Human USH2A PE40 incorporation in zebrafish ush2a transcripts. (a) The level of human USH2A PE40 incorporation into the zebrafish ush2a transcript is analyzed by RT-PCR using cDNA derived from ush2 <sup>a</sup>PE40 + M1/PE40 + M1, ush2aPE40/PE40, ush2ahum/hum, and wild-type larvae (5dpf). Beta actin amplification is shown as loading control (lower panel). (b) Quantitative RT-PCR analyses to determine the absolute number of PE40-containing ush2a transcripts in ush2aPE40 + M1/PE40 + M1, ush2aPE40/PE40, ush2ahum/hum, and wild-type larvae (5dpf). Data are expressed as percentage of PE40-inclusion (mean SD). Two-tailed unpaired Student's <sup>t</sup>-test revealed significant differences between groups ( <sup>p</sup> <sup>¼</sup> 0.0001). ND not detectable


of the exon-inclusion amplicon and of the exon-exclusion amplicon. The amount of product expected from splice site optimization can be calculated as percentage of total ush2a transcripts (Fig. 2b). The latter can be calculated as the sum of wild-type product and product expected from splice site optimization (see Note 41).

#### 4 Notes


microinjection since it results in a higher injection speed and accuracy.


#### Acknowledgments

The authors would like to thank Dr. Jeroen den Hertog for sharing his zebrafish cell line Zendo-1. The authors acknowledge the facilities and personnel of the Zebrafish Facility at the Faculty of Science of the Radboud University Nijmegen. The authors received funding from the Foundation Fighting Blindness USA (FFB PPA-0517-0717-RAD to EvW) and Stichting Ushersyndroom (to EvW and EdV) for the research that led to the protocol.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Use of Small Animal Models for Duchenne and Parameters to Assess Efficiency upon Antisense Treatment

### Ngoc Lu-Nguyen, Alberto Malerba, and Linda Popplewell

#### Abstract

Duchenne muscular dystrophy (DMD) is a rare genetic disease affecting 1 in 5000 newborn boys. It is caused by mutations in the DMD gene with a consequent lack of dystrophin protein that leads to deterioration of myofibers and their replacement with fibro-adipogenic tissue. Using antisense oligonucleotides (AONs) to modify out-of-frame mutations in the DMD gene, named exon skipping, is currently considered among the most promising treatments for DMD patients. The development of this strategy is rapidly moving forward, and AONs designed to skip exons 51 and 53 have received accelerated approval in the USA. In preclinical setting, the mdx mouse model, carrying a point mutation in exon 23 of the murine Dmd gene that prevents production of dystrophin protein, has emerged as a valuable tool, and it is widely used to study in vivo therapeutic approaches for DMD. Here we describe the methodology for intravenous delivery of AONs targeting dystrophin through tail vein of mdx mice. Furthermore, the most relevant functional analyses to be performed in living mice, and the most informative histopathological and molecular assays to evaluate the effect of this treatment are detailed.

Key words Antisense oligonucleotides, Duchenne muscular dystrophy, Dystrophin, Exon skipping, mdx

#### 1 Introduction

Duchenne muscular dystrophy (DMD) is due to mutations on the DMD gene and the consequent loss of the dystrophin protein in skeletal and cardiac muscles [1]. The loss of dystrophin makes the muscle fibers extremely fragile and susceptible to cycles of tissue degeneration and regeneration causing depletion of muscle stem cells and massive deposition of fat/connective tissue finally leading to muscle weakness, respiratory insufficiency, and cardiac failure [2]. Mutations of the DMD gene often lead to the disruption of the reading frame. Exon skipping, based on the selective removal of exons flanking the out-of-frame mutations using antisense oligonucleotides (AONs), has been used to re-frame the mRNA transcript to allow the expression of a partially functional Becker

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_20, © The Editor(s) (if applicable) and The Author(s) 2022

muscular dystrophy (BMD)-like protein [3, 4]. To study this approach in vivo, AONs targeting exon 23 of the mouse Dmd gene have been tested by local and systemic delivery in mdx mice, the most used animal model of DMD, from neonatal to very late (>15-month old) stages of development [5–8].

In this chapter, we focus on systemic intravenous administration of AONs that allows their body-wide distribution. We provide a detailed description of intravenous infusion of AONs through the mouse tail vein. Phosphorodiamidate morpholino oligomers (PMOs) with or without the conjugation with a cell-penetrating peptide have been used for this approach [4–8]. We also list methods for assessing the animal locomotor activities, forelimb strength, treadmill and wheel exercise, and in situ muscle physiology. Postmortem protocols for RT-PCRs measuring the level of exon skipping for DMD exon 23, Western blot for quantifying the level of dystrophin protein, and immunofluorescence for evaluating muscle histology and the effect of the treatment on muscle size, muscle fibrosis, and myofiber cross-sectional area are additionally described.

#### 2 Materials



#### 2.4 Postmortem

#### Tissue Processing


#### 2.5 RNA Extraction and RT-PCR



#### 2.6 Protein Extraction and Western Blot



#### 2.7 Immunohistochemistry Staining and Histological Analyses


#### 3 Methods


	- 1. Behavioral and locomotor measurements using open field animal activity monitoring system (SOP ID No.: DMD\_M.2.1.002).
	- 2. Use of treadmill and wheel exercise to assess dystrophic state (SOP ID No.: DMD\_M.2.1.003).
	- 3. Use of grip strength meter to assess the limb strength of mdx mice (SOP ID No.: DMD\_M.2.2.001).
	- 4. Measuring isometric force of isolated mouse skeletal muscles in situ (TREAT-NMD, SOP ID No.: DMD\_M.2.2.005).
	- 1. Weigh mouse before sacrificing by CO2 exposure and confirm the death by neck dislocation.
	- 2. From each mouse, we usually collect heart, diaphragm, extensor digitorum longus, gastrocnemius, soleus, and tibialis anterior.
	- 3. Tissues from one side of the body are frozen immediately in liquid N2 for RNA and protein extraction while tissues from the other side are embedded in OCT and subsequently frozen in liquid N2-cooled isopentane for cryo-sectioning (see Note 5).
	- 4. All samples are kept at -80 C until use.
	- 5. If required, collect total blood by cardiac puncture into 1-ml syringe (see Note 6). Transfer the blood into an ice-cold 1.5-ml tube, keep on ice for at least 4 h or overnight prior to serum extraction: (1) centrifuge at 8000 <sup>g</sup> for 10 min at 4 C; (2) transfer supernatant to fresh tubes; (3) repeat centrifugation; (4) collect the supernatant and store at -80 C.

#### 3.3 Postmortem Tissue Processing

3.4 RNA Extraction and RT-PCR Quantifying Exon Skipping Efficiency


3.5 Protein Extraction and Western Blot Quantifying Dystrophin Expression


3.6 Immunostaining for Dystrophin, Laminin, or Collagen VI

For histological analyses, muscles are sectioned and co-immunostained for dystrophin/laminin to assess the percentage of dystrophin positive fibers and the muscle cross sectional area or immunostained for collagen VI to assess the amount of muscle fibrosis.


3.7 Picro Sirius Red Staining for Detecting Collagen I and III

Deposition of muscle fibrosis can also be assessed following staining with picro sirius red that stains collagen in red and cytoplasm in yellow:


#### 3.8 Histological Analyses

	- 2. Ideally a mosaic image of the whole muscle section is acquired by overlapping and stitching individual images of that section together.
	- 3. For quantifying dystrophin expression: use ZEN software to score the mean intensity of dystrophin and subsequently normalize to the mean intensity of laminin of the same section.
	- 4. For quantifying dystrophin-positive fiber numbers: use ImageJ, manually count the number of positive fibers, and evaluate as the percentage of the number of total fibers within the same image field that are positive with laminin staining (see Note 18).
	- 5. For quantifying the frequency distribution of fiber size: see TREAT-NMD, SOP ID No.: DMD\_M.1.2.001.
	- 6. For quantifying the level of central nucleation: use ImageJ, manually count the number of fibers having centralized nuclei (as shown by DAPI staining) and evaluate as the percentage of the number of total fibers within the same image field that are positive with laminin staining.
	- 7. For quantifying muscle fibrosis: use ZEN software to score the mean intensity of collagen VI and evaluate as the percentage of wild-type values obtained in the same way. For measuring the collagen area positive with collagen VI or sirius red staining: use ImageJ and select the area covered by the section. Then using the threshold function, highlight only the portion of the section positive for collagen staining and evaluate as the percentage of the total area of the muscle cross-section (see Note 19).

#### 4 Notes


weekly for several weeks to boost the antisense efficacy and make a more robust exon skipping.


PBS, 0.05% Tween-20 can be used to dilute primary and secondary antibodies.


#### Acknowledgments

The authors thank the Muscular Dystrophy UK (London, UK) for funding the study (grant reference number: RA/893) and Sarepta Therapeutics Inc. (Cambridge, Massachusetts, USA) for their sponsorship. The authors declare that they have no conflict of interest.

#### References


8. Lu-Nguyen N, Ferry A, Schnell FJ et al (2019) Functional muscle recovery following dystrophin and myostatin exon splice modulation in aged mdx mice. Hum Mol Genet 28(18): 3091–3100

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 21

# In Vivo Models for the Evaluation of Antisense Oligonucleotides in Skin

### Jeroen Bremer and Peter C. van den Akker

#### Abstract

Here, we describe an in vivo model in which antisense oligonucleotides were preclinically evaluated in reconstituted patient and healthy control skin. The aim was to investigate the effect of antisense oligonucleotides upon local or systemic administration. This allows for clinically relevant evaluation of antisense oligonucleotides in an in vivo setting. In this model, primary human keratinocytes and fibroblasts were placed into silicone grafting chambers, implanted onto the back of athymic nude mice. After sufficient cells were expanded, within a few weeks, human skin grafts were generated with a high success rate. These mice bearing grafts were subsequently treated with antisense oligonucleotides targeting exon 105 of the COL7A1 gene which encodes type VII collagen. Patients completely lacking expression of type VII collagen develop severe blistering of skin and mucosa, i.e., recessive dystrophic epidermolysis bullosa. In this chapter, we describe the in vivo model used for the preclinical evaluation of antisense oligonucleotides as therapeutic approach for recessive dystrophic epidermolysis bullosa.

Keywords Epidermolysis bullosa, Therapy, Exon skipping, Antisense RNA, Skin equivalent mouse model, Splice modulating

#### 1 Introduction

Antisense oligonucleotide (ASO)-mediated exon skipping has been shown to have great potential as therapeutic approach for the devastating heritable skin blistering disease dystrophic epidermolysis bullosa (DEB) [1, 2]. Severe recessive DEB (RDEB-gen sev) is caused by biallelic null variants in the COL7A1 gene, which encodes type VII collagen (C7). C7 is an extracellular matrix molecule that secures attachment of the epidermis to the dermis by the formation of anchoring fibrils. Imaginably, the complete absence of C7 results in severe blistering of skin and mucosa and early demise [3]. The aim of ASO-mediated exon skipping is to remove the exon from the transcriptome in which the disease-causing variant resides. As a result, a slightly shorter protein is expressed which is functional [4]. In this chapter, we describe the evaluation of ASO targeting

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_21, © The Editor(s) (if applicable) and The Author(s) 2022

skin in an in vivo setting. The model is based on the cell sorting principle, in which cells are of the same cell type cluster; in this case, primary human fibroblasts and keratinocytes [5]. Athymic nude mice were used for several reasons, foremost, these mice are highly suitable for xenografting as the mice do not possess a thymus and therefore do not display graft rejection. Second, morphological changes, like pigmentation or blistering, can be easily observed as the mice have little to no fur. These mice are bred and kept in individually ventilated cages (IVC), as they are vulnerable to pathogens.

Initially, a submerged culture was established in silicone grafting chambers that were implanted onto the back of these mice. This submerged culture allows for proliferation, clustering, and attachment of fibroblasts and keratinocytes onto the muscle fascia. After 10 days, the silicone grafting chamber is removed, which creates an air–liquid interface. It is well known that an air–liquid interface is essential for differentiation of the keratinocytes and form a stratum corneum. These principles are similar to the principles of 3D skinequivalent culture in vitro.

The major advantage of this model is the ability to evaluate ASOs that are systemically administered. Additionally, this model allows treatment for longer periods than in vitro. The major disadvantage of this model is the need for, in comparison with in vitro 3D-culture, high numbers of patient cells. Primary cells were used; however, cell lines or iPSC-derived cells could be used as well. When using primary cells, the lower the passage, the better-defined strata of skin as the proliferative potential and differential is the highest. In this chapter, we describe the grafting model that was used to evaluate ASO-mediated exon skipping upon systemic treatment using primary cultured fibroblasts and keratinocytes.

#### 2 Materials


#### 3 Methods


Fig. 1 Grafting procedure. In this figure, the steps of the grafting procedure are explained. (a) First, a fullthickness skin excision is made. (b) Then the silicone grafting chamber is implanted, followed by injection of the cells in grafting medium in the chamber. (c) Subsequently, the chamber will be filled with wound fluid and after (d) 7 days, the grafting chamber is removed. (e) A scab will be formed on the wound which will (f) fall off around 10 days after removal of the chamber and a patch of human skin is revealed


#### 3.3 Removal of the Ten days post operation, the grafting chambers are removed.


#### 4 Notes

Grafting Chambers


#### References


DF, Palisson F, Schwieger-Briel A, Sprecher E, Tamai K, Uitto J, Woodley DT, Zambruno G, Mellerio JE (2020) Consensus reclassification of inherited epidermolysis bullosa and other disorders with skin fragility. Br J Dermatol 183(4): 614–627. https://doi.org/10.1111/bjd.18921


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Delivery of Antisense Oligonucleotides to the Mouse Retina

#### Alejandro Garanto

#### Abstract

The eye is the organ in charge of vision and, given its properties, has become an excellent organ to test genetic therapies, including antisense oligonucleotide (AON) technology. In fact, the first AON receiving FDA and EMA approval was meant to treat an eye condition. Currently, dozens of clinical trials are being conducted for a variety of subtypes of inherited retinal disease. Although most of them are based on gene augmentation therapies, a phase 3 and two phase 1/2 clinical trials using AONs are ongoing. Since the retina is a layered structure of nondividing cells, obtaining human retinal tissue and expanding it in the lab is not possible, unless induced pluripotent stem cell technology is used. Mouse models have helped to elucidate the function of many genes, and the retinal structure is quite similar to that of humans. Thus, drug delivery to the mouse eye can provide valuable information for further optimization of therapies. In this chapter, the protocol for intravitreal injections of AONs is described in detail.

Keywords Retina, Intravitreal injection, Mouse, Inherited retinal diseases, Antisense oligonucleotide, Drug delivery, Intraocular injection

#### 1 Introduction

The eye is the window to the world around us. Vision occurs thanks to a thin neuronal layer located at the back of the eye called the retina. When light enters the eye, the photosensitive cells (photoreceptors) capture the photons and convert them into chemical and electrical signals that travel to the brain, where the final image will be generated [1].

Inherited retinal diseases (IRDs) are monogenic disorders affecting approximately 1 in 3000 individuals worldwide [2]. So far, mutations in more than 250 genes have been associated with IRDs (RetNet: https://sph.uth.edu/retnet/). IRDs are highly heterogeneous, but in general, the first symptoms appear during the first decade of life, and the disease progresses with age, often leading to total blindness.

Although no cure is available for IRDs, the eye is at the forefront of the development of molecular therapies. Its accessibility, containment, immune-privileged status, together with the window

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_22, © The Editor(s) (if applicable) and The Author(s) 2022

of opportunity that IRDs present, make the eye a model organ for therapeutics. An example is Luxturna, an adeno-associated virus (AAV)-based therapy, which has become the first gene therapy treatment with market approval for an eye disease [3]. This achievement has sped up the development of many other potential treatments currently under investigation at both preclinical and clinical levels for a variety of retinal diseases. Another example of molecules that have shown promising results in humans is antisense oligonucleotides (AONs). These 18–21 nucleotide oligonucleotides are able to bind to the pre-mRNA and, among other functions, can modulate splicing or degrade RNA transcripts [4]. After the promising results obtained in a phase 1/2 clinical trial [5], a phase 2/3 clinical trial (NCT03913143) using AONs to correct a splicing defect in CEP290-associated IRD has recently started. Furthermore, two other phase 1/2 clinical trials have also commenced for specific mutations in the IRD-causing genes USH2A (NCT03780257) and RHO (NCT04123626).

When considering testing AON efficacy in the mouse eye, it is important to remember that AONs are sequence-specific molecules. In the past, we developed a humanized mouse model for the intronic variant c.2991+1655A>G in CEP290 [6]. This model contained a ~6 kb fragment of the human CEP290 including the intronic mutation, and the splicing defect associated with this mutation was only slightly recapitulated [6]. Despite, using a previously efficacious AON characterized in patient-derived cells [7], a single dose of 60 μg of AON was delivered to the retina intravitreally, and pseudoexon skipping was observed up to 30 days after injection [8].

Delivery to the eye can be performed in multiple ways. The most common ones are intravitreal and subretinal injections. For AONs, intravitreal injections are a good choice since the entire retina can be targeted (Fig. 1). There are several ways to perform intravitreal injections, and in this chapter, the protocol we successfully use in our lab is described step by step. For this procedure, no sophisticated equipment is required.

#### 2 Materials

	- 2. Stereomicroscope.
	- 3. Balance to weigh the animals.
	- 4. Microdissection scissors (e.g., Vannas 8 cm str).
	- 5. Dumont #5 forceps.
	- 6. Dumont #5 curve 45-degree forceps.

Fig. 1 Schematic representation of an intravitreal (left) and subretinal (right) injections. The figure was made with Biorender


#### 2.2 For Harvesting Eyes

	- 2. 2-ml Eppendorf tubes.
	- 3. Scissors.
	- 4. Small blades.
	- 5. OCT compound.
	- 6. Molds for cryoblocks.
	- 7. Liquid nitrogen.
	- 8. Polystyrene box.
	- 9. Isopentane.
	- 10. Aluminum boxes (to store the cryoblocks individually (14.5 mm) or in groups (29 mm)).

#### 3 Methods



#### 3.3 The Procedure 1. Take the first animal.


Fig. 2 Schematic representation of the steps to perform an intravitreal injection in mice. (a) The 30G needle is used to make a small hole in the eye to be able to introduce the Hamilton needle with a blunt end in the intravitreal space. It is important to only perform the incision with the very tip part of the needle (marked with an asterisk). (b) Schematic representation of the intravitreal injection of the left eye. The figure was made with Biorender

The cut should be done in the area behind the ora serrata, to make sure we inject in the retina.

	- 2. Check that all animals are awake.
	- 3. Place them back in the corresponding rack and recheck again that labels are correctly placed.
	- 4. Clean the needles first with MQ water (to remove any kind of possible tissue left), then with 70% ethanol, and then with MQ water again and allow to dry (see Note 19).
	- 5. Discard all the 30G and 33G needles used in the proper container.
	- 6. Clean the scissors and forceps with 70% ethanol and water.
	- 7. Discard the bench diaper, the tip, and syringes used to inject carprofen.
	- 8. Clean the working space.

3.6 Harvesting the Tissue and Read-Outs Depending on the molecule injected and the purpose of the experiment, the harvesting time will vary. For instance, it is known that when injecting viruses such as adeno-associated viruses (AAVs), the highest expression is obtained after 3–4 weeks [9]. In contrast, the effect of an AON can be already detected a couple of days after injection [8]. Therefore, the timing on tissue collection will be based on the specific project and the expected efficacy of the therapeutic molecule.

> The way to sacrifice the mice can also modify the read out. In our group, we sacrifice the animals by cervical dislocation to collect the samples as quickly as possible and avoid possible changes triggered by CO2 inhalation (the other type of euthanasia approved in our facility).

> AON efficacy assessment can be done at RNA or protein level. For example, checking by RT-PCR if the splicing is redirected, or by Western blot whether protein levels were increased or decreased, depending on the purpose of the study. For that, the retina needs to

Fig. 3 Schematic representation on how to place the right eye in the cryomold. The figure shows the different areas of the eye in the position of the mouse. When placing the eye in the mold, we put the dorsal (D) and the ventral (V) areas as indicated. Nasal (N) area will be at the bottom of the mold and the temporal (T) at the top. Arrows indicate the side of the mold used as reference to place the ora serrata of the eye in parallel. This is the common procedure we use in our lab, but the position of the eye can be done in different ways. Also the eye can be marked with a dye or by carefully burning the cornea at a specific position to recognize the orientation. The figure was made with Biorender

be harvested, and this can be done easily following the instructions previously described elsewhere [10]. Once the tissue is collected, it can be processed according to standard protocols for RNA isolation and protein lysates [6].

Morphological studies can also be performed. These tests are important for example, to study if the newly created protein localizes at the proper place or if the AON itself is creating any toxic effect. For that, the entire eye will be collected and subsequently sectioned to analyze these aspects by conventional eosin/hematoxylin or toluidine-blue staining or immunohistochemical studies. In order to prepare the samples, the eye needs to be sectioned. Here, there are two options. (A) The eye is fixed and dissected, removing the cornea and lens, cryopreserved in sucrose, and embedded in OCT, as previously described elsewhere [10], or (B) embedded directly in OCT. In general terms, fixed eyes will give better morphology than non-fixed. However, in our experience, often primary antibodies work only in unfixed tissue (especially those that are home-made). Therefore, choosing the best protocol to follow will depend on the reagents that will be employed for the different readouts. Since option A has already been described in detail previously [10], option B is explained below:


#### 4 Notes


the PBS 1, and the other for the AON, is recommended. This will ensure no traces of AON in the control eye and will contribute to reduce the overall intervention time.


#### Acknowledgments

The author wants to acknowledge Prof. Rob W.J. Collin and Dr. Irene Va´zquez Domı´nguez for critical review of this chapter. The group is financially supported by the Foundation Fighting Blindness (PPA-0517-0717-RAD), the Curing Retinal Blindness Foundation, as well as the Landelijke Stichting voor Blinden en Slechtzienden and Stichting Oogfonds via Uitzicht 2019-17, together with Stichting Blindenhulp, Rotterdamse Stichting Blindenbelangen, and Dowilwo. The funding organizations had no role in the design or conduct of this research and provided unrestricted grants. All figures were made with BioRender.

#### References


Unexpected CEP290 mRNA splicing in a humanized knock-in mouse model for Leber congenital amaurosis. PLoS One 8(11): e79369. https://doi.org/10.1371/journal. pone.0079369


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Delivery of Antisense Oligonucleotides to the Mouse Brain by Intracerebroventricular Injections

### Tom Metz, Elsa C. Kuijper, and Willeke M. C. van Roon-Mom

#### Abstract

The use of antisense oligonucleotides (AONs) is a promising therapeutic strategy for central nervous system disorders. However, the delivery of AONs to the central nervous system is challenging because their size does not allow them to diffuse over the blood–brain barrier (BBB) when injected systemically. The BBB can be bypassed by administering directly into the brain. Here we describe a method to perform single and repeated intracerebroventricular injections into the lateral ventricle of the mouse brain.

Key words Intracerebroventricular injections, Stereotactic surgery, Cannula, Antisense oligonucleotides

#### 1 Introduction

There has been a recent revival of interest in the use of antisense oligonucleotides (AONs) to treat neurodegenerative disorders with one approved central nervous system AON therapy and several in clinical trials [1]. This is largely due to the remarkably widespread distribution and cellular uptake of AONs once delivered into the brain. However, for drugs to reach the nervous system, they first have to cross the blood–brain barrier (BBB). Since the molecular weight of AONs is approximately 6000–10,000 Da, they are too large to cross the BBB by simple diffusion when delivered systemically. During the study of therapeutic efficacy of AONs in mouse models, AONs are often infused intracerebroventricularly (ICV). The BBB is bypassed by injecting the AON directly into the lateral ventricle, after which the AONs pass the ependymal cell layer that lines the ventricular system and enters the brain parenchyma.

One group of disorders for which these AON therapies are being studied in mouse models are the polyglutamine disorders. In these disorders, the disease is caused by CAG triplet repeat expansions in the coding region of a gene that is subsequently translated into an expanded stretch of glutamine amino acids in

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_23, © The Editor(s) (if applicable) and The Author(s) 2022

the protein. The disease-causing proteins for each of these polyglutamine (polyQ) disorders are different, but in each case, the expanded stretch of glutamines results in a toxic gain of function of the protein leading to neurodegeneration. To date, a total of nine polyQ disorders have been described: dentatorubralpallidoluysian atrophy (DRPLA), Huntington's disease (HD), spinal bulbar muscular atrophy (SBMA), and spinocerebellar ataxias (SCA1, 2, 3, 6, 7, and 17) [2, 3]. There is an inverse correlation of disease onset and polyQ length in the protein; the longer the CAG repeat, the earlier the age of onset of the disease [2]. Protein aggregates are found in the nucleus and cytoplasm of cells, indicating that protein misfolding is a common feature of these disorders. Using RNA breakdown [4] or RNA splice modulating AONs [5, 6], it is possible to reduce the toxicity of the mutant proteins, which can hopefully halt the disease progression in patients.

Because of the limited volume that can be injected into the mouse brain, multiple injections can be required to reach the desired dose. Repeated injections of AONs lead to a widespread distribution throughout the entire mouse brain, and the protein modifying effect can be detected in the cortex, cerebellum, and brainstem [6]. In this chapter, we describe a method for ICV delivery of AONs through a single injection or repeated injections using a cannula (Fig. 1).

#### 2 Materials


	- 2. Tubing (PlasticsOne C313CT).
	- 3. Cannula guide (PlasticsOne C315GS-5-SP).
	- 4. Dummy cannula (PlasticsOne C315DCS-5-SPC).
	- 5. MRI-compatible cannula guide (PlasticsOne C315GS-5-Pk/ SPC) (see Note 2).
	- 6. MRI-compatible dummy cannula (PlasticsOne C315DCNS-5/SPC) (see Note 2).
	- 7. Dental cement (DiaFil Flow 1928–5005-01 DiaDent).
	- 8. Primer (OptiBond® All-In-One 33381-E; Kerr Dental, Bioggio, Switzerland).
	- 9. Analgesia (Carprofen 50 mg/mL).
	- 10. Ocular lubricant (Added Pharma 220201).
	- 11. Petroleum jelly.
	- 12. Small piece of paper.

Fig. 1 Location and tools for intracerebroventricular injection. (a) Schematic and photographic outline of the injection site (I), bregma (B), and lambda (L) as points of reference. (b) Cannula types required for intracerebroventricular injection. The infusion cannula is connected to the syringe tubing and guided through the cannula guide to inject into the lateral ventricle. The cannula guide is attached to the skull and can be closed using a dummy cannula

	- (a) Noninvasive ear bars.
	- (b) Stereotactic arm that fits the cannula guide.
	- 2. Drill (Meisinger 310104001001005).
	- 3. Macroscope + light.
	- 4. UV light source.

#### 2.4 Solutions 1. Reconstituted AON in DPBS (Gibco 14190144).

#### 3 Methods

The limit of a bolus injection in the adult mouse brain is about 10 μL. In order to reach a sufficient dose of AON in the mouse brain, multiple injections over a longer period of time can be used (see Note 3). To make consecutive injections more easy and more animal-friendly, a cannula guide is placed on the mouse skull (see Note 4). The following injections can be given without surgery (see Note 5). The animal only has to be anesthetized and injected, which will not take longer than 10 min.

#### 3.1 Preparation 1. Prepare the stereotactic setup.


#### 3.2 Surgical Procedure


#### 3.3 Consecutive Injection


#### 4 Notes


Fig. 2 MRI of mouse brains holding an ICV cannula. (a) Axial image of mouse brain depicting the lateral ventricles (La) showing the protrusion of the cannula (C, white dashed line) into the right lateral ventricle. (b) Axial image of mouse brain showing skull malformation by intrusion of cement (Ce, white dashed line) oppressing the brain

before applying. Make sure the skull is very dry before applying primer, if necessary use a cloth.


#### Acknowledgments

The research of the group is supported by: Campagne Team Huntington. "Antisense oligonucleotide disease modifying treatment for Huntington's disease"; AFM Telethon. "Final proof of concept for the advancement of antisense oligonucleotide treatment for SCA3 towards the clinic" (Project number 20577); and ZonMW Memorabel. "RNA modulating therapy for Alzheimer's disease" (Project number 733050818).

#### References


DA, Baumann T, Gerlach I, Schobel SA, Paz E, Smith AV, Bennett CF, Lane RM (2019) Targeting huntingtin expression in patients with Huntington's disease. N Engl J Med 380(24): 2307–2316. https://doi.org/10.1056/ NEJMoa1900907


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part V

Safety and Toxicology

# Intrathecal Delivery of Therapeutic Oligonucleotides for Potent Modulation of Gene Expression in the Central Nervous System

### Zachary Kennedy, James W. Gilbert, and Bruno M. D. C. Godinho

#### Abstract

Therapeutic oligonucleotides hold tremendous potential for treating central nervous system (CNS) disorders. The route of administration of oligonucleotides significantly impacts both distribution and silencing efficiency. Here, we describe a technically simple, clinically relevant method to administer oligonucleotide compounds into the CNS via direct intrathecal injections. This method achieves distribution throughout the CNS rapidly and permits high-throughput testing of oligonucleotide efficacy and potency in mammals.

Keywords siRNA, Antisense oligonucleotides, CSF infusion, IT dosing, Intrathecal injections, Modified oligonucleotides, Gene silencing, Therapeutic oligonucleotides, CNS administration

#### 1 Introduction

Therapeutic oligonucleotides hold tremendous potential for treating disorders of the central nervous system (CNS), but numerous factors can hinder the delivery of these compounds. Among the major determinants affecting delivery of drugs to the CNS is the route of administration.

Drug delivery to the CNS can be performed through two general approaches: (a) systemic, where oligonucleotides are initially delivered outside of the CNS, or (b) direct, where oligonucleotides are delivered directly within the CNS. Systemic delivery strategies (e.g., intravenous or subcutaneous) are technically simple and easy to perform, but require significantly higher doses and result in very low efficacy in the CNS, mostly due to the inability of oligonucleotides to pass the blood–brain barrier [1–3]. Direct delivery methods, on the other hand, are comparatively more technical, but require much lower doses and enable significantly better

Zachary Kennedy and Bruno M. D. C. Godinho have contributed equally to this work.

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_24, © The Editor(s) (if applicable) and The Author(s) 2022

distribution, efficacy, and potency throughout the CNS than systemic methods [4, 5]. The most commonly used direct delivery methods, especially in rodent models, are intracerebroventricular (ICV) injections or lumbar intrathecal (IT) injections.

From a technical perspective, IT injections are easier to perform than ICV injections—they require less setup/procedural time and do not require stereotactic equipment [6, 7]. IT administration is also favored from a clinical perspective because it is less invasive than ICV injections; IT injections do not require brain surgery and can be performed as an outpatient procedure. IT injections can be performed with or without a catheter. Currently, IT injections are the only clinically approved route of administration for therapeutic oligonucleotide treatment of CNS disorders [8].

Given its ability to deliver to the CNS, the comparatively quick operation time, and the clinical relevance of the route of administration, IT injections are an attractive delivery method for which to screen oligonucleotide compounds for CNS applications. In this chapter, we describe how to perform direct IT injections in mice without using a catheter. Special attention is given to the preparation of the oligonucleotide compounds, prepping the mouse for the injections, and ensuring successful delivery.

#### 2 Materials



#### 3 Methods

Oligonucleotides must be prepared aseptically in a laminar flow cabinet to avoid contamination and maintain sterility. All procedures performed in live animals must be approved by the Institutional Animal Care and Use Committee (IACUC) prior to executing the experiment.

#### 3.1 Preparation of Test Oligonucleotides It is recommended that test oligonucleotides be treated with a calcium solution to minimize acute in vivo toxicities (see Note 1). ASOs can be treated immediately after synthesis, whereas siRNAs should be treated after duplexing of the guide and passenger strands (see Note 2).


While the aforementioned CaCl2 wash steps minimize toxicity, recovery of oligonucleotides from the columns is often incomplete, resulting in reduced yields. If the initial amount of oligonucleotides is low, an alternative strategy would be to evaporate oligonucleotides in SpeedVac (using the no temperature option for drying rate) and resuspend at the desired concentration with the chosen buffer.

#### 3.2 Direct Intrathecal Injection


Fig. 1 Intrathecal injection procedures. (a) Setup and materials needed for intrathecal injections. (b) Placement of anesthetized mice. After shaving the mice, the mouse is placed on a 15-mL conical tube placed under the lower stomach, allowing for easy visualization of the hip and spinal cord protrusions. Asterisk indicates injection site. Dotted lines indicate the superficial depression formed by spinal cord and iliac crests. (c) Placement of hands during procedure. Mouse is grasped by pinching the hips and lifting slightly, allowing for


When establishing this methodology in the lab, it is recommended to first practice using Evans blue dye. A successful injection will result in immediate distribution of the dye throughout the spinal cord and periphery of the brain (see Fig. 1e). It is also important to consider that distribution of therapeutic oligonucleotides is highly dependent on its conjugated ligand modality. Thus, in studies investigating biodistribution, oligonucleotides are often labeled with fluorescent tags, such as Cy3. Figure 2 shows the distribution of a Cy3-labeled Di-siRNA scaffold (20 nmol in 10 μL) 48 h after a single IT injection.

3.3 Suggestions for Tissue Collection and Processing For microscopy, the whole spine may be fixed in decalcifier/formalin after perfusion of the animal with cold 1 PBS. For RNA and protein analyses, the spinal cord can be either dissected or flushed out of the spine. After identifying the different segments of the spinal cord, these can be divided longitudinally to generate two samples per region for: (1) RNA assessment and (2) protein evaluation. RNA analyses can be carried out by RT-qPCR, bDNA, or other methods. Protein analyses can be performed by Western blot, ELISA, or other methods.

Fig. 1 (continued) maximum exposure of the cartilage between the L4 and L5 spinal segments. Successful needle insertion is often accompanied with a subtle tail-flick, which can be observed at the base of the tail or the very tip (see arrows), (d) Placement of needle during procedure. Dotted lines indicate the superficial depression formed by the spinal cord and iliac crest, where the hip meets the spinal cord. (e) Ventral view of brain and spinal cord immediately after IT injection of Evans blue dye

Fig. 2 Distribution pattern of Cy3-labeled oligonucleotide within the spinal cord. Horizontal cross sections of spinal cord segments following intrathecal injection of Cy3 labeled Di-siRNA oligonucleotides (20 nmol). Mice were euthanized 48 h post injection and tissues processed for microscopy. Tiled fluorescent images were acquired using 10 objective and displayed as overlay images: Cyan (DAPI, Nuclei), red (Cy3-labeled oligo). Scale bar 1 mm

#### 4 Notes


#### Acknowledgments

The authors would like to thank Dr. Anastasia Khvorova of the RNA Therapeutics Institute (University of Massachusetts Medical School) for continuous mentorship and research support. The authors would also like to thank Jake Metterville of Wave Life Sciences, Greg Cottle, and Van Gould of the Department of Animal Medicine (University of Massachusetts Medical School) for expert advice on technical details and animal welfare compliance. The authors acknowledge Dr. Emily Haberlin for editorial feedback on the manuscript.

#### References


1, randomised, first-in-man study. Lancet Neurol 12(5):435–442. https://doi.org/10. 1016/S1474-4422(13)70061-9


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 25

# Preclinical Safety Assessment of Therapeutic Oligonucleotides

### Patrik Andersson

#### Abstract

During the last decade, therapeutic oligonucleotide drugs (OND) have witnessed a tremendous development in chemistry and mechanistic understanding that have translated into successful clinical applications. Depending on the specific OND mechanism, chemistry, and design, the DMPK and toxicity properties can vary significantly between different OND classes and delivery approaches, the latter including lipid formulations or conjugation approaches to enhance productive OND uptake. At the same time, with the only difference between compounds being the nucleobase sequence, ONDs with same mechanism of action, chemistry, and design show relatively consistent behavior, allowing certain extrapolations between compounds within an OND class. This chapter provides a summary of the most common toxicities, the improved mechanistic understanding and the safety assessment activities performed for therapeutic oligonucleotides during the drug discovery and development process. Several of the considerations described for therapeutic applications should also be of value for the scientists mainly using oligonucleotides as research tools to explore various biological processes.

Keywords Oligonucleotide drugs, ASO, siRNA, Antisense, Non-clinical safety assessment, Preclinical safety assessment, Toxicity

#### 1 Introduction: Oligo Classes, Chemistries, and Designs

Oligonucleotide drugs (OND) of different classes range from 10–12 up to 100 nucleotides in length, often with chemical modifications of the backbone and ribose sugar. The chemistry and design used are dictated by the desired mechanism of action, resulting in different classes of therapeutic oligos with specific properties. The most common classes in clinical studies target RNA and rely on Watson-Crick hybridization for selectivity and affinity, where antisense oligonucleotides (ASOs) and short interfering RNAs (siR-NAs) are the most common, with several examples of approved products [1, 2].

ASOs are single-stranded ONDs of 12–20 nucleotides in length, often with a phosphorothioate (PS) backbone and 20 ribose

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_25, © The Editor(s) (if applicable) and The Author(s) 2022

modifications like OMe, MOE, LNA, and cEt that improve drug properties like metabolic stability, tissue uptake, and increased affinity to the target transcript [3–5]. The ASO class can be further subdivided into ASO gapmers that trigger RNase H-mediated cleavage and degradation of target transcripts and steric blocking ASOs that modulate splicing events or inhibit activity of, e.g., microRNA (miR) [6–8]. For ASOs with a steric blocking activity, use of other chemistries resulting in neutral backbones like Phosphorodiamidate Morpholinos (PMO), Peptide nucleic acids (PNA) and tricyclo-DNA (tcDNA) is possible and quite common [9–13].

siRNAs and microRNA-mimics have a double-stranded design with each strand approximately 20–24 nucleotides in length and rely on loading of the antisense strand into RISC for activity [14, 15]. The design of siRNA results in cleavage and subsequent degradation of the target transcript, whereas miR mimics regulate gene expression by binding to miR sites in mRNA, inducing degradation and regulating protein translation [8].

In addition to hybridization dependent ASOs, miR mimics, and siRNA, there are several classes of hybridization independent ONDs including aptamers [16, 17] and immunostimulatory CpG oligos [18–21]. Rather than binding to RNA, the threedimensional structure of folded RNA of a given sequence is combined with chemical modifications to achieve specific binding to proteins. Other therapeutic approaches such as mRNA therapy and the guide RNA in various gene editing approaches (e.g., CRISPR/ CAS9) utilize nucleotides and but are commonly not classified as ONDs.

This chapter will focus on safety assessment of the hybridization dependent PS backbone ASOs and siRNA.

#### 2 Delivery

The activity of ASOs, siRNA, and miR mimics rely on reaching the interior of the target cells. Uptake per se is not enough; the oligo needs to access the right subcellular compartments for activity, e.g., the cytosol for RISC loading or the nucleus for splice modulation or RNase H activity. Cellular uptake leading to pharmacodynamic effects are often referred to as "productive uptake" [22–24]. For ASOs, the best productive uptake after systemic delivery is often observed in the liver, in particular hepatocytes [24, 25]. Early siRNA candidates were delivered in protective formulations that also showed best uptake into hepatocytes [26]. Local delivery has successfully been used to bypass the blood–brain barrier, resulting in good therapeutic effects after local delivery to CNS [27–29], and the eye [30–33]. However, for use of therapeutic oligos beyond hepatocytes [3, 5, 34–37] and local delivery, achieving sufficient productive uptake has become one of the biggest challenges and numerous ways to overcome this has been proposed [36, 38, 39]. One approach is delivery in different types of formulations like the lipid nanoparticle (LNP) used to deliver Onpattro [26], the first siRNA receiving regulatory approval by FDA in 2018 [40]. However, although efficient delivery at low doses can be achieved, the LNP triggers proinflammatory flu-like responses that are managed by pre-medication before administration [41]. Since discovery of this LNP, a number of alternative formulations have been presented with different tissue distribution and improved efficacy:safety relationship.

An alternative way to improve the productive uptake is conjugation of the oligo to a targeting ligand, utilizing binding to cell surface receptors that internalize the oligo conjugate. An excellent example of this strategy is conjugation of the GalNAc carbohydrate, resulting in significantly improvements in productive uptake of both siRNA and ASOs [42, 43]. This GalNAc-mediated improvement in productive uptake is mediated by the binding of the conjugate to the asialoglycoprotein receptor (ASGR), which mainly is expressed on hepatocytes. Although non-conjugated ASOs show hepatocyte activity, adding GalNAc conjugates increased the clinical potency 20–30-fold for several re-formatted ASOs with hepatocyte targets [44]. Combining GalNAc conjugation with nuclease-resisting chemical stabilization has led to a tremendous increase in the utility of siRNA in the clinic, with several recent approvals with no need for the pre-medication required for LNP formulated Onpattro [2].

#### 3 Safety Assessment of Therapeutic Oligos

The focus of this chapter is the preclinical safety assessment of therapeutic oligos intended to enter clinical trials to get regulatory approval for use in patients. With increasing experience and mechanistic understanding, screening cascades, study designs and data interpretations for ONDs are constantly improving, leading to more potent clinical candidates with better safety profiles. As described below, safety assessment of clinical OND candidates is a highly regulated process that at first sight could be of less interest for scientists primarily using oligonucleotides as tools to dissect and understand basic biological process. However, several of the considerations for bringing safe candidates to clinical trials could also be of value when developing optimal tool oligos and study designs for basic research. This includes cross species activity, restricted uptake distribution, long tissue half-life and effect duration, hybridization dependent off-target effects and the need to screen away from sequence dependent toxicities.

3.1 Discovery Phase: Selecting the Oligo Candidate with the Best Balance Between Potency and Safety

The potential safety concerns with therapeutic oligos can be divided into:

	- 3. Sequence dependent, but hybridization independent (Sect. 3.1.3).

3.1.1 Sequence and Hybridization Dependent Effects: Assessing On- and Off-Target Safety

On-target toxicities, also known as exaggerated pharmacology, are dependent on sequence and RNA hybridization for ASOs, siRNA, and miR mimics. It can manifest in too strong intended effect or an adverse consequence of the pharmacological response in an unintended organ. A first assessment of potential on-target safety risks should be a theoretical exercise compiling available information on the biological role, tissue expression pattern, competitor information, etc. to assess the likelihood and potential adverse impact of the identified risks in the intended patient population. Considerations for assessing on-target toxicities for ONDs has been discussed by Kornbrust et al. [45].

Another safety concern dependent on Watson-Crick base pairing is hybridization dependent off-target risks. In contrast to the risk for on-target toxicity described above where the oligo has the intended, but exaggerated activity, this off-target risk relates to oligo activity on other transcripts than the intended target. Key features determining likelihood for hybridization dependent offtarget effects have been discussed in depth elsewhere [46–50] and several recommendations [51] are summarized below:


There are some specific considerations when assessing potential hybridization-dependent safety concerns. First, due to species differences in sequence, it is often difficult to achieve pharmacological activity with the same OND in other species and it is common practice to use surrogate molecules with sufficient potency in the model species of choice.

Second, the chemical modifications used in therapeutic oligos lead to slow tissue elimination and extended effect duration [52]. This is convenient from a delivery perspective, but the washout period needed for the adverse effects to cease would be equally long should on- or off-target toxicities be observed.

Third, the restricted productive uptake of ASOs and siRNA to many cell types needs to be considered, as there is a large difference in the uptake between different cells and tissues. The oligo distribution can change with other administration routes and delivery systems like conjugates or formulations, so understanding the productive uptake distribution for such new conditions is critical for proper risk assessment of potential on- and off- target toxicities.

A couple of toxicities that are dependent on plasma Cmax but independent of both hybridization and sequence can be observed at relative high doses of PS backbone ASOs. This includes prolongation of coagulation time and activation of the alternative complement system. Acutely, activation of the alternative complement system can lead to significant drops in blood pressure. Repeated complement activation can result in "consumption" of complement factor C3 with impaired complement-mediated clearance of antibody aggregates resulting in vascular inflammation [53]. Data from in vitro, in vivo, and clinical studies clearly show that cyno-

this lowered threshold of complement activation [54, 55]. Both these effects are driven by the plasma Cmax levels [56–60] and transient in nature. With increased potency of modern ASOs and adapted dosing in current clinical studies, plasma concentrations rarely exceed these activation thresholds [55, 61, 62].

molgus monkeys are significantly more sensitive than humans for

In contrast to these plasma Cmax-driven effects, other hybridization independent toxicities are highly dependent on the OND sequence. This includes proinflammatory manifestations and effects in high exposure organs such as liver and kidney that can sometimes be observed during the discovery phase. For siRNA, liver toxicity has been explained to be caused by off-target effects in the seed-region of the siRNA [48]. For PS backbone ASOs, liver toxicity is more frequently observed with higher affinity chemistry like LNA and cEt. It is clear from a number of published and unpublished observations that this liver toxicity observed during the screening process of PS backbone ASO gapmers is not caused by knockdown of the intended target transcript or liver concentration per se, see e.g. [63]. ASO sequence motifs associated with liver tox [64, 65] and different molecular mechanisms have been proposed, including cell death as a cellular consequence to exaggerated RNase H activity resulting from non-selective hybridization [66, 67]. An alternative mechanism proposed involves PS backbone-dependent binding to key intracellular proteins in a sequence and chemistrydependent manner. Being more hydrophobic, the higher affinity modifications showing higher incidence of liver toxicity also show higher affinity to a number of intracellular proteins compared to the same sequence with, e.g., MOE chemistry [68–71]. Importantly,

3.1.2 Sequence and Hybridization Independent Effects: Coagulation Time and Complement Activation

3.1.3 Sequence Dependent, But Hybridization Independent: Inflammation, Liver, and Kidney Toxicities

predictive in vitro models for liver and kidney toxicity have been developed [67, 72, 73], and design modifications reducing ASO toxicity without compromising potency have been proposed [69, 74] demonstrating that highly potent ASO sequences that do not show liver toxicity can be identified and progressed to clinical trials.

Immune-stimulatory effects have long been a prominent feature of ONDs, where responses may vary widely between species and depend on oligonucleotide design and sequence, as well as chemical modifications [75–85]. The immunomodulatory potential can deliberately be used to design nucleotide-based immunotherapies and vaccine adjuvants, often harboring so-called CpG motifs [18–21], but for most other OND these effects are unwanted. Despite avoiding established CpG motifs in the design phase, some therapeutic oligos induce clear proinflammatory effects that can show in several different ways in the clinic, including injection site or infusion related reactions, flu-like symptoms and thrombocytopenia [86–90]. These effects are dose-dependent and can occur at different time points after first administration of the drug. Rodents are particularly sensitive to the immunostimulatory effects of ONDs [91, 92]. Similar to the liver toxicity described above, oligo sequence is a key parameter defining the proinflammatory property of therapeutic oligos and subtle and systematic sequence modifications to a proinflammatory ASO resulted in clear differences in proinflammatory potential [93]. Chemical modifications can modify the immune stimulatory potential of ONDs of a given sequence: PS modification of the backbone has long been known to increase the immune stimulatory properties of ONDs [80, 85, 94], whereas the neutral backbone in PMOs does not evoke an immune response [95]. 5<sup>0</sup> -methylation of cytosine is frequently used to suppress the immune stimulatory effect of CpG DNA sequences [76, 79]. 2<sup>0</sup> OMe modification of ssRNA or siRNA sequences inhibit immune stimulation, whereby even single modifications can significantly reduce the cytokine upregulation [78, 84]. Other 20 ribose modifications (20 F, 20 H, 20 MOE, LNA) have also been described to reduce proinflammatory effects [76, 83]. Therapeutic oligos administered in lipid formulations have been shown to induce inflammatory responses, and humans seem to be more sensitive to these effects than both rodents and NHPs [96–98].

Thrombocytopenia (TCP), i.e., low concentration of circulating platelets, has been observed in NHP toxicity studies with ASOs. In most cases, the platelet counts show around 30% reduction from baseline and then stabilize at a non-adverse level. However, in some drug programs, a few individual monkeys have experienced severe TCP [99]. Severe TCP was observed in the phase 3 studies for volanesorsen and inotersen as well as for drisapersen [86, 100, 101]. These TCP events occurred in the highest dose group, and platelet counts increased after drug cessation. A combination of high dose of proinflammatory ASOs and possibly patient susceptibility factors seem to be the most likely cause: a high frequency of severe TCP in cynomolgus monkeys of Mauritian origin whereas no cases of severe TCP were observed when the same ASO was given to non-Mauritian cynomolgus monkeys in a follow-up study [102].

Strategies for preclinical safety assessment of therapeutic oligos have been discussed elsewhere [22, 89]. Results from a survey across 23 companies developing therapeutic oligos performed in 2018 showed that most companies follow the two species small molecule approach as outlined by the ICH M3(R2) guideline [103]. Although a guideline recently was adopted by Japanese regulators, most health authorities lack formal regulatory guidelines for therapeutic oligos, so the expectations and experience may vary between regions and even within health authorities [103]. However, white papers published by cross-pharma groups like OSWG (Oligo Safety Working Group) on best practice recommendations are frequently used as informal guidelines [45, 51, 96, 104–110].

For the common situation with a human active candidate having limited cross-species activity for meaningful assessment of potential on-target toxicity, a surrogate molecule can be included in parallel to the clinical candidate. Such surrogate molecules should be of the same design and chemistry as the clinical candidate and have a good general safety profile to allow meaningful assessment and documentation of potential on-target toxicities. Such surrogate molecules are mostly designed to be active in the rodent species of choice.

Despite lack of positive results in regulatory genotoxicity studies as discussed in the OSWG white paper by Berman et al. [105], several health authorities still request in vitro and in vivo assessment of genotoxicity [103].

For small molecules, in vitro and in vivo safety pharmacology studies are important to rule out adverse functional effects on key organs such as the CNS, cardiovascular and respiratory systems. Safety pharmacology assessment has been discussed in another OSWG white paper [106]. To the knowledge of the author, there is no information on systemically administered ASOs or siRNA showing activity in vitro or in vivo safety pharmacology studies, including activity on the hERG channel or any other ion channels important for cardiovascular or neuronal function. However, direct delivery into heart and CNS is a different story and could result in functional effects.

Similar to small molecules, the general toxicity studies for ASOs and siRNA are performed in a rodent and a non-rodent species. The duration of these Good Laboratory Practice (GLP) studies range from 1 to 3 months in the beginning of a program to chronic

#### 3.2 Development Phase: In-Depth Characterization and Documentation of the Oligo Candidate

3.2.1 Preclinical Safety Assessment During the Development Phase

studies of 6 and 9 months in duration for rodents and non-rodents, respectively. For double-stranded siRNA and microRNA mimics, the rat is by far the most commonly used rodent species whereas the mouse is rodent species of choice for most single stranded PS backbone ASO candidates [103]. Although rat is the most common rodent for toxicity studies of small molecules, rat specific lesions like Chronic Progressive Nephropathy (CPN) [91, 92] are aggravated by the high kidney concentrations resulting from systemic delivery of PS backbone ASO. CPN is of no human relevance [111] but can become problematic in toxicity studies of long duration.

Due to the highest likelihood of sequence-dependent crossover on activity and a robust historical background record, the non-human primate (NHP), is by far the most common non-rodent species used for both ASO and siRNA candidates, but other non-rodents have been evaluated [103], including the pig [112]. Other in vivo studies required before regulatory approval include developmental and reproductive toxicology studies (DART) and carcinogenicity studies. Considerations for the DART studies are described in an OSWG white paper [108] and is normally run in mouse, rat or rabbit. Carcinogenicity studies are commonly run as a lifelong (2 years) studies in rat and mice or a 2-year study in rat combined with a 6-month study in a transgenic mouse model. The relevance of these carcinogenicity studies for ASOs and siRNA has been questioned, but most health authorities are likely waiting for more data before discussing whether these studies can get a waiver or not.

3.2.2 Regulatory Perspective Several of the oligonucleotide products approved to date (e.g., eteplirsen, mipomersen, inotersen, volanesorsen) are aimed at treating rare, often genetic diseases for which no alternative treatment is available. In such cases, the presence of some safety signals has been judged to be acceptable. However, with some recent projects aiming at targeting significantly larger populations with more common disease, like the cholesterol-lowering siRNA inclisiran targeting PCSK9 and for which other treatments exist, the risk:benefit assessment will likely be different. At the same time as ONDs are considered for much larger and broader patient populations than before, exciting opportunities on the other end of the patient population spectrum are emerging. Milasen is a splice modulating ASO developed to treat a fatal neurodegenerative condition unique to a single patient [113]. With increasingly refined and optimized screening cascades, OND treatments for <sup>N</sup> <sup>¼</sup> 1 and other ultrarare conditions will most likely become more common practice.

> In summary, safety of ONDs depend on sequence, chemistry, design, and delivery approach. Several properties like limited species cross-reactivity, long tissue half-life and restricted productive

uptake distribution needs to be considered when designing interpreting results for hybridization dependent on – and off-target safety assessment studies. Together with liver and kidney toxicity, proinflammatory effects are the most commonly observed safety findings in preclinical studies. With improved mechanistic understanding and screening approaches, more potent OND candidates with better safety profile can be identified for treatment of an increasing range of diseases and patient populations.

#### References


381–393. h t tp s://doi.o rg/10.3233/ JND-160157


Nat Rev Drug Discov 16(3):181–202. https://doi.org/10.1038/nrd.2016.199


Nucl Acid Ther 28(3):146–157. https://doi. org/10.1089/nat.2018.0721


Soc 136(49):16958–16961. https://doi. org/10.1021/ja505986a


(2014) Sequence motifs associated with hepatotoxicity of locked nucleic acid—modified antisense oligonucleotides. Nucleic Acids Res 42(8):4882–4891. https://doi.org/10. 1093/nar/gku142


Assessment of formulated oligonucleotidebased therapeutics. Nucl Acid Ther 27(4): 183–196. https://doi.org/10.1089/nat. 2017.0671


Associations Oligonucleotide Working Group Survey on nonclinical practices and regulatory expectations for therapeutic oligonucleotide safety assessment. Nucl Acid Ther 31(1):7–20. https://doi.org/10.1089/nat. 2020.0892


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 26

# Preclinical Evaluation of the Renal Toxicity of Oligonucleotide Therapeutics in Mice

### Lucı´a Echevarrı´a and Aurelie Goyenvalle

#### Abstract

Antisense oligonucleotides (ASO) therapeutics hold great promise for the treatment of numerous diseases, and several ASO drugs have now reached market approval, confirming the potential of this approach. However, some candidates have also failed, due to limited biodistribution/uptake and poor safety profile. In pursuit of better delivery and higher cellular uptake, ASO are being optimized, and new chemistries are developed or conjugated with various ligands. While these developments may lead to candidates with higher potency, it is important to keep the safety aspects in sight and screen for potential toxicity in early phases of preclinical development to avoid subsequent failure in clinical development. Our understanding of ASO-mediated toxicity keeps improving with increased preclinical and clinical data available. In this chapter, we will focus on the assessment of renal toxicity in mice and describe methods to measure the levels of general urinary biomarkers as well as acute kidney injury biomarkers following ASO treatment.

Key words Antisense oligonucleotides (ASO), Safety, Toxicology, Kidney toxicity, Acute kidney injury biomarker, Preclinical evaluation, Mouse model

#### 1 Introduction

The field of synthetic antisense oligonucleotide (ASO) has advanced remarkably in the last decade, and ASOs represent a very promising therapeutic platform that keeps evolving rapidly, in particular in pursuit of delivery improvement. Many preclinical studies in the antisense area focus on improving ASO delivery and assessing their efficacy in target tissues, often neglecting the evaluation of toxicity, at least in early phases of development. However, safety assessment is particularly important when developing new generations of ASOs or novel delivery systems to avoid the potential failure of a new drug in further toxicological studies, as it happened with a peptide conjugated PMO (PPMO) targeting the human dystrophin exon 50 which was found to cause mild tubular degeneration in the kidneys of cynomolgus monkeys [1].

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_26, © The Editor(s) (if applicable) and The Author(s) 2022

Toxicological properties of ASO have been comprehensively and extensively summarized previously [2, 3], and our understanding of them has allowed the development of predictive tests to select the best preclinical candidates.

Following systemic administration, the highest concentrations of ASO, independently of their chemistry, are found in liver and kidney, which are therefore considered as high-exposure organs. Importantly, tissue concentration does not further increase upon re-administration once steady-state is reached [2]. Accumulated ASOs can often be visualized at the histopathology level as basophilic granules on tissue sections stained with hematoxylin and eosin, but these effects are regarded as nonadverse because of their reversible nature upon treatment cessation. In rodents treated with high dose of PS-ASO, it is also frequent to observe tissue macrophages with a foamy appearance, referred to as histiocytes, which store cytokines in response to an activated state [3].

Considering the high concentrations of ASO accumulating in the kidneys, including the charge-neutral backbone such as PMO [4, 5], they are regarded as a common organ for toxicity. The highest uptake is generally observed in the proximal tubular epithelial cells of the convoluted tubule, whereas uptake in tubular cells in kidney medulla is much lower [6, 7]. Renal effects therefore tend to be more tubular than glomerular, apart from the reported glomerulopathies in mouse and monkey studies with the 2<sup>0</sup> OMe PS Drisapersen developed for the treatment of DMD [8]. However, it appears that this toxicity was linked to the chronic complement activation and inflammatory effects of the ASO and therefore overpredicted in animal studies since humans are less susceptible to these effects. Much more common are the lesions observed in the proximal tubules, which typically appear in animals treated with much higher doses of ASOs than the clinically relevant doses. Renal toxicity is mostly regarded as an accumulation-related toxicity and mostly sequence unspecific, except for more acute tubular lesions reported with high-affinity ASO such as locked nucleic acid (LNA) [9]. These effects might be related to excessive accumulation of RNase H-dependent off-target transcripts and/or specific protein binding [2] and a predictive EGF-based assay has recently been developed to exclude this type of kidney-toxic candidates [10].

Besides this EGF-based assay, several specific and early biomarkers of toxicity can be evaluated in mice (treated with high doses of ASO) to predict toxicity in preclinical development and exclude nephrotoxic candidates [8]. Evaluation of renal toxicology typically includes macroscopic examination of the kidneys upon necropsy of the animals followed by microscopic examination and careful histopathology analysis. General biomarkers of renal toxicity can be measured in the serum or plasma of treated mice such as urea, albumin, creatinine, and total protein. In this chapter, we focus on urinary biomarkers of kidney toxicity and describe the methods to measure the levels of total protein, albumin, creatinine as well as specific kidney injury biomarkers as a way of evaluating the potential renal toxicity of antisense oligonucleotides in mice. For this assessment, urines are collected from ASO-treated mice either shortly after ASO injection to evaluate the potential acute kidney toxicity or after several weeks of repeated treatment to evaluate the potential long term renal toxicity induced by the accumulation of ASO in kidneys.

#### 2 Materials



#### 3 Methods


3.1 Evaluation of Creatinine Levels in Urine

Urine creatinine is measured using a creatinine assay kit (in our case R&D Systems), based on the Jaffe reaction where creatinine is treated with an alkaline solution to yield a bright orange-red complex. Intensity of the color at 490 nm corresponds to the concentration of creatinine in samples.


Fig. 1 Schematic representation of urine collection possibilities following ASO treatment in mice and possible urine analysis

20 mg/dL to 0.31 mg/dL. Mix each tube thoroughly before the next transfer. Use deionized or distilled water as the zero standard (0 mg/dL).


#### 3.2 Evaluation of Total Protein Level in Urine


(<sup>n</sup> standards + <sup>n</sup> unknowns) (<sup>n</sup> replicates) (volume of WR per sample) ¼ Total V required. Prepare the WR by mixing 50 parts of BCA Reagent A with 1 part of BCA Reagent B (50: 1; A:B).


Albumin levels from urine samples are measured using an albumin ELISA kit. All reagents must be at room temperature before use. We describe the method below using the albumin ELISA kit from Bethy Laboratories:


#### 3.3 Evaluation of Albumin Levels in Urine


Acute kidney injury (AKI) biomarkers levels are analyzed by multiplex assays (MILLIPLEX® MAP) using the Luminex® technology.

The multiplex kidney injury panels (panel 1: MKI1MAG-94K and panel 2: MKI2MAG-94K, Merck-Millipore) are used according to the manufacturer's instructions to measure levels of β-2 microglobulin (B2M), renin, kidney injury molecule 1 (KIM-1), interferon-gamma induced protein 10 (IP-10), vascular endothelial growth factor (VEGF) (panel 1) and Cystatin C, epidermal growth factor (EGF), Lipocalin-2-NGAL, clusterin and osteopontin (OPN) (panel 2).

β2-Microglobulin, Renin, Kim-1, IP-10, VEGF are measured using the panel 1 (ref MKI1MAG-94K from Merck-Millipore).

1. Allow all reagents to warm to room temperature (20–25 C) before use in the assay (except antibodies and beads).

3.4 Evaluation of Acute Kidney Injury Biomarkers (AKI) Level in Urine

3.4.1 Evaluation of β2- Microglobulin, Renin, Kim-1, IP-10, and VEGF Levels in Mouse Urines

	- (a) Reconstitute quality control 1 and quality control 2 with 250 μL deionized water.
	- (b) Invert the vial several times to mix and vortex.
	- (c) Allow the vial to sit for 5–10 min.
	- (a) Reconstitute the mouse kidney injury panel 1 standard with 250 μL deionized water.
	- (b) Invert the vial several times to mix. Vortex the vial for 10 s.
	- (c) Allow the vial to sit for 5–10 min.
	- (d) This will be used as standard 7 (see Note 6).
	- (e) Label 6 Eppendorf microfuge tubes standard 1 through standard 6.
	- (f) Add 150 μL of assay buffer to each of the six tubes.
	- (g) Prepare serial dilutions (1:4) by adding 50 μL of the reconstituted standard to the standard 6 tube, mix well and transfer 50 μL of standard 6 to the standard 5 tube, mix well and transfer 50 μL of standard 5 to the standard 4 tube, mix well and transfer 50 μL of standard 4 to the standard 3 tube, mix well and transfer 50 μL of standard 3 to the standard 2 tube, mix well and transfer 50 μL of standard 2 to the standard 1 tube and mix well.
	- (h) The 0 pg/mL standard (background) will be assay buffer.
	- (a) Add 200 μL of assay buffer into each well of the plate. Seal and mix on a plate shaker for 10 min at room temperature (20–25 C).
	- (b) Decant assay buffer and remove the residual amount from all wells by inverting the plate and tapping it smartly onto absorbent towels several times.

method for calculating analyte concentrations in samples. For diluted samples, final sample concentrations should be multiplied by the dilution factor (25 as per protocol instructions). If using another dilution factor, multiply by the appropriate dilution factor.

Cystatin C, epidermal growth factor (EGF), Lipocalin-2-NGAL, Clusterin, and Osteopontin (OPN) levels are measured using the panel 2 (ref MKI2MAG-94K from Merck-Millipore):

	- (a) For individual vials of beads, vortex for 1 min.
	- (b) Add 150 μL from each antibody-bead vial to the mixing bottle and bring final volume to 3.0 mL with assay buffer.
	- (c) Vortex the mixed beads well (see Note 5).
	- (d) There are five biomarkers in panel 1: using five antibodyimmobilized beads, add 150 μL from each of the five bead vials to the Mixing Bottle. Then add 2.25 mL assay buffer.
	- (a) Reconstitute quality control 1 and quality control 2 with 250 μL deionized water. Invert the vial several times to mix and vortex.
	- (b) Allow the vial to sit for 5–10 min (see Note 6).
	- (a) First, reconstitute the mouse kidney injury panel 2 standard with 250 μL deionized water. Invert the vial several times to mix. Vortex the vial for 10 s. Allow the vial to sit for 5–10 min. This will be used as Standard 6 (see Note 6).
	- (b) Label 5 Eppendorf microfuge tubes standard 1 through standard 5.
	- (c) Add 150 μL of assay buffer to each of the five tubes.

3.4.2 Evaluation of Cystatin C, Epidermal Growth Factor (EGF), Lipocalin-2-NGAL, Clusterin, and Osteopontin (OPN) Level in Urine

	- (a) Add 200 μL of assay buffer into each well of the plate. Seal and mix on a plate shaker for 10 min at room temperature (20–25 C).
	- (b) Decant assay buffer and remove the residual amount from all wells by inverting the plate and tapping it smartly onto absorbent towels several times.
	- (c) Add 25 μL of each standard or control into the appropriate wells. Assay buffer should be used for 0 pg/mL standard (background).
	- (d) Add 25 μL of sample (diluted) into the appropriate wells.
	- (e) Add 25 μL of assay buffer to all wells.
	- (f) Add beads: Vortex mixing bottle and add 25 μL of the mix to each well. During addition of beads, shake bead bottle intermittently to avoid settling.
	- (g) Seal the plate with a plate sealer.
	- (h) Wrap the plate with foil and incubate with agitation on a plate shaker (~700 rpm) overnight (16–18 h) at 2–8 C.
	- (i) Place the plate on magnetic holder (handheld magnet, EMD Millipore Catalog #40–285) and rest the plate on magnet for 60 s to allow complete settling of magnetic beads.
	- (j) Remove well contents by gently decanting the plate in an appropriate waste receptacle and gently tapping on absorbent pads to remove residual liquid.
	- (k) Wash plate with 200 μL of wash buffer by removing plate from magnet, adding wash buffer, shaking for 30 s, reattaching to magnet, letting beads settle for 60 s, and removing well contents as previously described after each wash. Repeat wash steps three times.
	- (l) Allow the detection antibodies to warm to room temperature.
	- (m) Add 25 μL of detection antibodies into each well.

#### 4 Notes


#### Acknowledgments

LEZ is an employee of SQY Therapeutics and AG is funded by the Institut National de la sante´ et la recherche me´dicale (INSERM); the Association Monegasque contre les myopathies (AMM); and the Duchenne Parent project France (DPPF).

#### References


through in situ RNA hybridization in multiple organ systems following systemic antisense treatment in animals. Nucl Acid Ther 23: 369–378. https://doi.org/10.1089/nat. 2013.0443


#### 384 Lucı´a Echevarrı´a and Aurelie Goyenvalle

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Protocol for Isolation and Culture of Mouse Hepatocytes (HCs), Kupffer Cells (KCs), and Liver Sinusoidal Endothelial Cells (LSECs) in Analyses of Hepatic Drug Distribution

Kjetil Elvevold, Ingelin Kyrrestad, and Ba˚rd Smedsrød

#### Abstract

Development of the new generation of drugs (e.g., oligo- and polynucleotides administered intravascularly either as free compounds or as nano-formulations) frequently encounters major challenges such as lack of control of targeting and/or delivery. Uncontrolled or unwanted clearance by the liver is a well-known and particularly important hurdle in this respect. Hence, reliable techniques are needed to identify the type(s) of liver cells, receptors, and metabolic mechanisms that are responsible for unwanted clearance of these compounds.

We describe here a method for the isolation and culture of the major cell types from mouse liver: hepatocytes (HCs), Kupffer cells (KCs), and liver sinusoidal endothelial cells (LSECs). The presently described protocol employs perfusion of the liver with a collagenase-based enzyme preparation to effectively transform the intact liver to a single cell suspension. From this initial cell suspension HCs are isolated by specified centrifugation schemes, yielding highly pure HC preparations, and KCs and LSECs are isolated by employing magnetic-activated cell sorting (MACS). The MACS protocol makes use of magnetic microbeads conjugated with specific antibodies that bind unique surface antigens on either KCs or LSECs. In this way the two cell types are specifically and separately pulled out of the initial liver cell suspension by applying a magnetic field, resulting in high purity, yield, and viability of the two cell types, allowing functional studies of the cells.

If the drug compound in question is to be studied with respect to liver cell distribution of intravascularly administered drug compounds the isolated cells can be analyzed directly after isolation. Detailed studies of receptor-ligand interactions and/or dynamics of intracellular metabolism of the compound can be conducted in primary surface cultures of HCs, LSECs, and KCs established by seeding the isolated cells on specified growth substrates.

Key words Mouse liver cells, Cell isolation, Kupffer cells, Liver sinusoidal endothelial cells, Hepatocytes, Cell culture, Magnetic-activated cell sorting, MACS

#### 1 Introduction

Modern drug treatment modalities frequently include i.v. administration of large molecule compounds or nano-formulations. In either case oligonucleotides may represent the active principle.

Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6\_27, © The Editor(s) (if applicable) and The Author(s) 2022

Lack of targeting control is a general challenge when these therapeutics are injected i.v. The main cause of targeting failure is unwanted uptake in liver [1]. Apart from the one noteworthy example of i.v. administered oligonucleotide therapeutics patisiran that successfully targets the major type of liver cells, the hepatocytes (HCs) [2], the general rule is that large molecules/nanoformulations are rapidly cleared from the circulation by specialized scavenger cells lining the wall of the several hundred million capillary-like liver vessels called liver sinusoids. The scavenger cells of the liver sinusoids represent two cell types: the liver sinusoidal endothelial cells (LSECs) and the Kupffer cells (KCs). The KCs are the resident mononuclear phagocytes of the liver and represent by far the largest population of macrophages in the body. These cells are geared to carry out phagocytosis of large blood-borne material (>200 nm). In contrast the LSECs, representing the great majority of endothelial cells of the liver, are unable to perform phagocytosis under normal conditions, but are among the most active endocytic cells of the body, using clathrin-mediated endocytosis and a set of unique endocytosis receptors [3].

Although the LSECs and KCs make up only 3.3% and 2.5% of the total liver volume, the numberwise distribution of same cells is 21% and 8.5% [4]. The following anatomical and physiological relationships strongly support the notion that LSECs are optimally geared to carry out extremely efficient blood clearance: (a) The liver receives as much as 25% of the blood volume that is pumped through the heart at any time unit; (b) The KCs and LSECs represent the surface that the blood encounters on its journey through the liver; (c) The total surface of the LSECs in the human liver facing the sinusoidal lumen is that of a tennis court [3]; (d) The ad- and ab-luminal surfaces of LSECs carry unique endocytosis receptors that recognize and internalize an array of blood-borne waste substances, including several nanoparticles as well as oligo- and polynucleotides [5–8]; (e) The endocytosis receptors in LSECs mediating blood clearance recycle back to the cell surface only seconds after they have delivered their cargo to the primary endosomes, making these receptors extremely efficient. Moreover, considering the effective intracellular transport toward lysosomes and their unusually high contents of lysosomal enzymes, there should be no surprise that the LSECs and KCs represent very active scavenger cells contributing importantly to the high blood clearance capacity of the liver.

Based on the remarkable scavenging activity of the LSECs and KCs, projects including development of large molecule compounds and nano-formulated therapeutics should include studies to determine (a) if the drug candidates accumulate preferentially in LSECs and/or KCs, (b) and if so, what receptors are involved in the recognition. Exact studies designed to determine to what extent any i.v. administered compound accumulates in the different types of liver cells require reliable methods to isolate pure preparations of LSECs, KCs, and HCs from a single liver.

We pioneered the method of preparing LSECs, KCs, and HCs from a single rat liver, using collagenase perfusion to disperse the liver cells, followed by density separation on Percoll, and selective substrate adherence [9]. This method, which has been used with or without modifications by several laboratories, was later adapted by us for isolation and culture of mouse LSECs, KCs, and HCs [10].

While the procedure for isolation of HCs is rather straightforward and will be only briefly mentioned later in the present methods description, techniques for isolation of LSECs and KCs are more elaborate and will therefore be dealt with in greater detail in the present chapter. The great majority of the techniques used nowadays to generate initial single cell suspensions of liver include perfusion with a collagenase solution through the portal vein. The further steps comprise either centrifugal elutriation [11], Percoll density separation followed by selective adherence [10], or magnetic-activated cell sorting (MACS). Fluorescence-activated cell sorting (FACS) is frequently used to isolate KCs and LSECs. However, this method subjects the cells to considerable shear stress, which is probably why cell cultivation and functional studies of LSECs and KCs purified by FACS have been so scantly described in the literature. It has been reported that FACS of murine liver cells to produce LSECs resulted in a higher purity compared to MACS but was associated with a lower yield and recovery rate [12]. See [13] for a systematic review on methods for isolation and purification of murine LSECs. What is clear is that the cell yield, viability, and purity are affected by several steps in the cell isolation procedure, including the enzyme and technique used to disperse the liver cells, buffers, centrifugation steps, and antibody used for sorting. In rat, highly pure LSECs are reported with MACS isolation [14, 15] using the LSEC-specific antibody SE-1 [16], which targets the FcγRIIb2 in these cells [17].

We here describe the MACS procedure that we have developed for isolation of mouse KCs and LSECs, using F4/80 and CD146 as cell markers, which allows simultaneous isolation of highly pure HCs, LSECs, and KCs. The method is reliable because specific antibodies are commercially available for recognition of either KCs or LSECs in a suspension of mixed liver cells. Moreover, the procedure is fast and reproducible, and gives good cell yields. In addition, the MACS isolation procedure requires equipment that is considerably less expensive and easier to operate than elutriation centrifuges or flow cytometry cell sorters. During the isolation procedure, the cells and buffers are kept cold to minimize intracellular metabolic processes. After the final isolation step the cells can be seeded in culture dishes and maintained for functional studies or pelleted and used for further analysis. This allows for both in vivo liver cell distribution of intravascularly administered compounds, as well as in vitro analysis of how compounds interact with isolated cultures of KCs, LSECs, and HCs.

#### 2 Materials


frozen until perfusion starts.

preparation). 5. Liberase™ Research Grade, rehydrated and aliquoted in 1 mg batches according to the manufacturer's protocol, and kept

Fig. 1 Equipment for the perfusion system


2.3 Buffers, Density Medium, Liberase™, Cell Culture Media, Coating of Cell Culture Dishes with Fibronectin, Magnetic-Activated Cell Sorting (MACS) Isolation System (Hardware and Buffers)

9. Prepare MACS buffer: Mix autoMACS Rinsing Solution (Miltenyi) and MACS BSA Stock Solution (Miltenyi) according to instructions by the manufacturer.

#### 3 Methods


Animals used for Experimental and Other Scientific Purposes.

Fig. 2 Position of the mouse organs

Fig. 3 Position of the portal vein and inferior vena cava


Fig. 4 Position of the canula in the portal vein

Fig. 5 Color shift of the organs

Fig. 6 Removal of the liver

Fig. 7 Removal of the gall bladder


#### 3.3 Cell Isolation 1. Very gently mix the cell suspension.


3.3.1 MACS Isolation of

KCs and LSECs

	- 2. Fill each of the two 5 ml tubes with MACS buffer and centrifuge (300 g for 10 min at 4 C; max acceleration/deceleration). Aspirate both supernatants completely.
	- 3. Resuspend each of the two pellets in 0.5 ml MACS buffer and apply to two LS columns for magnetic isolation according to the instructions given by the manufacturer.
	- 4. Wash the column with 3 - 3 ml MACS buffer. Discard the flow-through.
	- 5. Remove the column from the magnetic field and fill it with 5 ml MACS buffer. Immediately flush out the magnetically labeled cells by firmly pushing the plunger into the column. Collect the eluted volumes in two 15 ml centrifuge tubes—one tube for the cells incubated with anti-F4/80 MicroBeads, and the other for the cells incubated with anti-CD146 MicroBeads. Spin the tubes (300 g for 10 min at 4 C; max acceleration/deceleration), resuspend the resulting pellets in 1 ml AIM V medium, and count the cells. See Note 8 for details regarding yield, purity, and viability of isolated KCs and LSECs.
	- 6. At this point the KCs and LSECs may be analyzed for contents of drug administered prior to the liver perfusion procedure.
	- 7. If cells are to be used for in vitro analysis of drug uptake and intracellular processing, they may be cultured according to the following procedure.
	- 1. Seed 0.25 - 10<sup>6</sup> LSECs/200 μl AIM V medium/cm2 growth area in fibronectin-coated cell culture wells. Place in a humidified CO2 incubator at 5% O2.
	- 2. After 35 min carefully wash with PBS or medium.
	- 3. The LSEC cultures are now ready for experiments. Characteristic morphology of cultured LSECs is shown in Fig. 8a, b. See Note 9 for details regarding purity of cultured LSECs.

3.3.2 Culturing LSECs (Continue Here After Step 5 in Subheading 3.3.1)

Fig. 8 Characteristics of the LSEC cultures. Phase contrast (a, <sup>c</sup>, <sup>e</sup>) and scanning electron micrographs (b, <sup>d</sup>) of mouse liver cells following isolation and culture. Liver cells were isolated as outlined in Subheading 3.3 and established in monolayer cultures. (a, <sup>b</sup>) LSECs 2 h following seeding. The inset in b shows typical LSEC fenestrations (open trans-cytoplasmic pores), which are a hallmark of these cells. (c, <sup>d</sup>) KCs 1 h following seeding. (e) HCs 48 h following seeding. LSECs and HCs were seeded on fibronectin-coated tissue culture plastic, and KCs were seeded on non-coated culture plastic. KCs and LSECs were cultured in AIM V medium, in 5% O2 and 5% CO2 atmosphere, whereas HCs were cultured in William's E medium with supplements, 5% fetal bovine serum, 0.1 <sup>μ</sup>M dexamethasone, and 20% O2 and 5% CO2. Following the indicated cultivation times the cultures for scanning electron microscopy were gently washed with medium prewarmed to 37 C, fixed in McDowell's fixative for electron microscopy [18], and processed according to the protocol in [14], and examined in a Zeiss Sigma field emission scanning electron microscope, run at 2 kV. Phase contrast micrographs were taken with a 20 objective lens in a Nikon Eclipse TE300 inverted microscope, equipped with a Zeiss AxioCam MRc digital camera

3.3.3 Culturing KCs (Continue Here After Step 5 in Subheading 3.3.1)

3.3.4 Isolation and Culturing of HCs (Continue Here After Step 4 in Subheading 3.3)


#### 4 Notes


centrifugation step further will not give significantly increased yields of LSECs and KCs.


#### Acknowledgments

The authors wish to acknowledge Prof. Karen Kristine Sørensen at UiT The Arctic University of Norway for assisting in proof reading of the manuscript, and also for providing light and electron microscopic images shown in Fig. 8.

#### References


cells. J Histochem Cytochem 41(8): 1253–1257


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part VI

Intellectual Property

# Chapter 28

# Patent Considerations When Embarking on New Antisense Drug Programs

Laurence D. S. Gainey

#### Abstract

The aim of this chapter is to provide some key information on patents and the patent system to assist someone embarking on the design, development, and commercialization of new antisense drugs.

Here I outline certain key topics such as what is a patent? why patent? how do I protect my molecule with a patent? confidentiality, searching for the same or similar molecules in the prior art, data requirements, the patenting timeline and freedom to operate.

Keywords Patent, Intellectual property, Patentability, Prior art, Searching, Patent filing, Exclusivity, Freedom to operate, Confidentiality

#### 1 Introduction

When embarking on a research and development program for any therapeutic, including an antisense drug, it is crucial that appropriate consideration be given to patents and the patent system. A patent protecting a therapeutic is often the main tool to keep bioequivalent versions off the market for a limited period, thus allowing the innovator time to recoup and secure a return on their investment. It is also important to consider third-party patents in case these impede the ability to commercialize the therapeutic.

The aim of this chapter is to provide a basic understanding of the patent system for someone embarking on a research program to discover and develop a therapeutic antisense drug. However, as patent decisions for any particular project will be case specific, appropriate legal advice should be sought from a qualified patent attorney.

The chapter is split into sections that discuss:


The patenting of a putative therapeutic molecule is usually crucial to the commercial success of the therapeutic. It is therefore recommended that you engage the services of a patent professional with expertise in handling antisense oligonucleotide patent portfolios at the earliest opportunity and keep them appraised of developments in the research program throughout.

#### 2 What Is a Patent?

A patent is a property right given by a country's government to an inventor, or their successor, that protects a new and useful invention. It is a reward for publishing the invention and provides the recipient with an exclusive right, for a limited period of time, to prevent others from exploiting, such as making, using, importing, offering for sale or selling, the invention in the country. Once this period of exclusivity has expired the public are free to exploit the invention and this arrangement with the state—a limited period of exclusivity in exchange for publishing and teaching the world what the invention is, is often referred to as the "patent bargain" and is what helps to stimulate and drive innovation.

Some important features of a patent are:


authority. Also, exploitation of the invention may fall within the scope of someone else's patent and so authorization, such as via a license, from this entity may be needed to work the invention.


#### 3 Why Patent Inventions?


antisense oligonucleotide product or its use can also be very valuable to individual inventors, small companies, and academic institutions.

3.3 Collateral for Raising Funding Because of the opportunity to secure an exclusive market position as in Sect. 3.1, a strong patent portfolio is attractive to investors and so can assist with company flotations or raising further capital. 3.4 Incentive to Invest in Research and Development Without the existence of a patent protecting a therapeutic antisense oligonucleotide molecule or its use in treating a particular disease or patient group, there may be significant disincentives to invest in the research and development costs and lost commercial opportunities.

#### 4 Types of Inventions to Consider

A patent protecting the oligonucleotide drug molecule being developed, often defined by its sequence, is likely to be particularly desirable as this will protect the molecule regardless how it is used.

However, in addition to a new oligonucleotide defined by its sequence and/or particular chemical modifications (e.g., to the backbone, sugars, bases) throughout or at specific positions, it is important to recognize that valuable patenting opportunities might also be found for new types of chemical modifications, fusions or conjugates, manufacturing processes, formulations, drug delivery technologies, particular diseases to be treated (therapeutic uses), particular patient populations to target, dosage forms of the drug, and/or dosage treatment regimens, to name just some categories of invention. It is therefore important to think beyond the specific molecule/compound to see if there are other patentable opportunities that can be captured to enhance the patent portfolio protecting your entire research program.

#### 5 Patenting Requirements

Each country (jurisdiction) has its own legal requirements (legislation) of what constitutes a patentable invention, but typically the claims that define the invention must meet certain criteria, including (1) novelty (new); (2) inventive step (not obvious) over what is already known; and (3) industrial utility (be useful). In addition, the application must include an adequate description of how to carry out the invention.

Individual country legislation may also preclude the patenting of certain types of subject matter in that country. For example, certain countries do not permit patents directed to methods of treatment, surgery or diagnosis practised on the human body. However, antisense oligonucleotide molecules, particularly those with chemical modifications, constitute patentable subject matter all the key countries (including US, Europe, Japan, China).

#### 5.1 Novelty The claimed invention must not have been disclosed in the public domain prior to the filing date of the invention.

According to Article 54(1) of the European Patent Convention [2]:

An invention shall be considered to be new if it does not form part of the state of the art.

Thus, a claimed invention is novel/new if it does not form part of "the state of the art," the state of the art being everything that was available to the public by way of written (including via electronic means) or oral publication, use or any other way, in any country of the world and in any language before the effective filing date of the invention. The effective filing date of the invention is the date of filing of the first patent application (e.g., priority application or International application—see 7(b)) that appropriately discloses the claimed invention. Novelty is an objective measure. Each and every feature of the claimed invention must have been disclosed together in the prior art to preclude patenting based on novelty.

An ideal antisense molecule may be the culmination of optimum choice of a number of features including, target sequence, length, complementarity, and chemistry. Thus, even with recognized therapeutic targets, there is often scope for establishing novelty over the prior art molecules. What will likely determine the ability to secure a patent on the molecule will be inventive step or non-obviousness.

#### 5.2 Inventive Step (or Non-Obviousness) It is somewhat subjective, and the assessment of inventive step differs from one country to another but, in essence, it is an assessment of whether the difference between the claimed invention and the state of the art, as assessed by the person skilled in the art, is obvious.

According to Article 56, first sentence, of the European Patent Convention [2]:

An invention shall be considered as involving an inventive step if, having regard to the state of the art, it is not obvious to a person skilled in the art.

Because this assessment is to be made by the notional "person skilled in the art," it is intrinsically more difficult to gauge whether a claimed invention is inventive and is the main discussion point between a patent examiner and the patent attorney seeking to secure grant of the patent.

However, indicators of inventive step accepted by most patent offices include surprising and/or unpredictable results. To demonstrate this, it will likely be necessary to have comparative data which demonstrates that the new antisense molecule, as claimed, possesses some superior property, perhaps much greater affinity or half-life, compared to that of the closest prior art molecule.

#### 5.3 Industrial Utility The claimed invention must be capable of being made or used in any type of industry.

According to Article 57 or the European Patent Convention [2]:

An invention shall be considered as susceptible of industrial application if it can be made or used in any kind of industry, including agriculture.

In general, this is an easy criterion to meet, and it is difficult to see how an antisense oligonucleotide invention would not meet this requirement.

Finally, the application must include sufficient information to allow the person skilled in the art to work the invention across the whole area claimed without undue burden or requiring inventive skill.

According to Article 83 or the European Patent Convention [2]:

The European patent application shall disclose the invention in a manner sufficiently clear and complete for it to be carried out by a person skilled in the art.

Similar or equivalent provisions to those above, illustrated using the European Patent Convention [2], are to be found in the patent legislation of most counties.

#### 6 Confidentiality

5.4 Sufficiency/ Written Description

As noted above, to be patentable an invention must be novel.

The invention must therefore have not been disclosed to anyone outside of a confidentiality agreement or put into the public domain by any means. Numerous patent applications have failed to meet the novelty or inventive step criteria due to an inventor's own disclosure/publication.

Aside from publication in a scientific journal, other publications to be aware of include, website postings, or poster or oral presentation at a scientific conference. If the researcher is from an academic institution, other potential publications include: an internal academic presentation attended by individuals from outside the institution or under no obligation of confidentiality, a PhD thesis, a discussion with an academic from a different institution, indeed any oral disclosure to someone not required in law to keep the information confidential.

While some countries, such as USA, provide for a "grace period" that allow prior publications from the inventor, made within a limited period before the filing date of their patent application, to be discounted from the prior art when assessing patentability, thus allowing a patent application to be filed after an earlier publication, it is a mistake to rely on this facility because "grace period" provisions only exist in a few countries. As noted above, therapeutic drug development costs are very high. In order to recoup such investment, it is typically desirable or necessary to be able to sell the drug globally, and thus important that patent protection for the drug be secured in the major markets. Many of the major pharmaceutical market countries do not have a grace period provision. An ill-timed publication could therefore undermine the opportunity to secure a patent in certain of the major markets which will have serious commercial implications. For example, it may dissuade a potential licensee or could be a reason for ceasing development of the molecule.

It is therefore crucial that the invention is kept confidential, at least until the patent application has been filed (see Note 1).

#### 7 Patent Timeline

There are various routes available for seeking a patent in a particular country.

However, in situations where the applicant desires patent protection in multiple countries the following route is commonly adopted:


A typical timeline that uses the PCT system outlined above is shown in Fig. 1.

More details on the PCT system can be found on the PCT website [3].

#### 8 Data Requirements

Patent attorneys are often asked how much and what type of data must be included in the patent application. There is no hard and fast rule on this. Generally, it is case specific and will depend on various factors such as the complexity in the art.

The majority of patent systems typically do not mandate that the patent application contain any number of specific examples, nor that these examples be actual worked examples with real data. However, in order to be patentable, the invention must be sufficiently described and enabled (or supported) to permit a person of

Fig. 1 Basic timeline for Patent Cooperation Treaty (PCT) patent application filling

skill in the art to practice the invention. With predictable technologies, such as mechanical devices, disclosure of a single means for making or using the invention may suffice. The person skilled in the art can easily make the invention and understand whether and how it works. In more unpredictable fields, such as therapeutic agents acting within biological systems, it is likely that the amount and type of data needed to demonstrate that the invention works and that the person skilled in the art can practice the invention will be greater. Thus, in practice, with therapeutic-type inventions, the presence of worked examples with relevant data plays a significant part in satisfying the patent examiner (and/or court) that the invention as claimed is supported/enabled and works.

Antisense oligonucleotide inventions, as with other therapeutic molecules, particularly those in established fields would benefit from two categories of data.

The first are data that would convince the person skilled in the art that the molecule has the necessary biological effect to make it credible that it would work as claimed (supporting data). Thus, if a disease pathology is manifest by expression of a particular protein, data demonstrating that the antisense oligonucleotide can, for example, impede production of the protein would be advantageous.

It is not necessary that the application include actual clinical trial data demonstrating efficacy in the relevant subjects (e.g., human patients). Suitable in vivo or ex vivo animal data or even cell data from an in vitro system, demonstrating that the antisense molecule impedes protein production, for example, may be sufficient and should be included in the patent application.

The second are data that demonstrate that the claimed invention performs better than what is already known (comparative data). Comparative data is particularly helpful, indeed often necessary, if the claimed invention is close to and thus potentially prima facie obvious over what is already known (in the public domain). Data comparing the claimed invention with the closest prior art molecule can help support the position that the claimed invention meets the inventive step (non-obviousness) criterion.

While it may be possible to supply this comparative data during the patent examination/prosecution stage it is advantageous to have this in the application as filed. However, in order to be able to do this, the researcher must have identified which molecule is likely to be determined to be the "closest prior art molecule" and then to have synthesized and tested this molecule against those of the claimed invention.

#### 9 Searching

As will be appreciated from the above, the claimed invention is assessed against what is already known (in the public domain).

When embarking on a research program for a new antisense oligonucleotide, it is recommended that the researcher conduct or commission searches of the published literature, including published patent applications, to identify, for example, the various molecules against the target sequence that are in the public domain. This information can assist on two fronts. First, it offers the opportunity to conduct research in novel space and design novel molecules; secondly, it should identify the molecule(s) that are likely to be considered the closest prior art molecules against which the new molecule with be assessed for patentability.

Prior art and patent searching is a skill that requires access to and an understanding of how to search particular databases of published documents. The ways of conducting searches of the prior art is beyond the scope of this chapter. The researcher may have the tools (e.g., databases of publications) and skills to conduct their own prior art searching. Otherwise, it is recommended that this be commissioned from an appropriate search firm.

#### 10 Freedom to Operate (FTO)

The final topic to flag is that of freedom to operate. As noted above, a patent on your product (e.g., antisense oligonucleotide drug) is an exclusionary right. It does not give the right to commercialize the product, and one obstacle to exploitation may be the existence of one or more third-party patents that dominate the ability to make, use or sell the product. Examples of dominating third-party patents might be those that cover a broad class of molecules that your development candidate falls within or claims to the precise therapeutic use (e.g., disease to be treated) that a molecule such as the one you are developing is to be used for.

It is therefore also important to consider third-party patents in case these impede the ability to commercialize the product. An FTO assessment for third party patents starts with searching for the existence of relevant pending applications or granted patents. It is highly recommended that such searching be conducted by professional searchers and the search output should then be assessed by a suitably qualified patent attorney. Once potentially relevant thirdparty patent property, both pending applications and granted patents have been identified, strategies to mitigate the risk of patent infringement can be devised. This could include securing legal opinions on infringement and validity, initiating challenges to the validity of such patents and/or securing appropriate patent licenses to any dominating patents.

The most meaningful FTO assessment can only be carried out when the actual product and its method of manufacture have been identified. However, if FTO assessments are carried out at an early stage it may help to gauge risk and potentially allow the design of molecules that avoid problematic third-party patents. For example, if equivalent chemical modifications are feasible, only one of which is protected by third-party patents, it may be possible to utilize the patent-free modification to avoid the FTO risk.

#### 11 Summary

When embarking on a research program to develop a therapeutic antisense oligonucleotide, it is important to consider and incorporate a patent strategy that includes seeking protection for patents on the product and being aware of third-party patents that might be an impediment to commercialization.

It is advantageous to conduct prior art searches early on to ensure that your molecule is novel over those already known and to identify the closest prior art molecules against which your product will be assessed for patentability, in particular inventive step. It is imperative that the details of the invention are kept confidential until after the patent application has been filed, at least.

It is important to generate data that demonstrates that the molecule has the biological properties likely to be of use in the purported treatment and to include this data in the patent application. In vitro cell data may be sufficient.

It may also be advantageous to generate comparative data to demonstrate that the claimed molecule has some unexpected beneficial property or improvement over the closest prior art compound. If this data is included in the patent application, it could help to satisfy the patent examiner that the claimed molecule meets the inventive step criterion and thus greatly facilitate the patent prosecution. If it is not available at the time of filing it may be useful, if not actually necessary, to generate this data for use in the patent prosecution stage.

It is important to recognize that valuable patenting opportunities may exist for subject matter other than the antisense compound itself and its particular use and you should consider whether seeking patent protection for these would strengthening the overall patent portfolio.

Finally, you should engage the services of a patent professional with expertise in handling antisense oligonucleotide patenting at the earliest opportunity. They can provide counsel on the appropriateness of the data generated and the strategy for filing and prosecuting the patent application(s), including which territories to pursue for patent protection. They can draft and file the patent application(s) and manage the patent portfolio. They can also assist with any necessary prior art or third-party patent searching and advise on the best time to undertake the necessary freedom to operate assessments.

#### 12 Glossary

Applicant—the individual or legal entity that has filed for and has claimed initial ownership of a patent application.

Freedom to operate (FTO)—is the ability to develop, make, sell, offer for sale (i.e., market) the product without legal liabilities to third parties, including third party patent owners.

Licensor—the person or entity that grants formal permission to make or do something. In the context of a patent the right to perform certain acts protected by the patent.

Licensee—the individual who obtains the license from the licensor to make or do something.

Patent claims—a series of independent or linked (dependent) sentences that define the scope of what is being claimed as the invention. The patent claims form one section of the patent application.

Patent Cooperation Treaty (PCT)—is an international patent law treaty which provides a unified procedure for filing patent applications in each of its contracting states. It is administered under the auspices of the World Intellectual Property Organization (WIPO). A patent application filed under the PCT is typically referred to as an international application, or PCT application. The PCT application is merely a vehicle to facilitate grant of individual national patents. It is not, and cannot, become a patent itself. There is no such thing as an international patent.

Patent office—is a governmental or intergovernmental organization which processes and awards patents.

Patent portfolio—refers to the collection of patent applications and granted patents owned by an applicant.

Patent prosecution—is the process taken to establish whether the patent application meets the criteria for grant of a patent. It may also be referred to as patent examination.

Patent term (or "term")—is the length or period of time that a patent can remain in force. Once this term has finished the patent expires and the invention enters the public domain, so anyone is entitled to freely exploit the invention.

Patent term extension—is an extension of time to the usual patent term.

Person skilled in the art—is a fictitious individual used in many patent laws of the world as the individual that is used as reference to determine whether the claimed invention is patentable (e.g., inventive) and the application sufficiently disclosed, among other things. Often the person skilled in the art is a technician skilled in the technical area of the invention who is aware of the prior art and has common general knowledge but no scintilla of ingenuity or inventiveness. Sometimes the person skilled in the art can be a team of individuals. It is a legal fiction.

Prior art—refers to all information that has been made available to the public prior to the effective filing date of the claimed invention. The effective filing date being the date of filing of the application that first discloses the claimed invention. This could be, for example, the date of one of the priority applications or the substantive application.

Priority application or priority patent application—a patent application filed in a national or regional state (including PCT application) that can be used to claim priority to from a subsequently filed national, regional or international application.

Public domain—refers to information or inventions which are freely available to the public, in particular, where no exclusive intellectual property rights exist or are possible. It can refer to patented inventions whose term has expired or information which has been published and so no longer protectable by a patent.

State of the art—refers to information which is known, or is capable of being known, by the public. It is often a synonym for prior art (information known before the patent filing date).

Substantive application—is the national or international application which will be subjected to patent examination and whose text can no longer be changed. Such filing may claim priority from an earlier "priority" application.

Supplementary Protection Certificate (SPC)—is a distinct intellectual property right available in countries of the European Economic Area that extends the term of certain rights under a patent for pharmaceutical or agrochemical products that have had to undergo regulatory assessment. It enters into force immediately the patent on which it is based expires. The SPC is designed to compensate the applicant for delays incurred in securing regulatory approval for a product, and it effectively extends the monopoly right for the product for a limited period of time.

#### 13 Notes


#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# INDEX

#### A


#### B


#### C



#### D


Virginia Arechavala-Gomeza and Alejandro Garanto (eds.), Antisense RNA Design, Delivery, and Analysis, Methods in Molecular Biology, vol. 2434, https://doi.org/10.1007/978-1-0716-2010-6, © The Editor(s) (if applicable) and The Author(s) 2022

#### 420 ANTISENSE RNA DESIGN, DELIVERY, AND ANALYSIS Index


#### E


#### F


#### G


#### H


#### I


#### K


#### L


#### M


#### N


#### O

Off-target.................................................. 8, 9, 17, 70, 83, 98, 183, 297, 357–359, 363, 372

#### ANTISENSE RNA DESIGN, DELIVERY, AND ANALYSIS Index 421


#### P


#### Q


#### R


#### 422 ANTISENSE RNA DESIGN, DELIVERY, AND ANALYSIS Index


#### S


#### T


#### U


#### V

Variants ............................................ 35, 36, 90, 145, 146, 148, 150, 151, 157, 167–169, 208, 235, 241, 246, 251, 270, 281, 289, 315, 322

#### W


#### Z

Zebrafish......................................... 74, 84, 276, 281–298